UNIVERSITY OF • T JWNOIS LIBRARY AT URBANA-CHAMPAIGN AGRICULTURE NOTICE: Return or renew all Library Materials! The Minimum Fee tor each Lost Book Is $50.00. The person charging this material is responsible for its return to the library from which it was withdrawn on or before the Latest Date stamped below. Theft, mutilation, and underlining ot books are reasons for discipli¬ nary action and may result in dismissal from the University. To renew call Telephone Center, 333-8400 Digitized by the Internet Archive in 2018 with funding from University of Illinois Urbana-Champaign Alternates https://archive.org/details/1990agricultural6886unit HERTZBERG — NEW METHOD, INC. EAST VANDALIA ROAD, JACKSONVILLE, ILL. 62650 TITLE NO. U-0520 ♦ OO -'HO 42-9 U f ' CULTURE ACCOUNT NO. 07200-222 LOT AND TICKET NO. CM 06- 39 M AGRICULTURE *■ DEPARTMENT ■* 43-15 AGf "iBOOK * i|| . / I I/. 43-24 1991*N0.&ati -o91* 42-28 Q* 630wli n 3 a gh** no♦688-691*COP♦2* CLOTH COLOR 0075 CHARGING INFORMATION SPECIAL WORK AND PREP. 02AGX4 HEIGHT 10 3/8 STUBBING FRONT COVER HAND ADHESIVE MAP POCKET PAPER HAND SEW NO TRIM LENGTHWISE MAP POCKET CLOTH THRU SEW PAGES LAMINATED FOREIGN TITLE SPECIAL WORK THRU SEW ON TAPE EXTRA THICKNESS LINES OF LETTERING REMOVE TATTLE TAPE T HEIGHT PICA WRAP u 6 I 1 0 - i ; - Cr ‘ A '££- 2 - 3 °) tand Rangeland Birds of the United States Natural History and Habitat Use (J30 __ U.S Department of Agriculture Forest Service Agriculture Handbook 688 YRjtf# k > garjuv ^;..' 4 :: v»ui UNIVERSITY Of ILLINOIS AGRICULTURE LIBRARY Forest and Rangeland Birds of the United States Natural History and Habitat Use Richard M. DeGraaf Northeastern Forest Experiment Station USDA Forest Service Amherst, Massachusetts Virgil E. Scott Denver Wildlife Research Center USDI Fish and Wildlife Service Fort Collins, Colorado OQ r—i o £ ^02 o pQ P5< cr> CD CO % R.H. Flamre Rocky Mountain Forest and Range Experiment Station USDA Forest Service Fort Collins, Colorado 02 O H ►—t m O o* w V* 05 Q W Liz Ernst Denver Wildlife Research Center USDI Fish and Wildlife Service Fort Collins, Colorado Stanley H. Anderson Wyoming Cooperative Wildlife Research Unit University of Wyoming Laramie, Wyoming AGRICULTURE LIBRARY JUN 1 1 1991 Agriculture Handbook 688 UNIVERSITY OF ILLINOIS January 1991 Cover photo: Cedar Waxwings in nest. © R Reinhold Using This Book This book is designed to provide managers with information on the assemblage of bird species that might be expected in forest and rangeland habitats. In the first section, natural histories are provided for each bird species, including taxonomic information, range and status, habitat descriptions, nest site descriptions, and food habits. These natural histories thus provide a short review of the pertinent information on each species necessary for initial evaluation of management practices for a particular area. The illustrations in the publication are reproduced from BIRDS OF NORTH AMERICA by Robbins et al., illustrated by Arthur Singer, © 1983, 1966 Western Publishing Company, Inc., and are used by permission. Nomenclature, distribution, and taxonomic order follow the American Ornithologists’ Union (1983). After the natural history information, a series of matrices list bird species that either breed or winter in 20 forest cover types and 24 rangeland, desert, and other nonforest habitats. The authors believe this guide can provide a basic source of data to assist land managers in ecologically sound management of resources. CONTENTS Introduction . 1 Distribution of birds in the United States. 1 Importance of vegetation structure. 2 Forest and rangeland management. 4 Deciduous forests . 4 Coniferous forests . 6 Rangeland habitats . 8 Acknowledgments . 10 Natural History, by Species . 13 Bird/Cover Type Matrices.531 Eastern forest types .540 Eastern non forest habitats .546 Great Plains habitats .553 Western forest types.562 Southwestern and western nonforest habitats .567 Literature Cited.576 Bird Species Index.612 INTRODUCTION As North America was settled, many once-abundant birds began to disappear. Some were hunted, but habitat loss or alteration was respon¬ sible for most losses. Around the turn of the twentieth century, conser¬ vationists began efforts to preserve habitats so that birds, especially plumed wading birds, could survive. Legislation was initiated to assist in the conservation of birds and to manage hunted species. Today, many public and private conservation agencies are involved in managing bird populations and their habitats. Birds are important components of ecosystems. Birds disseminate seeds and prey upon innumerable insect and vertebrate pests. They are involved in energy transfer as they eat and are eaten. Nutrients are also distributed through the movement of birds. Vultures, crows, ravens, and other scavenging birds are important in natural decomposition cycles. Because birds are not isolated components of our natural systems but integral parts of them, it is vital to understand both their roles and their needs. Management of birds—whether for human enjoyment, consumptive use, or ecological considerations—requires data on their biology and habitat use, as well as an understanding of community interactions. Wildlife biologists, foresters, rangeland managers, and land-use planners use comprehensive information on the total community—including birds—m habitat management. Information must be available on the species present in the area, their habitat requirements, and how birds will respond to habitat alterations. The objective of this book is to summarize information on the natural histories and habitat needs of forest and rangeland birds to help managers evaluate the impacts of various actions in different vegetation types. Opportunities can be identified to emphasize birds in management actions and to minimize negative impacts. These data, however, cannot replace the need for on-the-ground evaluations when projects are planned that will alter avian habitats; the natural history and matrix information compiled here must be applied along with local field knowledge. Distribution of Birds in the United States There are several broad patterns in the ecological distribution of breeding birds in the United States. There is a generally increasing continuum of breeding bird species from the drier areas of the South¬ west and West to the more moist forests of the Northeast (Peterson 1975). Regions dominated by desert vegetation have relatively simple avifaunas that contain a few prominent species. Regions with complex mixed forests contain many rather evenly distributed species, although 1 much regional variation occurs along this gradient. In the Great Plains, the number of species increases from Texas to the Canadian border (Cook 1969, Peterson 1975). This latitudinal increase may reflect patterns of glaciation or a more heterogeneous landscape on the northern plains, but in general the avian community in grasslands is organized by the most obvious structural feature, grass height (Cody 1968). Mountainous regions of the United States have relatively diverse avifaunas, largely because their considerable topographic relief com¬ presses several different vegetation zones into a relatively small area, rendering them ecologically diverse (Cook 1969). Closely related habi¬ tats, or habitats with similar physical profiles or complexities, exhibit similar bird diversities (Peterson 1975). Importance of Vegetation Structure An avian community, as defined here, is an aggregate of species existing together in a definable ecological area that provides the species’ requirements. Each species can exist only where its specific require¬ ments are met. Within the general habitat provisions of food, water, and shelter, birds have various specific needs for nest sites, song posts, perch sites, and vegetation structure. Some species have relatively narrow ranges of tolerance for specific factors. A prime example is the Kirtland’s warbler, which breeds only in fire-regenerated stands of young jack pine in Michigan’s Lower Peninsula. Others, such as the American robin, which breeds throughout North America, have broad ranges of tolerance and so are widely distributed. Many components of the environment, including vegetation struc¬ ture, plant species composition, succession, and vegetation layering, affect the distribution of bird species. What is not so obvious is that there are two basic sets of factors, ultimate and proximate, that deter¬ mine whether a bird can reproduce in a given area. Many of the factors that actually determine reproductive success are not evident at the time the bird arrives or selects its breeding habitat. Keys to these ultimate factors, such as food availability for nestlings, are perceived in advance through proximate factors—aspects of the physical habitat, especially vegetation structure. Ever since Lack (1933) suggested that birds select breeding habitats by recognizing features they did not immediately require for survival, many studies have been conducted to identify the features or patterns of vegetation structure that bird species were “programmed' to seek. Beecher (1942) expressed a similar idea, and suggested that a bird did not “adapt” to a so-called new habitat but rather chose the habitat because of its programmed ability to recognize potentially satisfactory ultimate factors. MacArthur and MacArthur (1961) demonstrated that the vertical complexity of forest vegetation (the diversity of vegetation heights and 2 density of foliage at those heights) affects breeding bird diversity. The relationship of bird species diversity to foliage height diversity has been demonstrated in many forest habitat types (Karr 1968, Karr and Roth 1971, Willson 1974). Foliage height diversity may be an indicator of total foliage volume. The important consideration for managers, however, is that habitat alteration changes the number of bird species and their relative abundances, both of which affect diversity. Studies of habitat selection and resource partitioning by breeding birds include measurement of many descriptors of stand structure. These stand measurements — canopy height, layering and closure, tree diameter and species composition, understory height and volume, ground cover, e t c _are attempts to identify the proximate factors that birds select when settling on the breeding grounds. Horizontal diversity or patchiness (the distribution of successional stages, timber size classes, and openings) is also important to breeding bird composition. Roth (1976) demonstrated that the number of bird species increased faster than the degree of species overlap in a series of habitats from grasslands to forests, and that horizontal habitat patchiness was a better predictor of the numbers of bird species than was vertical habitat complexity. Both the vertical diversity or structure of forest stands and the distribution of stands of different size class or type are typically manipulated in forest management and can be altered as needed to manage the type and availability of bird habitat. The close relationship between habitat structure and bird species composition is useful for assessing the effects of forest management on breeding birds. For example, as stands of northern hardwoods (sugar maple, American beech, yellow birch) develop after clearcutting, each tree size-class—regeneration, seedlings/saplings, poles, and sawtimber—supports a different breeding bird species composition (DeGraaf 1987). In the Willamette Valley, birds respond to successional patterns as Oregon white oak is replaced by Douglas-fir and finally by true fir and we stern hemlock. Downy woodpeckers, black-capped chickadees, and white-breasted nuthatches breed in the oaks, while chestnut-backed chickadees, red-breasted nuthatches, and golden- crowned kinglets commonly breed in Douglas-fir stands (Anderson 1970). Long-term changes in bird populations occur in response to environ¬ mental change. As land uses change, or as succession proceeds, bird communities and populations will change. Most habitat management projects are, in effect, attempts to control succession: either setting it back to an earlier stage, arresting it, or allowing it to advance to a desired stage. This process can also be accompanied by short-term changes in which individuals adapt to changing conditions. Outbreaks of some insects, for example, might attract birds to a forest where they normally do not feed, such as the woodpeckers that congregate in areas of mountain pine beetle infestation. 3 Forest and Rangeland Management Most forest management activities are not directed primarily at wildlife, but proper planning can help maintain or enhance wildlife habitats while accomplishing other goals. In logging or thinning oper¬ ations, successional patterns are changed, creating habitat for different species of birds. Clearcutting reduces nesting, shelter, and feeding habitat for forest canopy birds, but the habitats of ground and shrub dwellers will be enhanced. Small rodents will increase in clearcuts and provide a food base for raptors. As the forest regenerates, woodland hawks and owls replace open-country hawks and falcons. Forest canopy birds will replace the ground dwellers. Even-aged forest management creates forest stands of uniform age or size classes. The forest will have a variety of openings and stands, each with a fairly homogeneous structure, especially in smaller size classes. Thus, over the entire forest, a variety of bird habitats exists. Uneven-aged forest management produces trees of markedly differ¬ ent ages or sizes in the same stand. Thus, each stand in the forest has a fairly diverse structure, but all stands are fairly similar. The result can be a more diverse bird community in each stand than in any one stand under even-aged management, but it is often less diverse than that in an entire forest under even-aged management, where many succession¬ al stages are represented. A northern hardwood forest under even-aged management, for example, supports approximately twice as many breeding bird species as does an extensive uneven-aged forest. The lack of distinct, early successional stages in the uneven-aged northern hardwood forest means that species associated with those habitats— willow flycatchers, cedar waxwings, eastern bluebirds, chestnut-sided and mourning warblers, among others—will not likely be present (DeGraaf 1987). Deciduous Forests A striking feature of the bird communities of deciduous forests is the high proportion—up to 75%—of migratory species. Thus the total bird population (the standing crop) is very high in the summer breeding season. Bird communities in coniferous forests are seasonally less vari¬ able. In the deciduous forest there is a general pattern of increasing bird density with plant succession (see Shugart et al. 1975 for examples). The general pattern of avian succession is generally acknowledged to be a manifestation of the habitat preferences and ecological requirements of forest birds. In northern hardwood types, as with other hardwood types, the composition of bird species varies with timber size class, the presence or absence of softwoods in the stand, stand area, the presence of cavity trees, openings, and other within-stand features. In New England, 4 even-aged sawlog stands have avifaunas very similar to those in uneven- aged stands because the diameter distributions in both are similar; there are essentially no differences in foliage profiles in stands more than 30 years old (Leak 1979, Aber 1979). Thus the breeding bird assemblages in stands beyond the pole stage are similar. On wetter sites, where red spruce comes into mature stands, the softwood component provides habitat for species—especially warblers—that are typically associated with coniferous types. Oak-pine and oak-hickory types, which together compose most of the inland forests of the United States, support 150 to 200 species of breeding birds. Many are associated with successional stages and wetland and open habitats within these broad types, but their avifaunas are nevertheless quite rich. In oak-hickory types, site quality determines the dominants in mature stands: red, white, and black oak on good sites, post oak and blackjack oak on poor sites, and white and post oak on intermediate sites. Succession in oak-pine and oak-hickory types is similar, except that pine seedlings come in after disturbance during the “brush” stage in oak-pine. Old-field successional stages are different from those after clear- cutting, especially in the oak-pine type. Old fields produce essentially pure pine stands that persist a long time before oak components appear. After clearcutting, however, all components of the oak-pine types are present throughout stand development (Evans 1978). Oak-hickory and oak-pine forests, largely due to these differences in their successional patterns, support somewhat different avifaunas. Many species associated with coniferous forests are found in the oak-pine type (see Evans 1978 for review). The larger difference between the two types is the greater value of oak-pine forests to wintering birds because the type is more southern than oak-hickory, and conifers provide additional cover. In the western United States, aspen stands provide especially rich bird habitats compared to coniferous types. Moist ground surface, high insect populations in the understory, edge effect, and nest hole avail¬ ability (depending upon woodpecker use and Fomes infection) have been identified as factors primarily responsible for the rich avifaunas in these stands (Winternitz 1976). In both the desert Southwest and the Great Plains, riparian wood¬ lands are essentially deciduous; cottonwoods are important in both regions, although Great Plains riparian woodlands contain more tree species as one moves eastward. These habitats contain bird communi¬ ties that are as much as seven times as rich as those in the surrounding Plains habitats in general (Tubbs 1980). In the Southwest, a variety of riparian habitats, each with a more or less unique assemblage of breeding bird species, is extremely important for wintering and migrant species also. Bird density is directly correlated with densities of cottonwoods in these wooded southwestern-riparian zones (Carothers and Johnson 1975). 5 Coniferous Forests Approximately 150 bird species are associated with coniferous forests across the United States; the greatest number of species is found in mature forests in the Northeast, and the lowest in young stands in the same region. In other regions of the United States, the number of species is both lower and fairly uniform (Wiens 1975). A general pattern in coniferous forest avifaunas is that they are characterized by a few abundant species. Approximately a quarter of the individuals are of a single dominant species, and one-third to one-half belong to the two most abundant species. The degree of dominance by a few species decreases with species richness. Several dominant species that occur throughout coniferous forests in the United States include: northern flicker, red-breasted nut¬ hatch, brown creeper, American robin, hermit thrush, golden-crowned kinglet, dark-eyed junco, and chipping sparrow. These species occur in all coniferous regions, and can be considered the group most represent¬ ative of this vegetation type across the United States (Wiens 1975). Analyses of the ecological structure of coniferous forest avifaunas— e.g., those of Baida (1969), MacArthur (1958), and Sturman (1968a, 1968b)—have revealed several patterns of habitat utilization. Foliage- gleaning species are the most abundant birds in all North American coniferous forests, while ground-gleaners, bark-gleaners, and aerial feeders are less abundant in decreasing order. Foliage feeders constitute the greatest proportion in mature north¬ eastern and southeastern coniferous forests. All ground-gleaners are most important in early successional stands, and bark-gleaners and aerial feeders are proportionately more numerous in western coniferous forests. Among foliage feeders, insectivores predominate in all forest types. Warblers constitute the major component of the breeding avifaunas of the northeastern and southeastern coniferous forests, but their relative densities are lower in the western forests. Both the number of warbler species and their densities are substantially lower in western forests than in eastern North America, and their paucity is not compensated by other foliage-gleaning insectivores. These differences likely reflect differences in prey availability between eastern and western North America (Wiens 1975). In the longleaf-slash pine forests of the Southeastern and Gulf Coastal Plains, bird species richness can be maintained by retaining dead trees, avoiding extensive monocultures, and controlling—rather than eradicating—understory vegetation (Wood and Niles 1978). Loblolly-shortleaf pine, a subclimax or developmental stage of oak- hickory, occurs on the Southeastern Coastal Plain and Piedmont—the old Cotton Belt. The red-cockaded woodpecker, the only endangered species closely associated with upland loblolly-shortleaf pine, occurs in clans of 2 to 10 birds in stands at least 80 years old and 35 to 160 acres in size (Meyers and Johnson 1978). 6 As the loblolly-shortleaf type develops, four stages each have distinct breeding bird assemblages: grassland, shrubland, pine forest, and hardwood forest (Johnston and Odum 1956). Maintaining stands in earlier stages by shorter rotation (about 35 years) is eliminating mature pine and hardwoods. Short rotations eliminate cavity trees, understory vegetation, fruits and mast, and deciduous trees. Many bird species winter in this type. Because species richness declines from early successional stands and begins to increase at about age 35, short rotations adversely affect both breeding and wintering birds (Dickson and Segelquist 1978, Noble and Hamilton 1976, Quay 1947). The ponderosa pine type, which has the widest distribution of any pine type in North America (Little 1971), occurs in extensive stands in northern California, eastern Oregon, and the intermountain region, and as scattered islands in the Southwest and Rocky Mountains. Because stands range from savannas to mixed pine-broadleaf transition forest to pure ponderosa pine and mixed conifer stands, the type has a wide array of bird species associated with it (Diem and Zeveloff 1980). Where ponderosa pine is an important commercial tree, maintenance of snags and cull trees is important for cavity-nesting birds; a higher proportion of the avifauna is composed of cavity-nesting species in western than in eastern forests. Where the ponderosa pine type grades into the pinyon-juniper type in the Southwest, the ecotone between the two types contains fewer bird species than does either community alone. In the pinyon-juniper type, the number of breeding species that nest in holes or forage on trunks and branches is directly related to the density of pinyon pines. In winter, bird species richness and density are strongly related to juniper berry production (Baida and Masters 1980). The Douglas-fir region west of the Cascade Range is intensively managed because timber values are high, especially in old stands. Under natural succession, grass and shrub stages are followed by Douglas-fir, which forms dense, even-aged stands that persist for centuries. Mortality eventually opens the stand, and true climax western hemlock and western redcedar invade and an understory is reestab¬ lished. After four to six centuries without disturbance, climax western hemlock replaces Douglas-fir (Franklin and Dyrness 1973). In intensively managed forests, fire is commonly used for slash re¬ moval and seed-bed preparation after harvest cutting. Genetically superior seeds or seedlings are planted, and herbicides, fertilizers, and pest control practices quickly produce even-aged stands of fast-growing Douglas-fir. These trees are harvested at 50 to 150 years of age. This managed succession—shortening the grass-forb and shrub stages and eliminating snags and old-growth forest—directly affects the avifaunal composition. The second (shrub) successional stage, in which approximately 40 per¬ cent of the bird species associated with the type nest, is abbreviated. Fire affects birds indirectly by modifying their habitats. Removal of woody vegetation creates clearings with low vegetation that favors some 7 birds. Along the border between Arizona and Mexico, for example, open- country birds such as American kestrel, roadrunner, curved-billed thrasher, harlequin quail, and chipping sparrow are most common on the Mexican side where fire control is less stringent. The species most common in Arizona are birds of brushland or dense forest, including the blue-gray gnatcatcher, black-throated gray warbler, Scott’s oriole, and rufous-sided towhee (Marshall 1963). Some species are attracted to new burns, including robins, bluebirds, several sparrows, flickers, several woodpeckers, mourning doves, and pine warblers. Prescribed fire has long been used to produce habitat for bobwhite quail in yellow pine in the Southeast (Stoddard 1963). Breeding-bird density and diversity are slightly higher in burned than in unburned chaparral (Laurence 1966). In lodgepole pine and spruce-fir forests in Yellowstone National Park, numbers of bird species increased for 25 years after fire, then begin to decrease (Taylor 1973). In the moist, temperate coniferous forests of the Olympic Mountains, large lightning fires are less common than in Wyoming, but a similar avian response occurs over time: more species are unique to the first 20 years after fire than to stages 100 to 300 years later. Habitat for ground- or shrub-foraging species is generally enhanced in the first few years after fire, while habitat for mature-forest birds is decreased. However, during the first 1 to 3 years after fire, the bird community in coniferous forests may be more similar to that in mature forests than to the ground/shrub community (Huff et al. 1984, Bock and Lynch 1970, Bock et al. 1978), with the greatest post-fire changes in the composition of bird species occurring after about 20 years in the western United States. Kirtland’s warbler depends completely on periodic fire to maintain a very specific nesting habitat in young stands of jack pine in Michigan’s Lower Peninsula. For detailed information on the effects of fire on bird habitat, see the compilations of Lotan and Brown (1984), Wood (1981), and Wright and Bailey (1982). Management of cavity trees has become standard silvicultural practice for bird habitat management, but continued availability of contiguous old growth for the northern spotted owl remains a real concern. A detailed review of Douglas-fir management and bird species composition is provided in Meslow and Wright (1975). Rangeland Habitats Rangeland avifaunas characteristically have few breeding bird spe¬ cies compared to forests. Approximately 40 species (excluding waterfowl, raptors, and galliforms) occur with moderate frequency across major rangelands of the United States, from the shrub steppes of the Great Basin to the shortgrass, tallgrass, and mixed-grass prairies of the Great Plains (Wiens 1973, 1974). A rather small group of species characterizes each rangeland type; sparrows are present in moderate to high numbers 8 in all rangeland habitats. There is substantial seasonal and annual variation in total bird density but not in the species composition of breeding birds. Rangeland avifaunas are often dominated by one or a few species, and high dominance is associated with low numbers of breeding species (Wiens and Dyer 1975). Grazing and grain production are the dominant uses of rangelands. Grazing affects breeding avifaunas in various ways depending upon the intensity and rangeland type. In general, however, where grazing regimes affect vegetative composition only slightly, effects on avifaunal compo¬ sition are slight. Where intensive grazing produces marked changes in vegetative composition, avifaunal composition changes markedly, usually toward that characteristic of more xeric habitats (Owens and Myres 1973, Wiens 1973). Fencing to control grazing intensity, timing, and location may create a mosaic of range conditions and therefore of bird communi¬ ties of differing structure and composition. Type conversion to remove woody vegetation and increase forage production—whether by herbicide, mechanical means, or fire—can be beneficial or detrimental to avian habitats depending upon extent, pattern, successional stages involved, and effects on special habitat needs of certain species. An example of a detrimental effect is the removal of Ashe juniper on the Edwards Plateau of Texas, which is required for nesting by the golden-cheeked warbler, a threatened species. A beneficial effect in fire-treated chaparral is increased species richness, especially where brush “islands” are retained (Bell and Studinski 1972). The history of western livestock grazing and big game populations and their habitats have been described in detail by Wagner (1978). Generally, bison, bighorn sheep, and cattle feed more heavily on grasses; mule deer and mountain goats on shrubs and trees; and pronghorn and domestic sheep generally feed on forbs. Elk and horse feeding habitats overlap those of all of these other herbivores. Foraging by rodents, rabbits, and hares can profoundly affect rangeland vege¬ tation, especially in desert rangelands (Norris 1950, Rice and Westoby 1978). The preponderance of evidence indicates that grazing is generally harmful to waterfowl habitat and nesting success (Brown and Johnston 1978, Weller et al. 1958). In the Southwest, heavy grazing has also caused serious declines in populations of lesser prairie-chickens (Brown 1978), greater prairie-chickens, Montezuma quail, California quail (Leopold 1977), and northern bobwhite (Phillips et al. 1964). Intensive grazing is considered the primary factor in the decline of the Columbian sharp-tailed grouse (Miller and Graul 1980). Attwater’s prairie-chicken uses grazed pastures more than ungrazed pastures because green herbaceous vegetation is made available by grazing (Kessler and Dodd 1978). In North Dakota, American bitterns, marsh hawks, and short-eared owls nest only in tall, dense ungrazed grasses and legumes (Duebbert and Lokemoen 1977). 9 The effects of grazing on avian habitats vary from place to place. In areas of higher precipitation, grazing may provide more habitat patch¬ iness and so be generally beneficial to birds. In areas of low precipita¬ tion, protection from grazing may be necessary for a species that benefitted from grazing in an area of higher precipitation. Effects of soil, slope, and exposure, as well as amount and seasonal distribution of precipitation may be more important than grazing in affecting the quality of bird habitat (Ryder 1980). Fire is an important tool in maintaining grasslands. Where fires are suppressed, grasslands may be replaced in successional stages by shrub-dominated communities, thereby providing habitat for different species of birds. For example, control of fire in the upper Midwest has reduced that habitat for sharp-tailed grouse. Prescribed fire maintains proper cover conditions for both prairie chickens and sharp-tailed grouse in tallgrass prairie and promotes the growth of preferred subclimax foods (Miller 1963). Creating isolated stands of trees and shrubs often enhances habitat for grassland birds. Thus riparian habitats and shelterbelts receive disproportionate use by birds. These habitats not only have their own bird communities, but are also used by grassland-nesting species as well. Cavity nesters also use riparian habitats extensively. Primary cavity users, those birds that excavate their own nesting and roost cavities, and secondary cavity nesters, which use cavities already present, are common in riparian habitats near grasslands. Fencerows also provide shelter and nest sites for grassland birds and add to the year-round diversity of bird communities. Little of the Northern Plains is forested, and shelterbelts have attracted bird species that not otherwise occur there. Mature shelterbelts resemble a late successional stage of the lowland hardwood forest in the north-central states. Perches for singing and hawking birds are available above the surrounding cropland or grassland. Foliage gleaners utilize the leaves of trees and shrubs. Raptors and hole-nesters are common in old shelterbelts. None of these microhabitats exist without woody vege¬ tation. Now, with the advent of center-pivot irrigation and large grain drills and harvesters, many shelterbelts are being removed. Acknowledgments We thank those who helped us in various ways. Contributors of information for the matrices of bird habitat use include G. N. Back, University of Nevada (Great Basin); L. Frederickson, University of Missouri-Columbia (wetlands of the central Mississippi drainage); A. J. Bjugstad, USDA Forest Service (Northern Plains); D. Patton and R. Szaro, USDA Forest Service (Southwestern Deserts); T. Keeney and W. Laudenslayer, USDA Forest Service (California); P. R. Canutt, USDA Forest Service (Pacific Northwest); W. Burbridge, USDA Forest Service 10 (Intermountain West); W. P. Zeedyk, USDA Forest Service (Southwest); J. C. Capp, USDA Forest Service (Rocky Mountains); K. Sideritz, USDA Forest Service (Northern Rockies); P. B. Hamel, Tennessee Natural Heritage program (South and Southeast); C. Keller and K. Gutzwiller, University of Wyoming. C. S. Robbins and D. Bystrack, USDI Fish and Wildlife Service, provided Breeding Bird Survey data for the entire United States. J. J. Green and C. J. E. Welsh compiled information on bird species occurrence for the matrices. R. D. Nelson and R. D. Lindmark, USDA Forest Service, and C. H. Halvorson, U.S. Fish and Wildlife Service, reviewed the manuscript draft. J. Scott voluntarily typed the natural history section. 11 . 175 ? 180 ? 175 ?- 170 ? I Shemya I. |llnaiaska l > ,Buldir I Adak I ! Prince / Patrick Jo' Metv'" e 'Kodiak l )\South- - ampton I Queen Charlotte Vancou'' 6 ' North \ ^ Dakota I *> Sout t\l Dakota' Nebraska \ lot Missouri Arizona O : Puerto \ Rico fexico Guadalupe I Ranges described in text generally follow boundaries and names shown on this map, modified from the A.O.U. Checklist of North American Birds, 6th Edition, 1983. 12 Common Loon Gavia immer winter RANGE: Breeds from western and central Alaska and the northern Yukon east across Canada to southern Baffin Island and Newfoundland south to northern California (at least formerly), northwestern Montana, North Dakota, and northern Iowa east to southern New England and Nova Scotia. Winters principally along coast from Aleutian Islands south to California, from Newfoundland south to Gulf Coast, and on lakes and bays near coasts. During migration occurs on inland waters throughout most of the United States. STATUS: Locally common. HABITAT: Breeds on or near freshwater lakes, ponds, and occasionally riverbanks, from tundra south in either open or wooded habitat. Territories may range from entire lakes of more than 100 acres to bays of 15 to 20 acres. SPECIAL HABITAT REQUIREMENTS: Bodies of water deep enough to escape from enemies by diving and large enough that it can take flight (up to 1/4 mile). NEST: Nests on the ground as close to water as possible, on islands, in sheltered places in coves, or on promontories or headlands. May locate nests on bare soil, on floating or matted vegetation, on muskrat houses or on rocks. Same nest site may be used, presumably by same pair, year after year. FOOD: Captures food during dives underwater. Generally eats approximately 80 percent fish with some crustaceans, vegetable matter, and insects, varying with locale. REFERENCES: Palmer 1962, Smith 1981, Terres 1980, Vermeer 1973, Wahl in Farrand 1983a. 13 Pied-billed Grebe Podilymbus podiceps im. RANGE: Breeds in southeastern Alaska and from central Canada south locally through temperate North America. Winters through most of breeding range from southern British Columbia and the central United States southward, casually farther north. Also winters throughout West Indies, Central and South America. STATUS: Common; most widespread grebe in North America. HABITAT: Inhabits ponds with much shoreline and emergent vegetation, marshes with areas of open water 15 to 25 inches deep, and marshy inlets and bays. Found on ponds, sloughs, flooded areas, marshy parts of lakes and rivers, and occasionally estuarine waters with weak tidal influence. SPECIAL HABITAT REQUIREMENTS: Marshes, sluggish streams, ponds 18 acres or less, and some emergent vegetation. NEST: A solitary nester, with generally only one pair nesting per pothole. Constructs a floating nest usually in shallow water, but sometimes on water several feet deep and well concealed in emergent vegetation. Builds nest around or anchored to reeds, rushes, or bushes and usually within 50 feet of open water. FOOD: Captures food while swimming and during dives. Primarily eats fishes, but also insects and some crayfish. REFERENCES: Faaborg 1976, Glover 1953, Palmer 1962, Sealy 1978, Terres 1980, Wetmore 1924. 14 Horned Grebe Podiceps auritus winter summer RANGE: Breeds in North America from central Alaska and northern Yukon to northern Manitoba south to eastern Washington, central Wisconsin, and extreme western Ontario. Winters from the Aleutian Islands and southern Alaska along the Pacific Coast to southern California and along the Atlantic Coast from Nova Scotia to southern Florida. During winter primarily marine, off ocean beaches and rocky shores as well as in sheltered inlets and bays. May be found in any water as a migrant. STATUS: Common in North America except in the Southwest. HABITAT: Inhabits ponds, marshes, sloughs, backwaters of streams and rivers, shallow bays of large lakes, and flooded places with some open water.’ Can alight on and take wing from small streams. Early in breeding season, often found where relatively little plant cover extends above water. SPECIAL HABITAT REQUIREMENTS: Small, shallow potholes of 18 acres or less. NEST: A solitary nester, with one pair occupying an entire pond. Builds nests commonly in quiet shallow water, usually well within a fringe of emergent vegetation, although sometimes very exposed. The floating, partly submerged nest is anchored to reeds or bushes or to bottom. Prefers to nest in small ponds with open water where territory can be observed visually. FOOD: Usually feeds in water 5 to 25 feet deep. Food caught during long dives under water consists primarily of small fish, crayfish, amphipods, prawns, shrimp, aquatic and land insects, and some amphibians and leeches; may also include some vegetable matter. In winter, nearly half of diet may be crustaceans. REFERENCES: Faaborg 1976, Ferguson 1981, Palmer 1962, Robbins et al. 1983, Terres 1980, Wetmore 1924. 15 Red-necked Grebe Podiceps grisegena winter summer RANGE: Breeds from western and central Alaska to south-central Ontario south to St. Lawrence Island, the Alaska Peninsula, central Washington to south-central Minnesota; rarely to southwestern Oregon, northern Michigan, southern Quebec and New Hampshire. Winters along Pacific and Atlantic Coasts and casually along Gulf Coast. STATUS: Populations decreasing or stable. HABITAT: Inhabits quiet inland waters on prairies, in woodlands, and extending out onto tundra. Less commonly found on prairie sloughs and marshes, backwaters of rivers, and flooded areas. SPECIAL HABITAT REQUIREMENTS: Shallow lakes and ponds (rarely less than 10 acres per pair) with at least some emergent vegetation. NEST: Usually a solitary nester although sometimes nests in loose colonies. Constructs an anchored floating nest on water 2 to 3 feet deep within or near edge of emergent vegetation such as cattails, sedges, rushes, and sometimes bushes. Sometimes builds nest on a muskrat house. FOOD: Dives and feeds in water at or near the bottom. In marshes and lakes, consumes primarily aquatic insects and some fishes. Also takes land insects, crustaceans, mollusks, aquatic worms, amphibians, and some vegetative matter. REFERENCES: Chamberlin 1977, Palmer 1962, Tate and Tate 1982, Terres 1980. 16 Eared Grebe Podiceps nigricollis winter summer RANGE: Breeds from south-central British Columbia and Manitoba south to Baja California and south-central Texas. Winters inland from California, Nevada, Utah, New Mexico, and central Texas and on the Pacific Coast from southern British Columbia south to Guatemala. Casual in eastern United States and also in Old World. STATUS: Large breeding colonies and winter flocks common. HABITAT: Inhabits marshy lakes and ponds, and large pools in streams or rivers in the prairie region of North America. Less typically inhabits marshes with some open water. During winter, found on salt lakes, bays, estuaries, and seacoasts. SPECIAL HABITAT REQUIREMENTS: Freshwater lakes and ponds larger than 18 acres, with shallow margins and emergent vegetation. NEST: Nests in compact colonies or sometimes singly, in sheltered areas, or in shallow water away from emergent vegetation. Builds a floating nest that is a platform of marsh vegetation built up from the bottom or anchored to reeds. FOOD: Gleans land insects from surface of water, or captures aquatic insects during dives. Also takes small crustaceans, small fishes, leech eggs, mollusks, and amphibians REFERENCES: Faaborg 1976, Grinnell and Miller 1944, Low and Mansell 1983, Palmer 1962, Ratti in Farrand 1983a, Terres 1980, Verner and Boss 1980, Wetmore 1924. 17 Western Grebe Aechmophorus occ iri * nf »Us (includes Clark’s C Aechmophorus da L18 RANGE: Breeds from southeastern Alaska and south-central British Columbia to southwestern Manitoba south to southern California, New Mexico, northwestern Iowa, and western Minnesota. Winters along the Pacific Coast from southern British Columbia to Baja California, and from Utah, Colorado, New Mexico, and western and southern Texas south into Mexico. STATUS: Locally abundant. HABITAT: Inhabits fairly extensive areas of open water bordered by tall emergent plants. Found on marshes, lakes, and bays; in winter may be found on salt, brackish, and freshwater where small fishes are abundant. SPECIAL HABITAT REQUIREMENTS: Open, fresh lakes bordered by rushes or tules. NEST: Nests in colonies of hundreds, even thousands of pairs at some lakes, but occasionally nests singly. Builds nests in colonies closely spaced, with territories consisting of only immediate vicinity of nest. Constructs nests in extensive areas of open water bordered by tules or rushes, in emergent vegetation, or on dry land. Prefers to nest near or on open shallow water about 12 inches deep. Anchors nests to, or builds on, submerged roots of bulrushes or other plants. FOOD: Captures food during dives underwater. Consumes primarily fishes. Also takes mollusks, crabs, marine worms, salamanders, and insects. REFERENCES: Grinnell and Miller 1944, Lindvall and Low 1982, Palmer 1962, Terres 1980, Wetmore 1924. 18 American White Pelican Pelecanus erythrorhynchos RANGE: Breeds from south-central British Columbia to central Manitoba and southwestern Ontario south locally to extreme northern California east to northern Colorado, northeastern South Dakota, and southwestern Minnesota. Sporadic on central coast of Texas and from central to southern California. Winters along the Pacific Coast from central California and southern Arizona south to Central America, and from Florida and the Gulf States south; casually throughout breeding range in western North America. STATUS: Locally common throughout breeding range. HABITAT: Found primarily on lakes, also rivers, estuaries, and shallow coastal bays and inlets. Loafs on beaches, sandbars, and driftwood. SPECIAL HABITAT REQUIREMENTS: In freshwater habitat, islands isolated from mammalian predators. NEST: Nests on the ground in colonies of a few to several hundred pairs on small, relatively flat islands, without tall (over 3 feet) obstructions, with loose earth suitable for heaping into nest mounds. Rarely, nests on floating islands of marsh plants. Colonies prefer open areas of annual grasses and forbs, shrubs, and nonvegetated areas. FOOD: Primarily consumes fish caught by scooping pouch into water while swimming. Groups often cooperate in fishing. Adults require about four pounds of food daily, and feeding areas may be located far from nesting areas. Also feeds on some salamanders and crayfish. REFERENCES: Knopf 1979, Knopf and Kennedy 1981, Lingle and Sloan 1980, Palmer 1962, Terres 1980. 19 Double-crested Cormorant Phalacrocorax auritus RANGE: Breeds in the southeastern Bering Sea, southern Alaska, and from southwestern British Columbia and northern Alberta to Newfoundland south along Atlantic and Pacific Coasts; very locally throughout interior of North America. Winters along the Pacific Coast from the Aleutian Islands and southern Alaska south to Baja California and Guerrero; on the Atlantic Coast from New England south; in the Mississippi and Rio Grande Valleys; and along the Gulf Coast south to Central America. STATUS: Widespread and locally common. HABITAT: Inhabits coastal areas, bays, estuaries, marine islands, freshwater lakes, ponds, rivers, sloughs, and swamps. (Only cormorant likely to be seen inland around freshwater lakes and rivers.) Has a pronounced preference for perching in trees, on rocks, buoys, or other objects that overhang or project from the water. SPECIAL HABITAT REQUIREMENTS: Undisturbed nesting site and convenient, dependable food souce within a foraging radius of 5 to 10 miles from roost or colony. NEST: Nests in colonies of a few to 3,500 pairs on rocky islands, cliffs facing water, or in stands of live or dead trees in or near water. In the Northeast and along the Pacific Coast, nests on the ground, on rocky islands, or on cliffs. Inland and in Florida, usually nests in trees. Breeding colonies may be located from below sea level to over 5,600 feet. FOOD: Captures food during dives in water, generally 5 to 25 feet deep but sometimes up to 72 feet deep. Prefers to hunt in water with a sandy bottom rather than over a rocky or gravelly bottom. Consumes primarily saltwater fish of little commercial value, plus freshwater yellow perch, bullheads, sticklebacks, carp, crappies, and sunfish. Also eats some salamanders, crustaceans, reptiles, mollusks, and sea worms. REFERENCES: Palmer 1962, Stallcup in Farrand 1983a, Terres 1980. 20 Anhinga Anhinga anhinga RANGE: Breeds from central and eastern Texas east to coastal North Carolina and south to southern Brazil and Ecuador. Winters in southeastern United States from central South Carolina, Georgia, Florida, and the Gulf Coast southward. Occasionally disperses north of breeding range. STATUS: Common throughout range. HABITAT: Inhabits quiet or slow-moving, often rather murky waters. Usually found in wooded freshwater swamps, streams, or tree-fringed lakes with water lilies, lotus, and other aquatic vegetation. Found in cypress swamps, freshwater sloughs of sawgrass and reeds with scattered willow clumps, or mangrove-bordered salt and brackish bays, lagoons, and tidal streams. Primarily a freshwater bird, but will range to marine coasts. Often perches with wings partly extended to dry. SPECIAL HABITAT REQUIREMENTS: Quiet, sheltered waters with some trees for perching. NEST: Nests in small groups with herons and egrets. May appropriate nests of common and snowy egrets or little blue herons, or construct its own. Nests are usually 3 to 10 feet above water. FOOD: Catches food by diving under water from the surface, while flying over water, or from a perch. Eats primarily fish, but also takes aquatic insects, crayfish, shrimp, leeches, tadpoles, frog eggs, water snakes, young alligators, and small terrapins. REFERENCES: Oberholser 1974a, Palmer 1962, Pough 1951, Sykes in Farrand 1983a, Terres 1980. 21 American Bittern Botaurus lentiginosus RANGE: Breeds from extreme southeastern Alaska, central British Columbia, and southern Mackenzie to central Quebec and Newfoundland south to southern California, New Mexico, Texas, and Florida, breeds rarely south of northern California, Utah, Plains States, Ohio River Valley, and Virginia. Winters from southern British Columbia, Utah, New Mexico, central parts of Gulf States, and southern New England south to southern California, Gulf of Mexico, and along Atlantic Coast. STATUS: Rather common, but elusive. HABITAT: Inhabits freshwater or saltwater marshes, bogs, swamps, wet meadows, or wherever the ground is wet and tall, emergent vegetation such as cattails, bulrushes, and reeds are present. Usually perches on the ground, sometimes on a log or stump, or on cattails 3 to 4 feet above water, rarely in trees. Generally solitary; will freeze with neck and bill pointing upward, blending into marsh vegetation. SPECIAL HABITAT REQUIREMENTS: Wetlands with tall, emergent vegetation. NEST: Usually a solitary nester, but may form loose colonies in favorable habitat. Typically nests on flimsy platform of cattails, reeds, or sedges, 4 to 5 inches above water in emergent vegetation, occasionally on the ground among grasses or in shrubs. FOOD: Stalks food in marshes, meadows, along edges of shallow ponds, or wherever the ground is wet. Also searches for grasshoppers in dry meadows. Consumes mollusks, spiders, crustaceans, fish, frogs, salamanders, snakes, lizards, small birds, small mammals, eels, and land and aquatic insects. REFERENCES: Armistead in Farrand 1983a, Grinnell and Miller 1944, Low and Mansell 1983, Palmer 1962, Robbins et al. 1983, Weller 1961. 22 Least Bittern Ixobrychus exit is RANGE: Breeds locally from southern Oregon to central Baja California and southern coastal Sonora in the west; in the east across Canada from southern Manitoba, southern Ontario, and southern New Brunswick south to Texas, the Gulf Coast, Florida, and the Greater Antilles. Winters from southern California, southern Texas, and northern Florida south to Panama and Colombia. STATUS: Locally common, but elusive. HABITAT: Inhabits freshwater marshes, bogs, and swamps with dense cattails, reeds, bulrushes, buttonbush, sawgrass, smartweeds, arrowheads, and other tall aquatic and semi-aquatic vegetation. Prefers marshes with scattered bushes or other woody growth. Less commonly found in coastal brackish marshes and mangrove swamps. Usually is hidden in tall vegetation, and slips away by walking or climbing through reeds or even by running through them 2 to 3 feet above water, grasping a single reed or several in each foot. SPECIAL HABITAT REQUIREMENTS: Freshwater wetlands surrounded by tall aquatic vegetation. NEST: Nests singly in dense stands of emergent vegetation 6 to 24 inches above water that is 3 to 38 inches deep, and close to open water. Uses natural clump of the previous year’s vegetation to form the foundation of the nest. Occasionally nests in bushes, and more rarely, on the ground. FOOD: Feeds on the open water side of emergents, and captures small fish. Also takes frogs, tadpoles, salamanders, leeches, mollusks, crustaceans, insects, lizards, slugs, and occasionally small mammals. REFERENCES: Low and Mansell 1983, Palmer 1962, Terres 1980, Weller 1961. 23 Great Blue Heron Ardea herodias RANGE: Breeds from southern Alaska, coastal and southern British Columbia, southern Keewatin, and central Manitoba east to Nova Scotia and south, except in high mountains. Winters from southern-coastal Alaska, coastal British Columbia, central United States, and southern New England south to northern South America. L38" STATUS: Common throughout range. HABITAT: Inhabits a wide variety of freshwater and saltwater habitats including ponds, lakes, streams, rivers, marshes, wet meadows, tidal flats, sandbars, and shallow bays, or wherever shallow water or marsh vegetation is present. SPECIAL HABITAT REQUIREMENTS: Open water or wetland habitats. NEST: Generally nests in colonies, preferably in an isolated patch of woodland or on an island. Builds nests in the tops of the tallest trees, live or dead, often above 50 feet, but also in bushes, on rock ledges, sea cliffs, in tule rushes, and on the ground. In colonies, may build dozens of nests, which are used repeatedly, in the crown of the same tree. In mixed heronries, typically nests in highest parts of trees while other heron species occupy lower parts of same trees. May travel as far as 10 miles from nest sites to foraging areas. FOOD: Usually stands motionless in shallow water and waits until prey comes within striking distance. Also forages in wet meadows, pastures, dry fields, and even along road shoulders and in suburban ponds. Consumes small fishes, frogs, salamanders, lizards, snakes, shrimp, crabs, crayfish, aquatic and land insects, leeches, and small mammals. REFERENCES: Low and Mansell 1983, Palmer 1962, Pough 1951, Sykes in Farrand 1983a, Terres 1980, Verner and Boss 1980. 24 Great Egret Casmerodius albus (formerly Common Egret) RANGE: Breeds from southern Oregon and southern Idaho south to southwestern Arizona, and from southeastern Saskatchewan, southwestern Manitoba, southern Ontario, and Maine south through the Gulf States to South America. Disperses after breeding to the north into Washington, Michigan, southern Ontario and Quebec, and the Maritime Provinces. Winters from northern California across southern United States, and south along Atlantic Coast from New Jersey through South America. STATUS: Common throughout range. HABITAT: Inhabits streams, ponds, lakes, rice fields, freshwater and saltwater marshes and lagoons, and mud flats. After feeding during day, flies singly or in small groups to a communal roost in trees or shrubbery. Gregarious during all seasons. SPECIAL HABITAT REQUIREMENTS: Open water or wetland habitats near woodlands. NEST: Nests singly or in colonies, often with other herons, ibises, wood storks, cormorants, and anhingas. Usually nests in woods or thickets near water so long as there is adequate support for the nest. Builds nest from 1 to 40 feet above ground, depending on substrate. Sensitive to disturbance by people when nesting and may flush at the slightest provocation. FOOD: Forages in freshwater, brackish, or saltwater swamps, along streams, and in ponds. Consumes fishes, frogs, salamanders, snakes, snails, crustaceans, insects, and small mammals. REFERENCES: Grinnell and Miller 1944, Low and Mansell 1983, Palmer 1962, Sykes in Farrand 1983a, Terres 1980. 25 Snowy Egret Egretta thula L20” RANGE: Breeds from northern California and Montana south to central and eastern Texas, along the lower Mississippi Valley, and from Maine south along the Atlantic and Gulf Coasts to South America. After breeding, disperses north to Oregon, Nebraska, Great Lakes, and Atlantic Canada. Winters from northern California, southwestern Arizona, the Gulf Coast, and coastal South Carolina south throughout the breeding range. STATUS: Common; breeding range expanding northward. HABITAT: Inhabits ponds; borders of lakes; freshwater, brackish, and saltwater marshes and swamps; stream courses; tidal flats; rice fields; and sometimes dry fields, where it associates with cattle. NEST: Nests in colonies (many coastal), sometimes with thousands of pairs or in smaller colonies with other herons, ibises, cormorants, and anhingas, or even singly. In western United States, commonly nests on the ground in cattail marshes; in other areas, may nest up to 30 feet high in trees and shrubs. FOOD: Forages by rushing about and shuffling its feet in shallow water to flush its prey out of hiding. Consumes small fish, frogs, lizards, snakes, shrimp, fiddler crabs, crayfish, grasshoppers, cutworms, and aquatic insects. REFERENCES: Grinnell and Miller 1944, Low and Mansell 1983, Palmer 1962, Sykes in Farrand 1983a, Terres 1980. 26 Little Blue Heron Egretta caerulea im. i RANGE: Breeds from southeastern New Mexico to central Kansas, southern Arkansas, southeastern Missouri, southwestern Kentucky, northwestern Tennessee, central Alabama, southern Georgia, and along the Atlantic Coast from Maine, south to the West Indies and South America; sporadically in central Minnesota. After breeding, disperses north in interior to North and South Dakota, Michigan, southern Ontario and southern Quebec, and to Nova Scotia on Atlantic Coast. Winters from southern Baja California, the Gulf Coast, and coastal Virginia south throughout most of breeding range. STATUS: Common; range is expanding. HABITAT: Prefers freshwater ponds, lakes, marshes, meadows, and marshy shores of streams, but also inhabits brackish and saltwater coastal habitats. Roosts in trees and shrubs at night. SPECIAL HABITAT REQUIREMENTS: Open water or wetland habitats. NEST: Nests in colonies of up to 100 in a variety of trees, usually hardwoods, almost invariably over or by freshwater. Tends to nest on the fringe of mixed colonies, often in company with the tricolored heron. Builds a flimsy platform nest, sometimes as high as 40 feet in trees and shrubs. Nests in willows, buttonbush, red maples, myrtles, and swamp privet. FOOD: Forages in a slow, methodical manner ashore, in mud, or in very shallow water; not given to wading as deeply as some herons. Seldom feeds in saltwater. When water disappears from marshes and swamps, will live solely on insects caught in grasslands. Diet includes fishes, frogs, lizards, snakes, turtles, shrimp, fiddler crabs, crayfish, aquatic insects, spiders, grasshoppers, crickets, and beetles. REFERENCES: Low and Mansell 1983, Palmer 1962, Sykes in Farrand 1983a, Terres 1980. 27 Cattle Egret Bubulcus ibis im. V LI T RANGE: Breeds locally from northwestern and central California, southern Idaho, northern Utah, Colorado, North Dakota, southern Saskatchewan, Minnesota, Wisconsin, southern Ontario, northern Ohio, and Maine south to Florida, West Indies, and Central and South America. After breeding, disperses north to southern British Columbia, southcentral Canada, and the Maritime Provinces. Winters throughout most of breeding range. STATUS: Common; a recent immigrant from the Old World, range is rapidly expanding. HABITAT: Frequents a great variety of habitats including pastureland, freshwater and salt marshes, fallow and plowed fields, orchards, citrus groves, road shoulders and median strips, vacant lots, lawns, and other open grassy areas. Least shy and least aquatic of North American herons; usually found in close association with large hoofed mammals, particularly cattle, and often perching on their backs. SPECIAL HABITAT REQUIREMENTS: Wetlands for nesting. NEST: Nests colonially, often with other herons and ibises, in both freshwater and saltwater habitats, on islands, in willows and tamarisks along watercourses, occasionally in cypress swamps with a lower growth of buttonbush, or in scrub oaks in marshlands. Also nests in redcedar, red maple, and in pines. Usually builds nests at heights of 5 to 12 feet, up to 30 feet in heronries. FOOD: Usually feeds in dry or moist open pastures among livestock, capturing insects and other prey disturbed as cattle walk and graze. May glean ticks or bugs off cattle. Consumes grasshoppers, leopard and cricket frogs, spiders, and some toads. REFERENCES: Palmer 1962, Sykes in Farrand 1983a, Terres 1980. 28 Green-backed Heron Butorides striatus (formerly Green Heron) RANGE: Breeds from southwestern British Columbia to northern California, southern Nevada and Utah, and north-central New Mexico, and from the western edge of the Great Plains States, southern Ontario and southern New Brunswick south to eastern Panama. After breeding, disperses north to eastern Washington, Idaho, and southern Canada. Winters from western Washington, coastal and southeastern California, southern Arizona, Texas and Louisiana, and northern Florida and South Carolina south to South America. STATUS: Common, locally abundant. HABITAT: Found in a wide variety of freshwater and saltwater habitats, primarily those in riparian deciduous zones. These include wet woodlands, lakeshores, ponds, rivers, streams, swamps, and marshes. Commonly alights on trees, stumps, or submerged debris, but roosts on or close to the ground. SPECIAL HABITAT REQUIREMENTS: Wetlands or open water habitats. NEST: Generally a solitary nester, but sometimes nests in colonies of 6 pairs or more. Nest may be built away from water in dry woodlands and orchards, on a low tussock or muskrat house, or in trees near water, often a dense tangle in crowns of middle-aged trees, typically 10 to 15 feet above ground, but up to 30 feet. FOOD: Captures prey while standing and waiting in shallow water, or by walking slowly, typically in a crouched position. Consumes fish, frogs, crayfish, mollusks, prawns, insects, leeches, earthworms, small snakes, snails, and mice. REFERENCES: Low and Mansell 1983, Palmer 1962, Terres 1980, Verner and Boss 1980. 29 Black-crowned Night-Heron Nycticorax nycticorax RANGE: Breeds from central Washington and east-central Alberta to southern Quebec, northeastern New Brunswick and Nova Scotia south locally through the United States to South America. Wanders a great deal. After breeding, disperses over most of the United States not within its breeding range, except northern Rocky Mountain region, and north into Canada. Winters in the Southwest and the lower Ohio Valley, Gulf Coast, and southern New England south throughout the breeding range. STATUS: Common throughout most of its range. HABITAT: Inhabits a wide variety of freshwater, brackish, and saltwater habitats almost anywhere a wader might exist, including lakes, ponds, marshes, wooded swamps, slow streams with pools, or rivers. Roosts by day, usually in a well-foliaged tree, not necessarily near feeding grounds. SPECIAL HABITAT REQUIREMENTS: Open water or wetland habitats. NEST: Nests in small to large colonies, usually with other heron species, in almost any habitat: wooded areas near coastal marshes, spruce groves on marine islands, hardwood forests on offshore islands, swamps, cattail marshes on prairies, clumps of tall grass on dry ground, apple orchards, and sometimes in city parks. Nests close together on the ground to over 160 feet high in trees, and may be well concealed or in the open. FOOD: Forages mainly at night, by standing and waiting, or walking slowly along shallow margins of lakes, mud-bordered bays, and in marshy places where there is standing or slow-running water. Eats fish, frogs, tadpoles, salamanders, snakes, toads, crayfish, crabs, shrimp, squid, clams, mussels, dragonflies, algae, succulent plants, young birds, and small mammals. REFERENCES: Grinnel and Miller 1944, Low and Mansell 1983, Palmer 1962, Sykes in Farrand 1983a, Terres 1980, Verner and Boss 1980. 30 Yellow-crowned Night-Heron Nycticorax violaceus RANGE: Breeds from central and northeastern Texas to southern Nebraska, southeastern Minnesota, east to the lower Ohio Valley, and eastern Tennessee, southeastern Pennsylvania and Massachusetts south to South America. After breeding, disperses north to eastern Colorado, Iowa, southern Ontario, and Atlantic Canada. Winters from the Gulf Coast and coastal South Carolina south throughout breeding range. STATUS: Much less common than black-crowned night-heron. HABITAT: Inhabits both freshwater and saltwater habitats, usually lush river swamps, but also tidal flats, stagnant backwaters or bayous of large cypress swamps, mangrove swamps, or dry, rocky, almost waterless areas on certain islands. SPECIAL HABITAT REQUIREMENTS: Wooded swamps. NEST: Nests in small to large colonies, sometimes with black-crowned, little blue, tricolored, and great blue herons, or singly, in trees or bushes and sometimes on the ground. Often nests in willows close to water, in mangroves, or in baldcypresses, usually 15 to 20 feet above ground. FOOD: Hunts at night but also frequently by day. Unlike other herons, rarely takes fishes, but feeds largely on crustaceans, mainly crayfish, and land and fiddler crabs. Also eats mussels, frogs, aquatic insects, snails, small snakes, lizards, leeches, and terrapins. REFERENCES: Low and Mansell 1983, Palmer 1962, Sykes in Farrand 1983a, Terres 1980. 31 Glossy Ibis Plegadis falcinellus RANGE: Breeds locally from Maine and Rhode Island south to Florida, and west on the Gulf Coast to Louisiana. Also inland, at least casually, in Arkansas. Wanders, at least, casually, to the Midwest and southern Canada. Winters from northern Florida and the Gulf Coast of Louisiana south to South America. STATUS: Locally common; initially an irregular breeding bird in North America in small colonies along Atlantic Coast, but recently has increased in numbers. HABITAT: Found in freshwater, brackish, and saltwater habitats, primarily marshes and estuaries. Prefers shallow pools bordered by shrubs and emergent vegetation. SPECIAL HABITAT REQUIREMENTS: Wetlands. NEST: Nests in small colonies, usually with herons or other waders in a variety of habitats; in willows or mixed growths of mangroves, tropical buttonwood and salt myrtle in Florida; in willows, gum, swamp maple, bay and buttonbush in cypress swamps of South Carolina; on islands of tamarisk, waxmyrtle, and salt myrtle; in cordgrass; in mixed stands of holly, redcedar, bayberry, wild cherry, sumac, salt myrtle, Virginia creeper, wild grape, and cat greenbrier on barrier beaches along New Jersey Coast; and in cattail marshes. Nests on platforms on the ground in marshes, up to 10 feet high in shrubs and trees growing in water, in sites well covered with vegetation. FOOD: Forages by probing in soft mud flats and in flooded fields. Eats mostly crayfish, but also snakes, grasshoppers, cutworms and other grubs, and leeches. REFERENCES: Burger and Miller 1977, DeGraff et al. 1980, Low and Mansell 1983, Palmer 1962, Terres 1980. 32 White-faced Ibis Plegadis chihi L 19- RANGE: Breeds locally from central California, eastern Oregon, southern Idaho, Montana, southern North Dakota, and southwestern Minnesota south to Mexico; and from eastern Texas, southern Louisiana east occasionally to Florida. Wanders, at least casually, north to southern Canada. Winters from southern California and the Gulf Coast of Texas and Louisiana south to Mexico. STATUS: Uncommon; nesting populations have been greatly reduced due to use of pesticides and herbicides by rice farmers. HABITAT: Inhabits wetland habitats, preferably marshes and sloughs or ponds surrounded by low bushes or willows, and emergent vegetation such as bulrushes. Also in tule or bulrush swamps, in centers of ponds, and in irrigated rice fields. Roosts in marshes in the evenings. SPECIAL HABITAT REQUIREMENTS: Freshwater marshes and sloughs. NEST: Colonial nester, sometimes with or near colonies of great blue and black-crowned night-herons, or snowy egrets. Generally nests in large beds of bulrushes or reeds several feet above water; on floating mats of dead plants, in cattails and hardstem and alkali bulrush; or infrequently on dry land. On land, prefers to nest on the ground among low shrubs and mixed forbs rather than in grass or cactus. Generally nests in areas well covered with vegetation. FOOD: Feeds by probing in freshwater marshes. Consumes insects, newts, leeches, worms, mollusks, crustaceans (especially crayfish), frogs, fishes, and some snails. After nesting season, feeds in larger marshes as well as in irrigated fields. REFERENCES: Burger and Miller 1977, Oberholser 1974a, Palmer 1962, Ryder 1967, Terres 1980. 33 Fulvous Whistling-Duck Dendrocygna bicolor RANGE: Breeds from southern California to southwestern Arizona, and from central and eastern Texas and the Gulf Coast of Louisiana, south to Mexico; locally in southern Florida. Wanders sporadically throughout North America. Winters from southern California, southern Arizona, the Gulf Coast, and southern Florida south to Mexico. STATUS: Fairly common, but population levels fluctuate. HABITAT: Inhabits marshlands, wet meadows, and in North America, primarily flooded agricultural land and rice fields. Does not ordinarily frequent woodlands. Loafs among dense bulrushes or far out on marshy ponds. SPECIAL HABITAT REQUIREMENTS: Broad, open marshlands. NEST: Prefers to nest in rice fields on low, contour levees, as well as a few inches over water among rice plants and wood growing between levees. Also nests in bulrushes, in knotgrass and dense beds of cattails, on hummocks in marshes, at the edge of ponds and swamps, or in rank tall grasses of wet meadows; rarely in tree cavities. FOOD: Feeds primarily at night, walking about on land gleaning seeds of grasses and weeds. Also tips up in shallow water, and visits cornfields for waste grain. Has a mainly vegetarian diet that includes rice, millets, nutgrass, knotgrass, signalgrass, water shield, and alfalfa. REFERENCES: Baldwin et al. 1964, Bellrose 1976, Cottam and Glazner 1959, Terres 1980, USDA 1981. 34 Black-bellied Whistling-Duck Dendrocygna autumnalis RANGE: Resident in southern Arizona, central and southeastern Texas, and south into South America. STATUS: Rather common within its breeding range, but only a straggler outside. HABITAT: Prefers open woodlands, groves or thicket borders of ebony, mesquite, retama, huisache, and cacti near banks and shallows of rivers, ponds, or marshes. In semiarid southern Texas, it has adapted to some constructed “water habitats” such as small reservoirs and stock tanks. May loaf on shores of small ponds and frequently perches in trees; usually does not alight or swim on deep water. SPECIAL HABITAT REQUIREMENTS: Shallow waters or wetlands, and natural cavities in trees or depressions in ground near water for nesting. NEST: Nests in cavities of elms, willows, live oaks, ebony, mesquite, hackberry, and other trees; also in nest boxes and on the ground. Nest trees may be standing in or located up to 3,000 feet from water, while ground nests are usually in grazed brush pastures, well hidden under low shrubs, and usually near water. Uses natural cavities with entrance holes ranging from 4 by 4.75 inches to 7 by 12.5 inches, and located about 9 feet above ground or water. FOOD: Often feeds at night by grazing in stockyards, pastures, or fields, or by tipping or standing in shallow water. Has a primarily vegetarian diet that includes many cultivated plants, stock foods, and native plants. Consumes about 8 percent animal material, including insects, mollusks, and snails. REFERENCES: Baldwin et al. 1964; Bellrose 1976; Bolen 1967a, 1967b; Bolen and Forsyth 1967; Johnsgard 1975b; Meanley and Meanley 1958; Terres 1980. 35 Tundra Swan Cygnus columbianus (formerly Whistling Swan and Bewick’s Swan) / im. L36" RANGE: Breeds from northwestern Alaska south to St. Lawrence Island and the Alaska Peninsula, and east near the Arctic Coast of Baffin Island, thence south around Hudson Bay to Churchill and the Belcher Islands. Winters mainly near coast from southern Alaska through British Columbia to Pacific states and northern Baja California (casual): also in southern Great Basin to northern New Mexico; and in mid-Atlantic states, rarely on Gulf Coast. STATUS: The most common and widespread swan in North America. HABITAT: Inhabits lakes, ponds, sluggish streams, and occasionally swamp bogs on open tundra while breeding; may be found along coas a estuaries when not breeding. During winter, primarily found on sizeable reservoirs; shallow, productive lakes of the interior; other sheltered freshwater habitats; or on coastal bays and estuaries. SPECIAL HABITAT REQUIREMENTS: Open water or wetlands on Arctic tundra. NEST: Builds nests on the ground along water’s edge, on hummocks in marshes or tidal meadows, or on low hills up to one-half mile from water; seems to prefer to nest on small islands in shallow tundra pools. Rarely nests on level stretches in marsh or meadow areas. FOOD: Feeds by plunging head under water and uprooting aquatic veqetation, preferably in shallow water, and occasionally by tipping up in deeper water, or by grazing in fields. Most commonly eats aquatic plants, but also eats waste corn, soybeans, shoots of winter wheat, grasses, sedges, and thin-shelled mollusks. REFERENCES: Bellrose 1976, Terres 1980, Tohish in Farrand 1983a, Verner and Boss 1980, USDA 1981. 36 Trumpeter Swan Cygnus buccinator RANGE: Breeds locally throughout Alaska and from southern British Columbia and southwestern Saskatchewan south to southeastern Oregon, eastern Idaho and northwestern Wyoming. Was introduced and is now established at Ruby Lake in Nevada and in southwestern South Dakota. Winters from southern Alaska, western British Columbia, southern Alberta (rarely) and Montana south to northern (casually southern) California, occasionally to Utah, New Mexico, and eastern Colorado. STATUS: Once near extinction, the population has increased to more than 4,000 birds. HABITAT: Typically found in open boreal forest; prefers large shallow, fertile marshes or lakes (up to 4 feet deep) with a profusion of submerged and emergent aquatic plants, and generally untimbered but well-vegetated shorelines. During winter, prefers shallow lakes, streams, and ponds with open water that are bordered by some level and open terrain. SPECIAL HABITAT REQUIREMENTS: Shallow, sheltered waters that do not have a fluctuating water level, and some margins of emergent vegetation. NEST: Nests on the ground on any site above the general level of the marsh terrain, preferably on muskrat houses surrounded by water 1 to 3 feet deep. Also nests on shore in sedges, bulrushes, cattails, rushes, or in horsetail. Has a nesting territory that ranges from 70 acres along irregular shorelines to 150 acres along straight shorelines. FOOD: Feeds primarily in shallow waters of lakes or open marsh, digging up roots and tubers of aquatic plants, or snapping off plant parts with bill; rarely feeds on land. Eats a variety of marsh and aquatic plants. REFERENCES: Banko 1960, Bellrose 1976, Johnsgard 1975b, Terres 1980, Van Wormer 1972. 37 Greater White-fronted Goose Anser albifrons L20” RANGE: Breeds from northern Alaska south to Bristol Bay and Cook Inlet, and east across northern Yukon, northern Mackenzie and southern Victoria Island to northern Keewatin. Winters from southern British Columbia south along the coastal states; on the Gulf Coast from Texas and Louisiana south to Mexico; and rarely in the lower Mississippi Valley from Missouri southward. STATUS: Common throughout range. HABITAT: Inhabits the borders of shallow marshes and lakes, riverbanks and islands, deltas, dry knolls, and hills near rivers and ponds in Arctic tundra. Generally found in areas characterized by dwarf birch, willows, bilberries, crowberries, Labrador tea, cassiope, raspberries, dryas, sedges, horsetails, cottongrasses, bluegrass, fescue, arctic grass, sphagnum moss in depressions, and reindeer moss and cetaria on drier sites. Rests on shallow ponds and sloughs in marshes. Winters in sheltered inland and coastal marshes and on open terrain and pasturelands with small bodies of water. SPECIAL HABITAT REQUIREMENTS: Wetlands in Arctic tundra. NEST: Typically nests in depressions on the ground in tall grass bordering tidal sloughs or in sedge marshes, usually within 300 feet of water, or on hummocks along rivers, streams, and lakes. Generally does not nest in colonies but may be found in loose colonies of 15 to 20 pairs in favored locations. FOOD: Primarily grazes on marsh grasses, freshly sprouted grain in fields, and fresh growth in burned-over pastures. Sometimes feeds heavily on aquatic plants and in grain fields after harvest. In the Arctic, consumes tundra plants, aquatic insects and their larvae, and berries. REFERENCES: Bellrose 1976, Dzubin et al. 1964, Johnsgard 1975b, Pough 1951, Terres 1980. 38 Snow Goose Chen caerulescens RANGE: Breeds from northern Alaska east along the Arctic Coast and islands of Canada to Baffin Island, south to Southhampton Island and along both coasts of Hudson Bay to the head of James Bay. Winters from the Puget Sound of British Columbia and Washington south to the interior valleys of California and Mexico; in southern New Mexico; from Kansas and Missouri south to the Gulf Coast; and along the Atlantic Coast from New York to Florida. During migration, found on large staging areas in the Dakotas, Minnesota, Iowa, and Nebraska. There are two races of the snow goose, the “lesser” and the “greater.” The lesser snow goose has two color phases—a dark phase, or blue goose, and a white phase—while the greater is believed to only have a white phase, and generally breeds farther north than the lesser. STATUS: Locally abundant. HABITAT: Inhabits islands of the Canadian Arctic Archipelago or is found within 5 miles of salt water on flat tundra of marsh grasses and sedges, in limestone basins, on islands of river deltas, or on plains usually drained by large rivers that open early in the season. During winter, uses both freshwater and saltwater marshes and wet prairies. SPECIAL HABITAT REQUIREMENTS: Wetlands on arctic tundra. NEST: Nests in a shallow depression on the ground in large, loose colonies, on dry sites, primarily in unspoiled, primitive areas. Nests, well concealed by tundra grasses and sedges, as close as 15 to 20 feet from each other on flat land. FOOD: Feeds by browsing in cultivated fields on winter wheat, in pastures on sprouting grasses, or on waste grain in stubble fields, also by digging out bulbous roots and soft parts of sedges, rushes, marsh grasses, and aquatic plants. REFERENCES: Bellrose in Farrand 1983a, Cooch 1964, Lemieux 1959, Terres 1980, Verner and Boss 1980. 39 Ross’ Goose Chen rossii L16 RANGE: Breeds primarily in the Queen Maud Gulf area of northern Mackenzie and northwestern Keewatin, but also on southern Southhampton Island and along the west coast of Hudson Bay south to Cape Churchill. Winters almost exclusively in the Central Valley of California and the Salton Sea; also in small numbers along the Rio Grande, New Mexico, and Gulf Coast of Texas. STATUS: Rare a few decades ago, now quite abundant in recent years. HABITAT: Inhabits island-studded lakes and deltas of low tundra country. Prefers islands that rise 10 to 20 feet above water level, are covered with rocks and shrubs interspersed with areas of open, level ground, and are surrounded by shallow water (under 5 to 6 feet deep) extensive enough to discourage predators from swimming across. During winter, found on freshwater and brackish marshes and on wet prairies, often in association with the snow goose. SPECIAL HABITAT REQUIREMENTS: Islands in Arctic tundra lakes. NEST: Nests on the ground in loose colonies, preferably on islands in mixed habitats of dwarf birch and rocks, or occasionally along river or lake shores if islands are unavailable. FOOD: Eats grain and new green growth of grasslands and grain fields. REFERENCES: Barry 1964, Bellrose in Farrand 1983a, Johnsgard 1975b, Ryder 1967, Terres 1980. 40 Canada Goose Branta canadensis L 16 - 26 ” RANGE: Breeds from the Arctic Coast of Alaska and northern Canada east to Baffin Island, south to central California, east to western Tennessee, southern Ontario and Quebec, and Newfoundland. Winters from south-coastal and southeastern Alaska, British Columbia and southern Alberta east to the Atlantic Coast of Newfoundland, and south to Mexico, the Gulf Coast and northern Florida. STATUS: Common; 11 subspecies of the Canada goose are currently recognized. HABITAT: Found in a variety of habitats near water, from forested and prairie regions to tundra, breeding on swamps, marshes, meadows, rivers, banks of lakes and ponds, and on islands. Winters in tidewater areas, marshes, inland refuges, and in flooded fields. SPECIAL HABITAT REQUIREMENTS: Elevated habitat feature or artificial structures near water for nesting. NEST: Usually nests on the ground near water (generally within 150 feet), preferably on a slightly elevated site that is isolated and affords good visibility of the surrounding area. Prefers muskrat houses for nesting but will also nest on small islands lacking tall growth, haystacks, rocky cliffs, hummocks, ridges of silt, pond banks, beaver lodges, and occasionally abandoned nests of ospreys, ravens, owls, or herons. Has successfully adapted to nesting on artificial structures. FOOD: Essentially a grazer, preferring young, green tender plants. Consumes various grasses and forbs, both terrestrial and aquatic. Consumes agricultural crops as primary food during migration and winter. Also consumes small amounts of insects, insect larvae, mollusks, and small crustaceans. REFERENCES: Bellrose 1976, Hansen and Nelson 1964, Terres 1980, VanWormer 1968. 41 Wood Duck Aix sponsa 9 L 13 Vi" RANGE: Breeds in western North America from southern British Columbia and southwestern Alberta south to central California and western Montana; in eastern North America from east-central Saskatchewan east to Prince Edward Island and Nova Scotia south (east of the Rockies) to central and southeastern Texas and the Gulf Coast. In the West, winters irregularly throughout the breeding range; in the East, winters primarily in the southern parts of the breeding range. STATUS: Common; population has increased in recent years primarily because of the availability of artificial nest structures and protection for most of the year. HABITAT: Inhabits woodlands near shallow, quiet inland lakes, swamps, river bottoms, ponds, marshes, and streams where nest sites are available. Important forest types are central and southern floodplain forests, red maple swamps, temporarily flooded oak forests, and northern bottomland hardwoods. Prefers areas with many perching sites. SPECIAL HABITAT REQUIREMENTS: Nest holes in trees or nest boxes in or near still or slow-moving water. NEST: Prefers to nest in natural cavities 20 to 50 feet above ground with entrance holes of 4 inches in diameter, cavity depths of 2 feet, and cavity bottoms measuring 10 by 10 inches. Uses nest trees in (or up to one-half mile from) water 3 to 18 inches deep. Readily accepts nest boxes provided with nesting materials of wood shavings or sawdust. FOOD: Eats about 90 percent plant material. Forages in ponds, marshes, sluggish streams, or along wooded banks for floating duckweeds, baldcypress cones and galls, seeds and tubers, wild rice, acorns, beechnuts, hickory nuts, grapes, berries, corn, and wheat. Also eats some invertebrates, such as spiders and aquatic insects. REFERENCES: Bellrose 1976, Grice and Rogers 1965, McGilvrey 1968, Palmer 1976b, Terres 1980. 42 Green-winged Teal Anas crecca cf 9 L lOW RANGE: Breeds from Alaska, northwestern and southern Mackenzie to north-central Labrador and Newfoundland south to central Oregon, Colorado, southern Ontario and Quebec, and Nova Scotia; Breeds locally from southern California east to southern New Mexico, Iowa, and Pennsylvania, and on the Atlantic Coast to Delaware. Winters from southern Alaska and southern British Columbia to New Brunswick and Nova Scotia south to Central America; also winters in the Hawaiian Islands. STATUS: Relatively common throughout range. HABITAT: Inhabits inland waters with dense rushes or other emergent vegetation on mixed and shortgrass prairies, and northern boreal forests. May be found resting on mudbanks or stumps, or perching on low limbs of dead trees. Winters in both freshwater and brackish marshes, ponds, streams, and estuaries. SPECIAL HABITAT REQUIREMENTS: Lakes, marshes, ponds, pools, and shallow streams. NEST: Nests in a depression on dry ground located at the base of shrubs, under a log, or in dense grass, usually 2 to 300 feet (but up to one-quarter mile) from water. FOOD: Feeds in shallow marshes or temporarily flooded fields by dabbling, or by probing on mud flats. Consumes a diet that is about 90 percent vegetative, consisting of seeds of aquatic plants, grains, berries, wild grapes, mast, and (to a lesser extent) the vegetative parts of aquatic plants. Also eats some insects, small mollusks, and crustaceans. REFERENCES: Bellrose 1976, Harrison 1975, Palmer 1976a, Terres 1980. 43 American Black Duck Anas rubripes (formerly Black Duck) RANGE: Breeds from northern Saskatchewan to Labrador and Newfoundland south to northern South Dakota, northern Illinois, central West Virginia, and on the Atlantic Coast to North Carolina. Winters from southeastern Minnesota and central Wisconsin to New Brunswick and Nova Scotia south to southern Texas, the Gulf Coast and south-central Florida. Winters as far north as open water and food are available. STATUS: Common, but the population is declining. HABITAT: Inhabits a wide variety of wetland habitats along the coast and in woodlands, including open boreal forests and mixed hardwoods; sometimes is found in stubble fields or berry barrens. Generally is extremely adaptable as long as there is some source of water. Prefers to winter on brackish marshes bordering bays, estuaries, and agricultural lands but may also be found on lakes, reservoirs, rivers, freshwater marshes, and old rice fields. NEST: Generally nests in a hollow on dry, slightly elevated ground in wooded areas, well-concealed in thickets, briars, shrubs or grasses, usually near (but possibly a mile or more from) water. Occasionally uses old crow or hawk nests and natural or excavated cavities in trees or tops of rotted stumps. FOOD: Forages by dabbling in shallow water, and by gleaning and grazing in fields. Has a diet that varies widely, depending on habitat. In fresh and brackish areas eats mostly plants; in marine environments eats mostly animals. Consumes primarily blue mussels, submerged aquatic plants, waste grains, acorns, seeds of marsh plants, crustaceans, earthworms, amphibians, and fishes. REFERENCES: Bellrose 1976, Benson 1968, Coulter and Mendall 1968, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976a. 44 Mottled Duck Anas fulvigula RANGE: Breeds along the Gulf Coast from southern Louisiana and Texas into Mexico; in peninsular Florida; and locally inland in southeastern Colorado, western Kansas, Oklahoma, and northeastern Texas. Winters in the breeding range and casually along the entire Gulf Coast into Mexico. In Florida, does not migrate. STATUS: Locally common. HABITAT: Primarily inhabits extensive coastal marshes with a good interspersion of ponds but is also found in rice fields and on ponds and stream banks in pasture and farmlands. In Florida, inhabits ponds and lakes of pine flatwoods, everglades, cultivated and fallow fields, fresh and brackish marshes, mangrove swamps, and baldcypress-watertupelo hummocks. SPECIAL HABITAT REQUIREMENTS: Freshwater and marine wetlands. NEST: Builds a nest that is usually well concealed in a clump of grass, or on the ground under a bush, in meadows, generally within 500 feet of water. Prefers to nest near coastal marshes, but will also nest near ponds, bayous, or ditches. Is shy and sensitive to human disturbance. FOOD: Feeds heavily on animal foods such as fish, crayfish, snails, and insects; also eats seeds of millet, rice, grasses, and aquatic plants. REFERENCES: Beckwith and Hosford 1957, Bellrose 1976, Sincock et al. 1964, Singleton 1968. 45 Mallard Anas platyrhynchos (includes Mexican Duck) RANGE: Breeds from northern Alaska east to southern Keewatin and across to southern Maine south to California, the southern Great Basin and New Mexico, and from Oklahoma east through the Ohio Valley to Virginia. Winters generally from southern Alaska and southern Canada south to central Mexico. Introduced and established in the Hawaiian Islands. STATUS: The most common and widely distributed duck in North America. HABITAT: Inhabits ponds, lakes, rivers, streams, marshes, wet meadows, and wooded swamps of primarily mixed and shortgrass prairie; also inhabits boreal forest region and sub-arctic deltas. Winters on inland ponds and rivers with some open water; less commonly in coastal marshes. NEST: Typically nests on the ground in dry or slightly marshy areas within 300 feet of water, sometimes as far as 1.5 miles away in grasslands. Conceals nest well in snowberry clumps, among weeds and grasses, in pastures, stubble, or cultivated fields, or in marsh vegetation; rarely in cavities, on hollowed tops of stubs, or in tree crotches. FOOD: Feeds by dabbling in shallow waters of ponds, sloughs, lakes, streams, and swamps, and by grazing and gleaning in grainfields and meadows. Consumes seeds, acorns, nuts, waste grains, aquatic insects, mollusks, tadpoles, frogs, small fish, and fish eggs. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976a, Terres 1980. 46 Northern Pintail Anas acuta (formerly Pintail) L I 8 I/ 2 ' RANGE: Breeds from northern Alaska across northern Canada to northern and eastern Quebec, New Brunswick, and Nova Scotia to California, across to the Great Lakes, St. Lawrence River, and Maine. Winters from southern Alaska south to northern New Mexico, and east to central Missouri and the Ohio Valley (uncommonly); along the Atlantic Coast from Massachusetts, south throughout the southern United States to South America. STATUS: Abundant in the West and common in the East. HABITAT: Found in a wide variety of habitats, but typically inhabits open country with low vegetation and with many scattered small, shallow bodies of water. Frequents lakes, rivers, marshes and ponds in grasslands, barrens, dry tundra, open boreal forest, and cultivated fields. Winters on freshwater and brackish coastal marshes, shallow lagoons, mudflats along rivers, and sheltered marine waters. SPECIAL HABITAT REQUIREMENTS: Drakes need mudbanks or exposed water margins for resting; also shallow wetlands for feeding. NEST: Often builds a nest in a hollow on dry ground, sometimes concealed by grasses or shrubs, usually within 300 feet (occasionally a half mile) from water. Nests in stubble fields, in a dry portion within a large marsh, or in lightly grazed pasture, but generally avoids timbered or extensively brushy areas. FOOD: Prefers to feed in shallow waters of marshes, ponds, and wet meadows, or in grainfields. Primarily a seed-eater; mostly (87 percent) consumes vegetative diet, consisting of seeds of pondweeds, sedges, grasses, smartweeds, and cultivated grains; also takes some fairy shrimp, snails, earthworms, mollusks, crustaceans, dipteran larvae, and other insects. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Krapu 1974, Palmer 1976a. 47 Blue-winged Teal Anas discors d Lir RANGE: Breeds from east-central Alaska and southern Mackenzie to southern Quebec and southwestern Newfoundland, south to northeastern California, east across to central Louisiana, central Tennessee and eastern North Carolina. Winters from southern California to western and southern Texas, the Gulf Coast and North Carolina on the Atlantic Coast south to South America. STATUS: Common throughout range. HABITAT: Prefers wetlands on rolling tallgrass prairie but is also found in mixed shortgrass prairie and boreal and deciduous forests. More of a shoreline inhabitant than one of open water, prefers calm water or sluggish currents to fast water. Uses rocks protruding above water, muskrat houses, trunks or limbs of fallen trees, or bare stretches of shoreline or mudflats as resting sites. Winters on shallow inland freshwater marshes and on brackish and saltwater marshes. SPECIAL HABITAT REQUIREMENTS: Marshes, sloughs, ponds, lakes, and sluggish streams. NEST: Builds nests on dry ground in dense grassy sites such as bluegrass, hayfields, and sedge meadows, where the vegetation ranges from 8 to 24 inches high at the onset of nesting, or under bushes, usually within several hundred yards of open water; occasionally on a sedge tussock or muskrat house, in slough grass, or in alfalfa fields. In good habitat nests communally. FOOD: Prefers to feed on mud flats, in fields, or in shallow water where there is floating and shallowly submerged vegetation plus abundant small aquatic animal life. Consumes a diet that is 70 percent vegetative, consisting of seeds of sedges; grasses, pondweeds, and smartweeds; stems and leaves of aquatic plants; and snails, mollusks, crustaceans, and insects. REFERENCES: Bellrose 1976, Bennet 1938, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976a. 48 Cinnamon Teal Anas cyanoptera cf RANGE: Breeds from southern British Columbia east to southwestern Saskatchewan (probably), and south into Mexico. Winters from central California, southern Nevada, central Utah, southeastern Arizona, southern New Mexico and central Texas south to South America. STATUS: Common in the West. HABITAT: Inhabits small, shallow wetlands, including areas with alkaline waters, but may also be found around larger and deeper lakes. Winters primarily on freshwaters, though occasionally found in marine habitats. SPECIAL HABITAT REQUIREMENTS: Shallow lake margins, ponds bordered by tule and grasses, sloughs, marshes, sluggish streams, reservoirs, and irrigation ditches. NEST: Nests on the ground in dense grasses under 2 feet high, in cattails or reeds near water, or in a hollow in the ground, often 100 feet or more from water. Broods may be moved as far as a mile from the nest site to good brood cover of lush emergent vegetation adjacent to water with abundant food. FOOD: Feeds by tipping up in shallow water, grazes in grass and in fields, or probes in mud for its food, which is 80 percent vegetative. Primarily consumes seeds and vegetative parts of pondweeds, bulrushes, sedges, smartweeds and grasses; also takes mollusks and insects. REFERENCES: Bellrose 1976, Grinnell and Miller 1944, Johnsgard 1975b, Low and Mansell 1983, Palmer 1976a, Verner and Boss 1980. 49 Northern Shoveler Anas clypeata 9 L14" RANGE: Breeds from northern Alaska to northern Manitoba, south to northwestern and eastern Oregon, northern Utah, Colorado, Nebraska, and Missouri, and central Wisconsin. Winters from the coast of southern British Columbia to central Arizona east to the Gulf Coast and South Carolina on the Atlantic Coast south to South America. STATUS: Fairly common; more abundant west of the Mississippi River. HABITAT: Prefers shallow prairie marshes, particularly those with abundant plant and animal life floating on the surface, but also occupies potholes, sloughs, and marshes in taiga, forests, and (less commonly) cultivated country. It tolerates a wide range of water conditions, from clean and clear to muddy; flowing to stagnant; considerably alkaline, and even heavily polluted. Likes to have mudbanks or low sloping shorelines with short or flattened vegetation for loafing. Winters in freshwater and brackish habitats. SPECIAL HABITAT REQUIREMENTS: Shallow waters with muddy bottoms, surrounded by dry grassy areas for nesting. NEST: Nests on dry ground in a slight hollow, preferably in short grasses within 300 feet of water, but will nest in hayfields, meadows, and rarely bulrushes if grasses are not available. Seldom nests in weed patches, and avoids woody vegetation such as willows. FOOD: A filter feeder; prefers to feed in shallow waters, but will actively feed in deep waters containing submergent and surface vegetation. Consumes a considerable amount of microscopic animal life such as ostracods, copepods, and similar crustaceans, and macroscopic animal life as well. Eats fingernail clams, mollusks, and insects for one qurter of the diet. Also eats grasses, sedges, water lilies, pondweeds, bulrush seeds, algae, and smartweeds. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976a, Poston 1974. 50 Gadwall Anas strepera $ L UV 2 " RANGE: Breeds from southern Alaska and southern Yukon to the New Brunswick-Nova Scotia border, south locally to southern California, northern Texas, central Minnesota, and northern Pennsylvania and on the Atlantic Coast to North Carolina. Winters from southern Alaska, southern British Columbia and Colorado to southern South Dakota, Iowa, the southern Great Lakes and Chesapeake Bay on the Atlantic Coast south to Mexico and the Gulf Coast. STATUS: Uncommon, but numbers have increased substantially during the past 2 decades and the range is extending eastward. HABITAT: Inhabits prairie marshes, sloughs, ponds or small lakes in grasslands in both freshwater and brackish habitats. Generally avoids wetlands bordered by woodlands or thick brush, preferring those bordered by dense, low herbaceous vegetation or shrubby willows and with grassy islands. Prefers to winter in freshwater, marshy habitats but can be found on open water of any kind. SPECIAL HABITAT REQUIREMENTS: Shallow water for feeding; marshes or grassy areas near water for nesting. NEST: Nests on the ground on a well-drained site on islands in lakes, in upland meadows or pastures, in alfalfa fields, or on prairies, usually within 150 feet of water. Prefers to nest in uplands rather than over water, especially in dense, coarse herbaceous vegetation and under shrubby willows. FOOD: Prefers to feed along shallow marsh edges with abundant aquatic plant life, but also feeds in open water more than other dabblers. Sometimes feeds in stubble fields for grain, or in woods for acorns. Consumes mainly leaves and stems of aquatic plants; also eats insects, mollusks, crustaceans, amphibians, and fishes. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976a, Terres 1980. 51 American Wigeon Anas americana 9 L 14' RANGE: Breeds from central Alaska and central Yukon to New Brunswick and southern Nova Scotia, south to northeastern California, central Colorado, South Dakota, southern Ontario, and northern New York, sporadically to the Atlantic Coast. Winters from southern Alaska to southern Nevada, sporadically across the central United States to the southern Great Lakes and Ohio Valley, and on the Atlantic Coast from Nova Scotia south throughout the southern United States to Central America. STATUS: Common. HABITAT: Inhabits freshwater wetlands and lakes from tundra to shortgrass and mixed prairie, preferring permanent to temporary waters. Commonly associates with diving ducks, and in winter frequents coastal marshes and bays, wet meadows, and shallow freshwater and brackish ponds. SPECIAL HABITAT REQUIREMENTS: Large lakes, ponds, marshes, sluggish streams and rivers, with open water and exposed shoreline. NEST: Nests in a hollow on dry ground on an island or on shore, in tall grasses or weeds, or at the base of a tree or bush, as far as 400 yards from water. FOOD: Feeds by grazing and gleaning in wet or dry pastures and fields, by dabbling on the water surface in shallow water along marsh edges and sloughs, and by scavenging for wild celery after diving ducks have torn plants loose from bottom. Primarily vegetarian, eats mainly leaves, stems, and buds of aquatic plants; also feeds on waste grains, mollusks, crustaceans, and insects. REFERENCES: Baldwin et al. 1964, Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Low and Mansell 1983, Palmer 1976a, Terres 1980, Verner and Boss 1980. 52 Canvasback Aythya valisineria RANGE: Breeds from central Alaska and northern Yukon to western Ontario and south to south-coastal Alaska; locally in inland areas to northeastern California across to northern Utah, central New Mexico, northwestern Iowa, and southern Ontario. Winters along the Pacific Coast from the central Aleutians and south-coastal Alaska south to Baja California, from Arizona and New Mexico to the Great Lakes, and, on the Atlantic Coast, from New England south to the Gulf Coast and Mexico. STATUS: Locally common; numbers are declining due to loss of breeding habitat through drainage and drought. HABITAT: Prefers shallow prairie marshes, 10 acres or less, or other permanent wetlands with stable water levels, bordered by cattails and bulrushes, with little, if any, wooded vegetation around the shoreline. Large lakes of 150 acres or more, marshes, and rivers with submerged beds of sago pondweed are favored during migration. Winters primarily on estuaries and sheltered bays, sometimes on deep, freshwater lakes, where wild celery and pondweeds thrive. SPECIAL HABITAT REQUIREMENTS: Marshes, ponds, lakes, and rivers bordered by emergent vegetation and with enough open water for taking off and landing. NEST: Usually nests over water 6 to 24 inches deep in bulrushes, reeds, or cattails, sometimes on a muskrat house, rarely on dry ground. Attaches nest to surrounding plants or built on a mat of floating dead plants, 3 to 60 feet from edge of open water. FOOD: Dives in shallow water, usually 3 to 12 feet deep, for food, which is 80 percent vegetative material. In the Northeast, prefers seeds and vegetative parts of wild celery; in the Southeast and the West, primarily consumes pondweeds; also feeds on water plantains, grasses, sedges, mollusks, and insects. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Evans and Bartels 1981, Johnsgard 1975b, Palmer 1976b, Stoudt 1982, Terres 1980. 53 Redhead Aythya americana d 9 L14 RANGE: Breeds from central British Columbia and southwestern Mackenzie to northwestern and central Minnesota, south to southern California, the Texas Panhandle and northern Iowa; locally in south- central and southeastern Alaska, and sporadically in eastern North America. Winters from British Columbia on the Pacific Coast, in the interior from Nevada to the middle Mississippi and Ohio Valleys and the Great Lakes, and from New England on the Atlantic Coast south throughout the southern United States to Mexico. STATUS: Common; population declining in the eastern United States. HABITAT: Inhabits freshwater marshes, sloughs, ponds, and shallow lakes bordered by hardstem bulrush, cattails, reeds, or sedges in prairies and intermountain parks. Winters primarily on freshwater and brackish lakes, rivers, and estuaries, in areas well protected from heavy wave action. SPECIAL HABITAT REQUIREMENTS: Wetlands at least 1/2 acre in size bordered by permanent, dense emergent vegetation, with stable water levels during the nesting season. NEST: Usually nests in emergent vegetation or on a floating mat of dead plant material over shallow water 6 to 14 inches deep, fairly close to shore, but occasionally on dry ground or over water 4 feet deep. Tends to be semi-parasitic, sometimes laying eggs in nests of other waterfowl, especially the canvasback; also incubates own nests. Prefers to rear broods in potholes at least 1 acre in size, with deeper waters than those used for nesting. FOOD: Feeds in marshes, sloughs, and ponds more than other diving ducks, mainly in water less than 6 feet deep. Has a diet that is 90 percent vegetative, consisting primarily of seeds and vegetative parts of aquatic plants and including some insects and shellfish. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Lokemoen 1966, Palmer 1976b, Tate and Tate 1982, Weller 1964. 54 Ring-necked Duck Aythya collaris RANGE: Breeds in east-central and southeastern Alaska, and from central British Columbia and northwestern and southern Mackenzie to Newfoundland and Nova Scotia, south to northeastern California, southeastern Arizona, northern Illinois, and Massachusetts. Winters on the Pacific Coast from southeastern Alaska, in the interior from southern Nevada to the lower Mississippi and Ohio Valleys, and on the Atlantic Coast from New England south through the southern United States to Panama. STATUS: Common. HABITAT: Inhabits shallow, dense bogs, swamps, and marshes, especially those with sweetgale or leatherleaf cover, from 1 to 2,000 acres in size, typically having a pH range of 5.5 to 6.8, and preferably near or in woodlands. It also uses small potholes, sloughs, and beaver flowages near larger wooded lakes or rivers with submerged and emergent vegetation. Winters on fresh or brackish marshes, lakes, and estuaries, rarely on strictly saline waters. SPECIAL HABITAT REQUIREMENTS: Wetlands with an expanse of open water. NEST: Nests on floating mats of vegetation, among hummocks, in clumps of marsh vegetation or on islands, on relatively dry sites usually within a few feet of water; seldom in emergent vegetation over water. FOOD: Prefers to feed in shallow water, usually less than 6 feet deep. Consumes a diet that is 80 percent vegetative, consisting mostly of a few plant groups; seeds, bulbs, and succulent parts of waterlilies, pondweeds, sedges, grasses, and smartweeds. Also consumes some aquatic insects and mollusks. REFERENCES: Bellrose 1976, DeGraff et al. 1980, Johnsgard 1975b, Mendall 1958, Palmer 1976b. 55 Greater Scaup Aythya marila d RANGE: Breeds from western Alaska to southern Keewatin, around Hudson and James Bays, and northern Quebec; casually or irregularly south to southeastern Alaska, northwestern British Columbia, central Manitoba, and southeastern Michigan. Winters along the Pacific Coast from the Aleutians and southeastern Alaska south to Baja California, in the eastern Great Lakes, from the Ohio and lower Mississippi Valleys south to the Gulf Coast, and on the Atlantic Coast from Newfoundland south to Florida. STATUS: Locally common. HABITAT: Inhabits lakes, ponds, and marshes from forested tundra to richly vegetated low tundra. During the breeding season, drakes rest along shorelines or on shoals, while in late fall and winter, both sexes form large rafts in open water, even in the open ocean well beyond the breakers. Winters on brackish and saltwater bays and estuaries, less commonly on large inland waters. SPECIAL HABITAT REQUIREMENTS: Scattered wetlands in forested to open tundra. NEST: Nests on the ground in a slightly elevated spot, in grasses on the tundra, usually near the shores of lakes or ponds, but up to 3,000 feet from water. Sometimes nests on islands, in marshes, or above or in water among rushes or wild rice. FOOD: An expert diver; dives to depths of at least 20 feet, remaining underwater seeking food for up to a minute. A saltwater bird most of the year; prefers to feed in shoals with water less than 20 feet deep or in shellfish beds, also in freshwater lakes and ponds. Consumes a diet that is about half vegetative and half animal, including snails, aquatic insects, tadpoles, small fishes, and seeds in summer. During winter, also eats mollusks, crabs, barnacles, and other crustaceans. Also feeds on muskgrass, sea lettuce, eelgrass, wild celery, and widgeon grass. REFERENCES: Cottam 1939, Johnsgard 1975b, Palmer 1976b, Terres 1980, Vickery in Farrand 1983a. 56 Lesser Scaup Aythya affinis cr 9 L12” RANGE: Breeds from central Alaska to northern Manitoba and western Ontario, south to southern interior British Columbia, northern Wyoming and northwestern and central Minnesota; casually or irregularly east to southern Ontario and west-central Quebec, and south to northeastern California and Colorado, northern Illinois and northern Ohio. Winters from southern Alaska and southern British Columbia and Utah to the southern Great Lakes Region and New England, south throughout the southern United States to South America. STATUS: Abundant. HABITAT: Inhabits grass-margined wetlands in prairie and forested habitats, with the largest breeding concentrations found in marshes of hardstem bulrush bordering lakes. Winters on sheltered bays, estuaries, coastal marshes, and freshwater lakes, preferring a more sheltered habitat than the greater scaup. SPECIAL HABITAT REQUIREMENTS: Lakes, ponds, potholes, marshes, and sloughs bordered by grasses. NEST: Usually nests in upland areas adjacent to water but also on islands, in wet meadows, in shallows at edges of bays and sloughs among bulrushes or on tussocks in marshes. Conceals nest well in hollows on usually dry ground, in grasses, nettles, low brush, even under driftwood. FOOD: An expert diver, generally feeds in water 5 to 6 feet deep, but may feed to depths of 15 to 20 feet. Consumes plant and animal foods about equally, including seeds and vegetative parts of pondweeds, grasses, sedges, widgeon grass, wild rice, bulrushes, snails and other mollusks, crustaceans, and aquatic insects. REFERENCES: Bellrose 1976, Johnsgard 1975b, Palmer 1976b, Terres 1980. 57 Harlequin Duck Histrionicus histrionicus 9 L12” RANGE: Breeds in western North America from western Alaska and northern Yukon south to Vancouver Island, eastern Oregon and western Wyoming; in the Sierra Nevada of California, and in eastern North America from southern Baffin Island south to central and eastern Quebec and eastern Labrador. Winters along the Pacific Coast from the Pribilof and Aleutian Islands south to central California; on the Atlantic Coast from southern Labrador south to New York (rarely to South Carolina); casually farther inland south to Kansas and West Virginia and Florida. STATUS: Uncommon to locally abundant. HABITAT: Prefers cold, shallow, rapidly flowing mountain streams in forested regions, but also inhabits ponds and lakes, along rocky Arctic shores, and in tundra. Loafs in shallows, or on a rock or log in a stream or ashore. Winters on marine waters, usually 6 to 12 feet deep, in heavy surf along rocky coasts. SPECIAL HABITAT REQUIREMENTS: Shallow, swift streams or rivers, or pothole glacial lakes. NEST: Nests on the ground on an island, in a recess in a stream bank, under bushes or trees, or occasionally in a hollow tree or a cavity among rocks. Usually nests within 6 feet (but sometimes up to 60 feet) from water, with overhead shelter. FOOD: A proficient diver, seems to prefer to feed in rough waters broken by rocks and surf. Mostly (98 percent) consumes crustaceans and mollusks. Also eats stone flies and other insects, fishes, and echinoderms. REFERENCES: Bellrose 1976, Cottam 1939, Johnsgard 1975b, Palmer 1976b, Terres 1980. 58 White-winged Scoter Melanitta fusca a im. $ L16 RANGE: Breeds from northern Alaska to southern Keewatin and northern Manitoba south to central Alaska, northeastern Washington, southern Saskatchewan, northern North Dakota and northern Ontario; possibly farther east. Winters on the Pacific Coast from the Aleutians south to Baja California, on the Great Lakes, and on the Atlantic Coast from Newfoundland south to South Carolina. STATUS: Common to locally abundant. HABITAT: Inhabits waters on open tundra or prairie with dense, low ground cover, preferably lakes larger than 110 acres, 3 to 12 feet deep and with dense submergent vegetation. Uses mixed tundra-taiga less frequently during nesting. In other seasons, can be found on marine and brackish waters along the coasts where the water is shallow over shellfish beds and the bottom is hard and sandy or gravelly, and occasionally on open freshwater. SPECIAL HABITAT REQUIREMENTS: Lakes, ponds, and sluggish streams in northern prairie or tundra. NEST: Nests in a hollow on the ground under dense, low spreading shrubs, preferably gooseberry, snowberry, rose, and raspberry, usually close to water, occasionally up to 450 feet and rarely a half mile away. May nest on islands, commonly uses old nest bowls, and in some instances may occupy active blue-winged teal or gadwall nests. Easily disturbed by human interference such as recreational boating on breeding lakes. Broods are reared in shallow, open water areas with submergent vegetation and some protection from wave action. FOOD: Prefers to dive for shellfish in shallow water, usually less than 6 feet deep. Primarily eats animal foods (90 percent), mostly mollusks; also eats crabs, some fishes, aquatic insects, and sea lettuce. REFERENCES: Brown and Brown 1981, Grosz and Yocom 1972, Palmer 1976b. 59 Common Goldeneye Bucephala clangula a 9 L13” RANGE: Breeds from western Alaska and northern Yukon to central Labrador and Newfoundland, south to central Alaska, northern Washington across to northern Michigan, Maine, New Brunswick, and Nova Scotia. Winters on the Pacific Coast from the Aleutian Islands south to southern California, on the Great Lakes, in the Mississippi and Ohio Valleys, and south to the Gulf Coast, and on the Atlantic Coast from Newfoundland south to Florida; irregularly elsewhere in the interior of the United States. STATUS: Common. HABITAT: Inhabits lakes, ponds, shallow rivers, slow-flowing streams, floodplain forests and bogs, preferably with weedy margins, near or in woodlands with large cavity trees. Breeds in a range that generally coincides with the boreal coniferous forest. Winters on bays, estuaries, rivers, and inland as far north as open water and food are available. SPECIAL HABITAT REQUIREMENTS: Large trees (minimum dbh of 20 inches) with cavities for nesting, near clear, cold, shallow water for feeding. NEST: Nests in tree cavities near or in water, preferably open-top, bucket cavities, but also in old pileated woodpecker cavities, in hollow stumps, in natural cavities, or when cavities are not available, in abandoned buildings, cavities among rocks, and nest boxes. Sometimes may lay joint clutches when cavities are scarce. Prefers nest tree entrances that are 6 to 40 feet above ground or water. Accepts nest boxes that are 9 by 9 by 24 inches in size, with an elliptical entrance 3 1/2 by 4 1/2 inches. FOOD: Prefers to dive for food in water 3 to 12 feet deep. Consumes animal food, as 75 percent of the diet, including crabs, crayfish, mussels, snails, insects, and some fishes. Also eats seeds, tubers, and leafy parts of pondweeds; seeds of pond lilies and bulrushes; and wild celery. REFERENCES: Carter 1958, Cottam 1939, DeGraff et al. 1980, Johnsgard 1975b, Palmer 1976b, Thomas et al. 1979. 60 Barrow’s Goldeneye Bucephala islandica cT 9 L13” RANGE: Breeds from central and southwestern Alaska, southern Yukon and southwestern Alberta south to south-coastal and southeastern Alaska, southern British Columbia, and northern Washington; locally at higher elevations to northwestern Wyoming, and in northeastern Quebec and northern Labrador. Winters along the Pacific Coast from south-coastal and southeastern Alaska south to central California; locally in the interior of western North America; and in the Atlantic region from the upper St. Lawrence drainage and Nova Scotia south to New York, rarely to South Carolina. STATUS: Uncommon. HABITAT: Inhabits lakes and ponds larger than 2 acres in montane, tundra, and subtundra habitats. Prefers moderately alkaline lakes, 5 to 15 feet deep, with a dense growth of submerged aquatic vegetation such as pondweeds and widgeon grass, and bordered by dense stands of bul¬ rushes. Winters on lakes and rivers, and in coastal estuaries and bays where rocky reefs and ledges in shallow water provide feeding grounds. SPECIAL HABITAT REQUIREMENTS: Cavities in trees near open water. NEST: Nests in natural cavities in trees and stumps, in abandoned flicker nest cavities enlarged by natural decay, pileated woodpecker holes, or nest boxes, preferably within 100 feet of water. Occasionally may nest up to a half mile from water. Nests in dead or dying Douglas- fir, aspen, cottonwoods, lodgepole pine, ponderosa pine, and less commonly scrub pine. If tree cavities are not available, nests in holes in the ground or in cavities in rock cliffs or under rocks. FOOD: Dives and forages in open water to depths of 3 to 10 feet for its food, which is 78 percent animal material. Primarily consumes insect nymphs and larvae, and water boatmen; crustaceans, especially crayfish; and some fishes, blue mussels, pondweeds, and wild celery. REFERENCES: Cottam 1939, Bellrose 1976, Johnsgard 1975b, Palmer 1976b, Terres 1980. 61 Bufflehead Bucephala albeola 9 L IOV 2 " RANGE: Breeds from central Alaska to northeastern Manitoba and northern Ontario south to northern Washington, southern Manitoba and locally in southern Ontario; also locally south to the mountains of northern California, and to Wyoming, Iowa, and Wisconsin. Winters from the Aleutian Islands on the Pacific Coast, the Great Lakes, and Newfoundland on the Atlantic, south in coastal states and the Ohio and Mississippi Valleys to the southern United States and Mexico. STATUS: Common. HABITAT: Primarily inhabits small, shallow, fresh or slightly alkaline lakes and ponds, preferably without broad margins of emergent or floating aquatic vegetation, in mixed coniferous-deciduous woodlands north and west of the Great Plains. Logs, stumps, rocks, open shore, and sometimes fence rails near water are used for resting and loafing. Winters in sheltered marine habitats, or on brackish or freshwater. SPECIAL HABITAT REQUIREMENTS: Tree cavities excavated by wood¬ peckers, especially northern flickers, near shallow, fertile waters in forested regions. NEST: Prefers to nest in aspen trees near water containing unaltered northern flicker holes, but will also use pileated woodpecker holes. Also nests in Douglas-fir, balsam poplar, black cottonwood, ponderosa pine, and a few other coniferous and deciduous trees. Nests generally within 75 feet of water and rarely in dense forest. In some areas, accepts nest boxes 7 inches in diameter and 16 inches deep with entrances 2 7/8 inches wide. FOOD: Prefers to forage in shallow water, diving to depths of 6 to 10 feet for food which is primarily animal material. In summer, mostly eats aquatic insects and larvae, but also includes water boatmen, shrimplike amphipods, some snails and small fishes, and seeds of pondweeds and naiads. During winter, mostly eats shrimp, snails, and other crustaceans. REFERENCES: Erskine 1971, Johnsgard 1975b, Palmer 1976b, Terres 1980. 62 Hooded Merganser Lophodytes cucullatus L 13" RANGE: Breeds from southern Alaska to Nova Scotia, south to Oregon and Idaho, east to Maine and Massachusetts, and locally in the Mississippi Valley and southeastern United States. Winters on freshwater from British Columbia and New England south to California, Texas, Florida, and northern Mexico. STATUS: Locally common or rare. HABITAT: Inhabits wooded, clear freshwater habitats, preferably water with sandy, gravelly, or cobbled bottoms. Prefers fast-flowing water, but also uses standing water as long as it is clear, small fish and invertebrates are abundant, and nest sites are available. Easily disturbed and thus tends to avoid areas of human activity. In Wisconsin, brood habitat was described as rivers with high levels of food resources, fast current velocities (0.80 to 0.93 feet/second), and wide (40 to 60 feet) and moderately deep (1 to 2 feet) channels with cobbled bottoms and heavy surrounding cover of mixed hardwoods. SPECIAL HABITAT REQUIREMENTS: Wooded, clear water streams, rivers, swamps, ponds, and lakes with cavity trees. NEST: A cavity nester that uses almost any hole or hollow tree, at any height, as long as it is large enough for the female and her nest. The nest tree is usually within a few yards of, or standing in, water. Prefers flooded shoreline with standing trees, and with snags or stumps interspersed but will nest in other locations, including nest boxes. FOOD: Captures food during short dives in shallow, often rapidly flowing water, or at water surface. Eats small fishes (mainly rough fish), crayfish and other crustaceans, many aquatic insects such as caddis fly larvae and dragonfly nymphs, also some seeds and parts of aquatic plants. REFERENCES: Bellrose 1976, Johnsgard 1975b, Kitchen and Hunt 1969, Morse et al. 1969, Palmer 1976b, Terres 1980, Vickery in Farrand 1983a. 63 Common Merganser Mergus merganser RANGE: Breeds in North America from central and south-coastal Alaska, northern Saskatchewan, and Newfoundland south to the mountains of central California and northern New Mexico. East of the Rockies, breeds south to southern Saskatchewan, central Michigan, southern Maine, and west-central Nova Scotia. Winters from the Aleutian Islands and south- coastal Alaska east across southern Canada to Newfoundland and south to southern California and the Gulf Coast from southern Texas to central Florida. Winters as far north as open inland waters are available. STATUS: Common throughout range. HABITAT: Prefers to breed in ponds associated with upper portions of rivers in forested regions, and clear, freshwater lakes with forested shorelines. Is transcontinental in character, but essentially confined to forested regions. SPECIAL HABITAT REQUIREMENTS: Clear, forested streams, rivers, and lakes with tree cavities. NEST: Generally nests in cavities in hollow trees near water, but also in dark recesses, on the ground, or in nest boxes. Height of tree cavity and species of tree are unimportant, but the number of suitable cavities available is definitely limited. May nest beneath boulders, in root tangles along undercut streambanks, in crevices in cliffs, or in chimneys, as long as the nest is concealed from above. One pair may use 2 to 3 miles of river during nesting. FOOD: An opportunistic forager, generally feeds in fairly shallow waters from 1 1/2 to 6 feet deep. Consumes a wide variety of fishes, primarily rough and forage fish, but may be detrimental in areas specifically managed for trout or salmon production. Also eats frogs, aquatic salamanders, crayfish, shrimp, and other small crustaceans, snails and other mollusks, leeches, worms, aquatic insects and their larvae, and the roots and stems of aquatic plants. REFERENCES: Bellrose 1976, Johnsgard 1975b, Palmer 1976b, Terres 1980, Timken and Anderson 1969. 64 Red-breasted Merganser Mergus senator RANGE: Breeds from northern Alaska to central Keewatin, northern Baffin Island, Labrador and Newfoundland south to the Aleutian Islands, northern British Columbia, central Minnesota, northern New York and Nova Scotia, casually south along the Atlantic Coast to Long Island. Winters primarily along coasts and on large inland bodies of water from southern Alaska, the Great Lakes and Nova Scotia south to Baja California, southern Texas, and the Gulf Coast. STATUS: Common. HABITAT: Prefers to breed on small islands or islets with low, prostrate woody vegetation or other natural features to cover the nest, and with open shores, gravel bars, or rocks to provide roosting and preening areas for drakes and young. Although it prefers inland waters, it may be found along the coasts on shores and on marine islets. Winters mainly in estuaries and sheltered bays, less frequently on inland freshwater. SPECIAL HABITAT REQUIREMENTS: Rivers, ponds, and lakes with some overhead cover nearby for nesting. NEST: Nests on the ground under low cover, generally within 33 feet of water, preferably on islands, but also on riverbanks and lakeshores, in marshes, on rocky islets, or in bank recesses. Nests may be under low conifer boughs; under or between boulders in shallow cavities; in tall grass, heather, or bracken; or under driftwood. FOOD: Feeds primarily on fish caught during dives underwater. Also eats fish eggs, frogs, annelids, nymphs, caddis flies, amphipods, crabs, prawns, and mollusks. REFERENCES: Bellrose 1976, Clapp et al. 1982, Johnsgard 1975b, Palmer 1976b. 65 Ruddy Duck Oxyura jamaicensis 9 RANGE: Breeds in east-central Alaska and from central and northeastern British Columbia to western Ontario and south to southern California, western and southern Texas, and southwestern Louisiana, with some scattered breeding east to Nova Scotia and south to Florida. Winters from southern British Columbia, Idaho, Colorado, Kansas, the Great Lakes, and on the Atlantic Coast from Massachusetts south throughout the southern United States to Mexico. STATUS: Common. HABITAT: Inhabits permanent freshwater and alkaline prairie marshes having dense stands of cattails, bulrushes, whitetop, and reeds, and relatively stable water levels. Both large and small marshes are used for nesting, from potholes less than one acre to 1,200-acre sloughs. Commonly loafs and rests on water well out from shore. Prefers large bodies of shallow freshwater and brackish water, especially those with areas of aquatic plant growth, during migration. Winters on ice-free inland waters, or on sheltered shallow brackish or saltwater coastal waters. SPECIAL HABITAT REQUIREMENTS: Open water areas close to dense emergent vegetation with muskrat channels or natural passageways to allow movement between the nest site and open water. NEST: Usually nests over shallow water in emergent vegetation, on a floating mat of vegetation, or on a platform built up from the floor of the marsh. Occasionally nests on a muskrat house, on floating logs, or on an old coot nest. “Dump” nests are common in marshes with fluctuating water levels. FOOD: Dives for food in waters 2 to 10 feet deep, and occasionally feeds on the water surface. Consumes about 72 percent plant foods, consisting of seeds, tubers, leaves, and algae. Also eats insects, snails, and some crustaceans. REFERENCES: Bellrose 1976, Clapp et al. 1982, Cottam 1939, DeGraff et al. 1980, Johnsgard 1975b, Joyner 1969, Palmer 1976b, Siegfried 1976. 66 Black Vulture Coragyps atratus L 22" W 54" RANGE: Resident from southern Arizona and western Texas to southern Illinois, southern Indiana and New Jersey south to the Gulf Coast, southern Florida, and South America. May retreat from northern range limits in winter. STATUS: Common, but the population is declining in the southern Atlantic Coast region; range is extending slightly northward. HABITAT: Nearly ubiquitous except in heavily forested regions. It is found in the southern Great Plains, southeastern pine forests, oak- hickory forests, and intermediate oak-pine forests. NEST: Does not construct a nest. Frequently lays eggs in hollow bases of trees or stumps, rarely more then 10 to 15 feet above ground, but also on the ground, under dense or thorny vegetation, in cavities of rocks, on the floor of caves, on cliff ledges, or in abandoned buildings. FOOD: Feeds primarily on carrion from city dumps, sewers, slaughterhouses, and roadkills along highways. Also kills and eats baby herons, domestic ducks, newborn calves, baby lambs, skunks, and opposums; feeds at times on ripe and rotten fruit and vegetables. REFERENCES: Armistead in Farrand 1983a, Brown and Amadon 1968, Harrison 1979, Heintzelman 1979, Scott et al. 1977, Tate and Tate 1982, Terres 1980. 67 Turkey Vulture Cathartes aura L 25 " W 72" RANGE: Breeds from southern British Columbia, western Ontario, extreme southern Ontario and Massachusetts south throughout the remaining continental United States to South America. Winters from northern California, Arizona, Texas, Nebraska, the Ohio Valley, and Pennsylvania south to the Gulf Coast, Florida, and South America. STATUS: Common. HABITAT: Uses a wide variety of habitats, from the tropics and forested habitats dominated by mixed or deciduous trees to open plains and deserts, and from lowlands to mountains. Preens and roosts in tall snags or trees with open branches. May gather in groups of up to 70 birds to roost at night. NEST: Does not build a nest. Lays eggs on the floor of caves (preferably one which has 2 entrances), on the ground inside dense shrubs, in hollow logs or stumps, on rocky outcrops or ledges, in swamps, in hollow snags, in old hawk nests, or on the floor in abandoned buildings. The eggs are usually well-hidden from view and inaccessible to predators. FOOD: Feeds almost entirely on carrion in any state, from fresh to putrid, sighted while soaring over open fields, ridges, roads, or any type of clearing. Although partial to carrion of small mammals, amphibians, reptiles, birds, and fish, will consume carrion of large animals. Gathers quickly after the death of an animal to feed. REFERENCES: Brown and Amadon 1968, DeGraff et al. 1980, Grinnell and Miller 1944, Heintzelman 1979, Sprunt 1955, Terres 1980. 68 California Condor L 45" W 120" RANGE: Coastal ranges of California from Monterey and San Benito Counties south to Ventura County, ranging, at least casually, north to Santa Clara and San Mateo Counties, and east to the western slope of the Sierra Nevada and the Tehachapi Mountains, with breeding sites apparently confined to Los Padres National Forest in Santa Barbara, Ventura, and extreme northern Los Angeles Counties. STATUS: Endangered. All known individuals are now in captivity. HABITAT: Inhabited rugged canyons, gorges and forested mountains of southern California mainly between 985 and 8,860 feet in elevation; nested primarily between 2,000 to 4,500 feet in elevation. Spent a great deal of time roosting, preferably on dead conifers 40 to 70 feet tall, but also in live conifers and on cliffs. Needed a long, unobstructed space for taking off downhill from the roost site, which was located in areas protected from the wind, and near food, water, and nest sites. SPECIAL HABITAT REQUIREMENTS: Tall conifers or cliffs for roosting, open grasslands for feeding, cliffs for nesting, and freedom from human disturbance. NEST: Nesting areas were characterized by extremely steep, rugged terrain, with dense brush and groves of Douglas-fir surrounding high sandstone cliffs. Nested in a cavity in rock or among boulders on cliffs, with space enough to hold 2 full-grown condors, with perches available for both young and adults, protection from the elements, and space below for taking off. FOOD: Fed in open grassland because of its need for space for taking flight. Sighted food while soaring over the countryside; 95 percent of diet was carcasses of cattle, sheep, ground squirrels, deer, and horses in any state of decay. Preferred to drink water from clear pools at the tops of waterfalls, but when pressed would obtain water from any source. REFERENCES: Koford 1953, Mackenzie 1977, Wilbur 1978. 69 Osprey Pandion haliaetus L 22" W 54" RANGE: Breeds fron northwestern Alaska and northern Yukon to central Labrador and Newfoundland south locally to Baja California, central Arizona, southern Texas, the Gulf Coast, and southern Florida. Winters from central California, southern Texas, the Gulf Coast and Florida south to South America. STATUS: Locally common to uncommon; population declining due to destruction of habitat, pesticides, human disturbance, and reduction of food resources. HABITAT: Nearly cosmopolitan distribution, occurring on every continent except Antarctica. Occupies a wide range of habitats in association with water, primarily near lakes, rivers, and along coastal waters with adequate supplies of fish. SPECIAL HABITAT REQUIREMENTS: Elevated nest sites near water with rich fish resources. NEST: Nests in loose colonies or singly, and uses a wide variety of structures to support large stick nests, which may be 60 feet or more above ground. Prefers a snag in or near water, with a broken top or side limbs able to support the nest. Prefers tall snags that provide good visibility and security. Also nests on pilings, utility poles, duck blinds, buildings, steel towers for transmission lines, windmills, channel markers, fences, a wide variety of living, partially dead, or dead trees, wooden platforms in marshes, on cliffs, and sometimes on the ground. Nest site may be used by the same pair year after year. FOOD: Feeds almost exclusively on fish; flies 50 to 100 feet above (preferably shallow) water, then hovers and plunges into the water to catch fish. Also eats frogs, snakes, ducks, crows, night-herons, and small mammals. REFERENCES: DeGraff et al. 1980, Heintzelman 1979, Sprunt 1955, Zarn 1974a. 70 American Swallow-tailed Kite Elanoides forficatus RANGE: Breeds locally from South Carolina south to Florida, and west to Louisiana; occasionally to Great Lakes and New England and also Central and South America. Winters in South America. STATUS: Locally common. HABITAT: Inhabits open river bottom forests with adjacent semi-prairie land, freshwater marshes bordering large lakes, lowland cypress swamps, and pine glades. SPECIAL HABITAT REQUIREMENTS: Very tall living trees for nesting. NEST: Nests in the very tops of tall, slender living trees, usually 60 to 100 feet above ground, but up to 200 feet high. In Florida, usually nests in pines or in black mangroves. Selects trees in open, thinly wooded areas, or along the edge of trails or openings so the birds can approach the nest unimpeded. Other kites are tolerated near the nest, but not other hawks or eagles. FOOD: Feeds entirely on the wing, primarily on flying insects, but also sweeps low over fields, forest canopies, and prairies to catch grasshoppers, crickets, cicadas, small snakes, lizards, and frogs. Also snatches young birds and eggs, and drinks while skimming the surface of a lake or pond. REFERENCES: Brown and Amadon 1968, Heintzelman 1979, Oberholser 1974a, Terres 1980. 71 Black-shouldered Kite Elanus caeruleus (formerly White-tailed Kite) L14” RANGE: Resident locally from northwestern Oregon south (west of the deserts) to Baja California, and from southern Oklahoma, western Louisiana, east-central and southeastern Texas south to South America. Strays to adjacent states, also to Florida, and in the Mississippi Valley north to Missouri and southern Illinois. STATUS: Rare to locally fairly common; once reduced in numbers almost to the point of extinction in the United States. Year-round irrigation of agricultural land has improved habitat conditions in recent years. HABITAT: Inhabits open country around freshwater marshes, moist meadows, alfalfa fields, and cultivated bottomlands, with scattered clumps of trees. In the western Sierra Nevada in California, found below 1,000 feet in blue oak-savannah, digger pine-oak, and riparian deciduous types. SPECIAL HABITAT REQUIREMENTS: Trees with dense canopies for nesting near a permanent water source and an abundant population of voles (Microtus spp.). NEST: Nests in oak, willow, eucalyptus, cottonwood, or other hardwood trees, from 18 to 59 feet above ground, usually near a marsh, streambank, or canal, and areas where voles are abundant. With good vole populations, breeding pairs need a minimum of 20 acres around the nest site for hunting. FOOD: Searches for food by flying and hovering at less than 100 feet above ground. Feeds primarily on voles, but also eats other small mammals, small snakes, lizards, frogs, and large insects such as grasshoppers, beetles, and crickets. REFERENCES: Brown and Amadon 1968, Heintzelman 1979, Oberholser 1974a, Terres 1980, Verner and Boss 1980. 72 Snail Kite Rostrhamus sociabilis (formerly Everglade Kite) LI 5” RANGE: Resident in southern Florida, primarily at Lake Okeechobee and Loxahatchee National Wildlife Refuge, and locally throughout the Everglades basin and the upper St. John’s River. STATUS: Endangered due to habitat destruction, hunting, and drought. HABITAT: Highly specialized; inhabits permanent freshwater marshes with broad expanses of cattails, sawgrass, or other tall, emergent grasses, and with scattered clumps of bushes or small trees with a low and distant horizon for visibility. SPECIAL HABITAT REQUIREMENTS: A permanent water source supporting adequate quantities of the apple snail. NEST: Often nests in loose colonies, sometimes in or adjacent to colonies of herons, egrets, and anhingas. Males construct nests up to 8 feet above water in cattails, reeds, or bulrushes, or in willows or other shrubs or trees growing in water. Also nests in areas with tree islands dominated by dead trees and shrubs, or on artificial nest platforms. FOOD: Feeds exclusively on the apple snail, which it sights while flying slowly low over water, or while perching on an old stake, a mound of aquatic debris, or a cattail clump. REFERENCES: Brown and Amadon 1968, Heintzelman 1979, Mackenzie 1977, Sprunt 1955, Stieglitz and Thompson 1967, Terres 1980. 73 Mississippi Kite Ictinia mississippiensis L12” RANGE: Breeds from central Arizona to north-central Kansas, southern Illinois, western Kentucky, the northern portions of the Gulf States and South Carolina south to central and southeastern New Mexico across the Gulf Coast and north-central Florida. Expanding range along its northern border. Winters in South America. STATUS: Common in the southern Great Plains, uncommon in the Southeast. HABITAT: Inhabits forests, open woodlands, and prairies. Found on the prairies of Kansas and in baldcypress swamps and pinelands in the Gulf States, scrub oak country in Oklahoma, and mesquite-sand sagebrush rangeland in Texas. NEST: In the East, nests in riparian habitats, or in large pines, oaks, and sweetgums of large woods. In the Great Plains, nests in shelterbelts, farm woodlots, lawn trees in towns, or any small grove of trees. Also nests in scrub oaks and mesquite. Depending on the tree, places nest from 10 to 135 feet above ground. In Arizona, nests in cottonwoods taller than 50 feet in open groves or in scattered clumps surrounded by dense riparian scrubland of saltcedar and velvet mesquite 6.5 to 33 feet tall. FOOD: Primarily consumes insects caught in the air, including grasshoppers, locusts, cicadas, katydids, large beetles, and dragonflies; also eats small snakes, lizards, and frogs. REFERENCES: Brown and Amadon 1968, Glinski and Ohmart 1983, Heintzelman 1979, Kaufman in Farrand 1983a, Oberholser 1974a, Parker and Ogden 1979, Sprunt 1955, Terres 1980. 74 Bald Eagle Haliaeetus leucocephalus im. RANGE: Breeds from central Alaska and northern Yukon across Canada to Labrador and Newfoundland, south locally to the Aleutian Islands, southern Alaska, central Arizona, southwestern and central New Mexico, Baja California, and the Gulf Coast; very locally distributed in the interior of North America. Winters generally throughout the breeding range, but most frequently from southern Alaska and southern Canada southward. STATUS: Endangered and threatened in parts of the lower 48 states. HABITAT: Closely associated with lakes and large rivers in open areas, forests and mountains, and along seacoasts. In Alaska and Canada, where human disturbance is slight, habitat is composed of a narrow strip of land along lakeshores and rivers that provides trees for nesting, fishing, and loafing. Needs large trees adjacent to water, preferably snags, but also live trees or boulders that provide good visibility, for perching. Winters in coastal habitats and inland where ice-free waters allow access to fish. SPECIAL HABITAT REQUIREMENTS: Large bodies of water containing abundant fish resources, large trees for nesting, perching, and roosting, and freedom from human disturbance. NEST: Prefers to build a large, heavy nest 10 to 150 feet above ground in very tall living trees, usually close to water. If suitable trees are not available, nests are built on rocky cliffs or on the ground. Shows strong attachment to the nest site, and characteristically adds new material to the nest each year. FOOD: Feeds primarily on fish it catches or takes from an osprey. Will feed on waterfowl and other birds, carrion, small- to medium-sized mammals, and turtles. Inland, subsists mainly on dead waterfowl during winter. REFERENCES: DeGraff et al. 1980, Evans 1982, Fielder 1982, Grubb and Kennedy 1982, Heintzelman 1979, Mackenzie 1977, Sprunt 1955. 75 Northern Harrier Circus cyaneus (formerly Marsh Hawk) RANGE: Breeds from northern Alaska to southern Quebec and Newfoundland south to Baja California, southern Arizona, southern and eastern Texas, southern Illinois, and southeastern Virginia. Winters from Alaska (casually) and southern British Columbia east to South Dakota, southern Ontario, and Massachusetts south through the United States to South America. STATUS: Common; populations are increasing slightly in the Southwest, and declining in the Northeast and Midwest. HABITAT: Typically inhabits sloughs, wet meadows, fresh or salt marshes, swamps, prairies and plains. Generally roosts on the ground or perches on very low objects such as fence posts or tree stumps. During the non-breeding season, inhabits areas far removed from nesting habitat. Roosts in undisturbed fields or marshes in winter. SPECIAL HABITAT REQUIREMENTS: Open country with herbaceous or low woody vegetation for concealing nests. NEST: Nests singly or sometimes semi-colonially, on the ground in a variety of sites, but usually near or above water. Nests in tall grass in open fields, in swamps with low shrubs and clearings, sometimes built up over water on a stick foundation, sedge tussock, or willow clump, or on a knoll of dry ground. FOOD: Hunts for food, primarily on the wing, over fields, marshes, and meadows, taking a wide variety of prey including mammals, birds, amphibians, reptiles, insects, and fishes. Mostly eats small mammals. REFERENCES: DeGraff et al. 1980, Evans 1982, Heintzelman 1979, Low and Mansell 1983, McAtee 1935, Sprunt 1955, Tate and Tate 1982, Terres 1980. 76 Sharp-shinned Hawk Accipiter striatus RANGE: Breeds from western and central Alaska and northern Yukon to southern Labrador and Newfoundland, south to central California, southern Texas, the northern parts of the Gulf States, and South Carolina. Winters from southern Alaska, the southernmost portions of the Canadian Provinces south through the United States to Panama. STATUS: Fairly common; the population appears to be recovering from earlier declines that occurred until the early 1970’s in the eastern United States. HABITAT: Primarily inhabits coniferous and mixed conifer-birch-aspen forests of the Canadian and Transition life zones northward to the Arctic tree line. Less commonly inhabits other woodland types except in mountainous areas. During migration and in winter it may occur in almost any type of habitat containing trees or shrubs. SPECIAL HABITAT REQUIREMENTS: Dense coniferous-deciduous forest. NEST: Usually nests in trees with dense foliage, primarily conifers, from 6 to 90 feet, typically 30 to 35 feet above ground and below a well- developed canopy. Nests may be in small groves of conifers surrounded by deciduous trees. Generally constructs a new nest each year in the immediate area of the previous year’s nest. FOOD: Feeds primarily on birds sighted while flying over forest floor, meadows, and brushy pastures. Sparrow-sized birds are taken most often, but occasionally attacks birds larger than itself. Also eats a few small mammals, reptiles, and insects. REFERENCES: DeGraff et al. 1980, Evans 1982, Heintzelman 1979, Jones 1979, Platt 1976, Reynolds et al. 1982, Tate and Tate 1982. 77 Cooper’s Hawk Accipiter cooperii L 15Vi” W 28" RANGE: Breeds from southern British Columbia and central Alberta to southern Quebec and Maine south to Baja California, Mexico, Louisiana, central Mississippi, Alabama, and Florida. Winters from Washington, Colorado, and southern Minnesota to New England south through the southern United States, to Costa Rica. STATUS: Uncommon. HABITAT: Inhabits various types of mixed and deciduous forests and open woodlands including small woodlots, riparian woodlands in dry country, open arid pinyon woodlands, and forested mountainous regions. May use almost any type of habitat containing trees or shrubs during winter and in migration. SPECIAL HABITAT REQUIREMENTS: Mature coniferous or deciduous woodlands. NEST: Usually nests in deciduous or coniferous trees near the edge of a wooded area, with large open fields and water nearby. Places nest from 20 to 60 feet above ground (usually 35 to 45 feet). Occasionally uses old crow nests. FOOD: Hunts from inconspicuous perches, and catches its prey, primarily birds, by surprise. Consumes medium-sized birds such as thrushes, jays, starlings, and quail primarily but also takes smaller birds and larger birds up to the size of ruffed grouse. Also eats chipmunks, red squirrels, rabbits, other small mammals, amphibians, and insects. REFERENCES: DeGraff et al. 1980, Evans 1982, Heintzelman 1979, Jones 1979, Reynolds et al. 1982. 78 Northern Goshawk Accipiter gentilis (formerly Goshawk) 42" RANGE: Breeds from western and central Alaska and northern Yukon to Labrador and Newfoundland, south to southern Alaska, central California, southern New Mexico, western South Dakota, northern Minnesota, and northwestern Connecticut, and in the northern Appalachian Mountains. Winters throughout the breeding range may extend as far south as the Gulf States during periodic invasions related to food shortage. STATUS: Uncommon to rare but increasing; range is expanding southward in Appalachians. HABITAT: Inhabits mixed hardwood and coniferous forests in temperate and boreal regions, from sea level to tree line. Prefers woodlands with intermediate canopy coverage interspersed with fields or wetlands, especially in remote areas. SPECIAL HABITAT REQUIREMENTS: Extensive mixed woodlands with large trees for nesting. NEST: Prefers to nest in large hardwood trees 30 to 40 feet above ground, where clear, level access is afforded by a stream or other opening. Frequently selects birch, maple, aspen, and beech for nesting trees; occasionally selects juniper, pine, spruce, and fir. Usually builds a new nest each year, but may build on top of an old hawk nest. FOOD: Hunts for prey in dense woodlands, clearings, and open fields. In one study, its diet consisted of 54 percent birds, 37 percent mammals, and 9 percent insects. Eats grouse, quail, pheasants, small hawks, owls, crows, gulls, ducks, doves, thrushes, rabbits, squirrels, chipmunks, mice, woodchucks, muskrats, weasels, shrews, grasshoppers, and caterpillars. REFERENCES: Cramp and Simmons 1980, DeGraff et al. 1980, Evans in Farrand 1983a, Heintzelman 1979, Jones 1979, McAtee 1935, Shuster 1980, Terres 1980. 79 Common Black-Hawk Buteogallus anthracinus (formerly Black Hawk) L 20" W 48" RANGE: Resident in central Arizona, southwestern Utah, southern New Mexico, and western Texas, south through Central America to Colombia (northernmost populations move southward during winter). STATUS: Rare; threatened in Arizona and New Mexico. HABITAT: An obligate of riparian areas. Optimum habitat consists of a flowing stream bordered by mature riparian forests. Also inhabits broad alluvial valleys, narrow rocky canyons, or marshes near the coast. SPECIAL HABITAT REQUIREMENTS: Mature, relatively undisturbed habitat with a permanent water source and tall (75 to 100 feet) trees for nesting. NEST: Nests in trees from 15 to 100 feet above ground, preferably within a grove of trees rather than in a lone tree. Builds nests in cottonwood, sycamore, alder, mesquite, willow, velvet ash, ponderosa pine and Douglas-fir. May use same nest for successive years. FOOD: Prefers to fish in streams of low to moderate gradient, less than one foot deep with scattered boulders and some low or fallen branches. Usually locates prey while flying but also hunts from a perch. Eats a varied assortment of prey, including beach and land crabs, frogs, fishes, crayfish, reptiles, small mammals, birds, and insects. REFERENCES: Heintzelman 1979, Oberholser 1974a, Schnell 1979. 80 Harris’ Hawk Parabuteo unicinctus RANGE: Resident in southern Kansas, and from southeastern California (recently reintroduced), southern Arizona, southern New Mexico, and central Texas south to South America. STATUS: Fairly common. HABITAT: Inhabits arid desert scrub of mesquite, palo verde and large cacti in the Southwest, river woodlands, and brushy flatlands. Infrequently found in yucca, cactus, creosotebush deserts, and oak- juniper habitats. Commonly perches on tops of telephone poles, trees, and bushes; on large cacti; and in snags. SPECIAL HABITAT REQUIREMENTS: Thorn-scrub habitats. NEST: Nests from 5 to 30 feet up in cactus, mesquite, hackberry, yucca, Spanish-bayonet, paloverde, ironwood, cottonwood, ebony, and other trees. May nest in pairs or in trios with an extra male that also brings prey to the nest. FOOD: Consumes a diet comprising 57 percent mammals, 35 percent birds, and 7 percent lizards. Feeds on rabbits, wood rats, mice, night- herons, teal, flickers, and other birds and mammals. REFERENCES: Harrison 1979, Heintzelman 1979, Oberholser 1974a, Sprunt 1955, Terres 1980, Terrill in Farrand 1983a. 81 Gray Hawk Buteo nitidus L 15" W 35" RANGE: Breeds in southern Arizona and southern Texas (the northwestern extreme of its breeding range) south to South America. Northernmost populations usually migrate south in nonbreeding season. STATUS: Rare. HABITAT: Is found primarily in riparian willow, cottonwood, and sycamore groves in the San Pedro and Santa Cruz river drainages of Arizona. Inhabits mature woodlands of river valleys and nearby semiarid mesquite and scrub grasslands. SPECIAL HABITAT REQUIREMENTS: Stands of cottonwood and willow near rivers for nesting. NEST: The small nest is placed from 20 to 40 feet up in cottonwood, hackberry, or mesquite along streams or rivers. FOOD: Prefers to feed on lizards and small snakes, and frequently eats beetles and large grasshoppers. Also catches and eats rabbits, squirrels, mice, quail, young doves, and fishes. REFERENCES: Heintzelman 1979, Oberholser 1974a, Terres 1980, Terrill in Farrand 1983a. 82 Red-shouldered Hawk Buteo lineatus L 16" W 40" RANGE: Breeds from northern California south, west of the Sierra Nevada divide, to Baja California; and from eastern Nebraska, central Minnesota, southern Ontario, and southern New Brunswick south to Mexico. Winters primarily from eastern Kansas and central Missouri to southern New England southward, but also sporadically throughout breeding range. STATUS: Common, but population is unstable. HABITAT: Inhabits moist, well-drained woodlands, wooded river swamps, bottomlands, and wooded margins of marshes, often close to cultivated fields. Seems to prefer mature forests and is usually more common in lowland areas than in mountainous regions. SPECIAL HABITAT REQUIREMENTS: Riparian deciduous woodlands with tall trees for nesting. NEST: Nests 20 to 60 feet above ground in tall trees. Usually builds nest 35-45 feet above ground on a main fork and close to the tree trunk. Has built nests in oak, pine, baldcypress, mangrove, cottonwood, birch, beech, sycamore, yellow-poplar, ash, sweetgum and maple. Occasionally uses an abandoned hawk, crow, or squirrel nest as a foundation for a new nest; often uses the same nest site year after year. FOOD: Perches on a fence post, tree, or telephone pole and overlooks a meadow, marsh, open field, or forest to sight prey. Feeds primarily on small mammals but also takes rabbits, squirrels, small birds, frogs, small snakes, toads, lizards, fishes, and large insects. REFERENCES: Bednarz and Dinsmore 1982, DeGraff et al. 1980, Forbush and May 1955, Heintzelman 1979, McAtee 1935, Portnoy and Dodge 1979, Sprunt 1955, Stewart 1949, Tate and Tate 1982. 83 Broad-winged Hawk Buteo platypterus A RANGE: Breeds in central Alberta and central Saskatchewan, and from central Manitoba to New Brunswick and Nova Scotia south to eastern Texas, the Gulf Coast and Florida. Winters in southern Florida and from Mexico to South America. STATUS: Common throughout range. HABITAT: Inhabits continuous dry woodlands of oaks, beeches, maples, and mixed coniferous-hardwoods around lakes, streams, and swamps. In migration when conditions are favorable, forms large flocks, or “kettles,” soars to the top of thermals, and then glides to another, thus saving energy during the long flight to its wintering area. NEST: Normally nests near water in a variety of tree species, from 25 to 90 feet, but as low as 3 to 10 feet, above ground. Nest site preference is probably related to life form of the tree species and characteristics of the site rather than to prevalence of a particular tree species. Black and yellow birch are commonly selected for nesting in New England. Sometimes uses old crow, hawk, or squirrel nests. FOOD: Hunts from perch in deep, shady woodlands or while flying over treetops or open meadows. Feeds largely on small mammals such as mice, moles, and shrews, occasionally red squirrels and chipmunks; also eats snakes, frogs, lizards, large larvae of night-flying moths, caterpillars, grasshoppers, beetles, crickets, fiddler crabs, crayfish, sometimes small fish, and some small birds. REFERENCES: DeGraff et al. 1980, Evans in Farrand 1983a, Forbush and May 1955, Heintzelman 1979, Matray 1974, Sprunt 1955, Terres 1980. 84 Short-tailed Hawk Buteo brachyurus dark phase L 14“ W 35" RANGE: Resident locally in peninsular Florida, from St. Marks and San Mateo south to Lake Okeechobee. Winters mostly south of Lake Okeechobee and in Central and South America. STATUS: Rare. HABITAT: A tropical species that barely reaches the United States in Florida, this buteo is found primarily in mixed woodland-savannah habitats, but also in mangrove and baldcypress swamps adjacent to forests, and along streams and the borders of lakes. Often perches in tall trees. NEST: Nests in the topmost branches of tupelos, pines, magnolias, or baldcypresses, from 40 to 100 feet above ground or in the top of mangroves. FOOD: Feeds on a great variety of birds, but primarily meadowlarks and red-winged blackbirds. Also feeds on some small mammals, reptiles, and amphibians. REFERENCES: Heintzelman 1979, Ogden 1974, Ogden in Farrand 1983a, Sprunt 1955, Terres 1980. 85 Swainson’s Hawk Buteo swainsoni L 18" W49" dark phase light phase RANGE: Breeds locally in east-central Alaska, Yukon, and Mackenzie, and from central Alberta and central Saskatchewan to western Illinois south to southern California, central and southern Texas, and western Missouri. Winters primarily on the pampas of southern South America, casually north to the southwestern United States and southeastern Florida. STATUS: Common; population has decreased in the southern Great Plains. HABITAT: Inhabits prairies, plains, deserts, large mountain valleys, savannahs, open pine-oak woodlands, and cultivated lands with scattered trees. NEST: Nests in isolated trees, in shrubs and trees along wetlands and drainages, in windbreaks in fields and around farmsteads, in giant cactus, or on the crossbars of telephone poles. Occasionally nests on the ground, on low cliffs, on rocky pinnacles, or on cutbanks. May build nest up to 100 feet above ground in cottonwoods, or lower in willows or other shrubs. May repair and use same nest year after year; sometimes builds on old black-billed magpie nests. FOOD: Hunts primarily from perches such as fence posts or low trees and from a vantage point on the ground. Diet consists of small mammals, birds, fishes, salamanders, frogs, snakes, and insects. REFERENCES: Dunkle 1977, Evans in Farrand 1983a, Heintzelman 1979, Sprunt 1955, Tate and Tate 1982, Terres 1980. 86 White-tailed Hawk Buteo albicaudatus L 21" W 48' RANGE: Resident from central and southeastern Texas south to South America. STATUS: Uncommon. HABITAT: Inhabits saltgrass flats of coastal grasslands, and open grassy ranges and chaparral country with scattered mesquite, yucca, and large cacti farther inland. Perches on bushes, trees, utility wires, or on the ground. NEST: Nests on prairies, in brush, or in fringes of timber in sizeable bushes or small trees, 5 to 15 feet above ground. Prefers to nest on a ridge with a view all around. Nest plants include large cactus, yucca, and scrub oaks. May use the same nest in successive years. FOOD: Feeds extensively upon rabbits; also eats cotton rats, lizards, snakes, frogs, grasshoppers, beetles, cicadas, and other insects. REFERENCES: Heintzelman 1979, Oberholser 1974a, Terres 1980. 87 Zone-tailed Hawk Buteo albonotatus L19" W 47' RANGE: Breeds from central Arizona, southern New Mexico, and western Texas south to South America. Very rare north of Mexico in winter. STATUS: Locally fairly common. HABITAT: Inhabits deep, rough, and rocky wooded canyons and tree- lined rivers along middle slopes of desert mountains, especially in open deciduous or pine-oak woodland. SPECIAL HABITAT REQUIREMENTS: Large trees for nesting. NEST: Builds a bulky nest in large trees, usually cottonwoods along streams and rivers, 25 to 100 feet above ground in leafy top. Rarely nests in mesquites. FOOD: Eats chipmunks, quail and small birds, lizards, small fishes, and frogs (little is known about diet). REFERENCES: Heintzelman 1979, Oberholser 1974a, Terres 1980. 88 Red-tailed Hawk Buteo jamaicensis dark phase f- - L 18" W 48" RANGE: Breeds from western and central Alaska and central Yukon to New Brunswick and Nova Scotia south to Central America. Winters from southern Canada throughout the remainder of the breeding range. STATUS: Common, but population is declining. HABITAT: Inhabits a wide variety of different habitats throughout its range, preferring mixed country of open pasture, fields, meadows, or swampy areas interspersed with coniferous or deciduous woods. Inhabits deserts and plains with scattered trees and open mountain forests, generally avoiding dense, unbroken woodlands and tundra. NEST: Usually nests in a tall tree in or at the edge of a woodland, or in an isolated tree in an open area. Frequently selects the largest and tallest tree (of a wide variety of species) available. Constructs nest next to the trunk in a crotch from 35 to 90 feet above ground. In treeless areas, nests on rocky cliffs, shrubs, or cactus. FOOD: Frequently hunts for prey while perching in snags, live trees, or on poles in rather open areas or at forest edges. Also locates prey while soaring. Primarily eats small mammals; also eats birds, reptiles, and some insects. REFERENCES: Austin 1964, Bednarz and Dinsmore 1982, DeGraff et al. 1980, Evans in Farrand 1983a, Fitch et al. 1946, Forbush and May 1955, Heintzelman 1979, Terres 1980. 89 Ferruginous Buteo regalis Hawk RANGE: Breeds from eastern Washington, southern Alberta, and southern Saskatchewan south to eastern Oregon, Nevada, northern and southeastern Arizona, northern New Mexico, north-central Texas, western Oklahoma, and Kansas. Winters primarily from the central and southern parts of breeding range south to Mexico. STATUS: Common; population is stable or declining slowly. HABITAT: Inhabits the semiarid western plains and arid intermountain regions; prefers relatively unbroken terrain, with scattered trees, rock outcrops, or tall trees along creek bottoms available for nesting sites. Generally winters on the southern plains. SPECIAL HABITAT REQUIREMENTS: Open country with elevated nesting sites. NEST: Prefers tall trees for nesting; will use a wide variety of sites, including ground nests on riverbed mounds, cutbanks, low hills, buttes and small cliffs, in short trees in open country, powerline structures, and haystacks. Tree nests are usually in the upper canopy, from 6 to 55 feet above ground. Nests are often used year after year. FOOD: Hunts from a perch, while soaring, during low, rapid flight over open country, or while systematically searching and hovering at 40 to 60 feet. One study found its diet to be 70 percent mammals, 27 percent birds, and 3 percent reptiles. Feeds primarily on rabbits, ground squirrels, and prairie dogs; also takes mice, rats, gophers, birds, snakes, locusts, and crickets. REFERENCES: Blair and Schitoskey 1982, Evans 1982, Heintzelman 1979, Snow 1974a, Sprunt 1955, Tate and Tate 1982, Weston 1969, Woffinden and Murphy 1983. 90 Rough-legged Hawk Buteo lagopus RANGE: Breeds from western and northern Alaska, northern Yukon, and northern Labrador south to northern and southeastern Mackenzie, east to northern Quebec and Newfoundland; also from Kodiak Island and Umnak in the eastern Aleutian Islands and the Arctic Islands north to Prince Patrick, Victoria, Bylot, and southwestern Baffin Islands. Winters from south-central Alaska (casually) and southern Canada south to southern California, southern Arizona east to southern Texas, Missouri, Tennessee, and Virginia, casually to eastern Texas and the Gulf Coast. Concentrates in areas of high prey density during winter. STATUS: Most common hawk of the American arctic. HABITAT: Inhabits open tundra and mountainsides; does not inhabit forests unless there is much open ground. Essentially an open country dweller that occupies a large range in its seasonal wanderings. In winter, prefers conifer groves for roosting and open, treeless areas for hunting. NEST: Nests primarily on cliffs along river bluffs, but is flexibile in selecting nesting substrate. Locates nests usually under overhangs on rocky cliffs, outcroppings, and ledges; occasionally nests in stunted trees or on the ground. Often returns to the same nest for many years. FOOD: Hunts for food in wet meadows, bogs, and riparian areas. Generally seeks prey from the air rather than from a perch. Microtine rodents such as brown lemming, collared lemming, tundra vole, Alaska vole, red-backed vole, and other small mammals comprise the bulk of the diet. Shifts to other prey when rodents become scarce. Also consumes young ptarmigan, arctic ground squirrels, and sometimes small rabbits. During the breeding season, may consume up to 20 percent of diet as birds; during winter, consumes mammalian prey almost exclusively. REFERENCES: Heintzelman 1979, Sprunt, 1955, Terres 1980, White and Cade 1971, Zarn 1975. 91 Golden Eagle Aquila chrysaetos RANGE: Breeds from northern and western Alaska east to Labrador, south to southern Alaska, Baja California, western and central Texas, western Oklahoma, and western Kansas; in eastern North America to New York and New England. Winters from south-central Alaska and the southern portions of the Canadian provinces south throughout the breeding range, rarely to coastal South Carolina. STATUS: Fairly common in the West, rare in the East. HABITAT: Inhabits open country, from barren areas to open coniferous forests, primarily in hilly and mountainous regions, but also in rugged deserts, on the plains, and in tundra. Prefers large trees with large horizontal branches and cliffs for roosting and perching. In the West, often moves down from the mountains onto the plains and valleys for winter. SPECIAL HABITAT REQUIREMENTS: Elevated nest sites, especially cliffs, that are isolated from human disturbance and are close to hunting areas. NEST: Usually nests on cliff ledges, preferably overlooking grasslands, but also nests in trees or on the ground. In the western mountains, nests at elevations of 4,000 to 10,000 feet above sea level. May use the same nest year after year, or pairs may use alternate nests in successive years. FOOD: An opportunist; hunts for a variety of prey by soaring over open country or by sighting prey from perch. Feeds primarily on mammals (mainly lagomorphs), but also marmots, prairie dogs, ground squirrels, weasels, woodrats, skunks, and mice, rarely on larger mammals. Also eats grouse, pheasants, owls, hawks, rock doves, magpies, and other birds, as well as rattlesnakes and some carrion. REFERENCES: DeGraff et al. 1980, Heintzelman 1979, Jollie 1943, McGahan 1968, Snow 1973, Terres 1980. 92 American Kestrel Falco sparverius RANGE: Breeds from western and central Alaska and southern Yukon to northern Ontario, southern Quebec, and southern Newfoundland south to Mexico. Winters from south-central Alaska, southern British Columbia, and northern United States south throughout the breeding range to Panama. STATUS: Common. HABITAT: Widely distributed in habitats that include deserts, forest openings, marshes, grasslands, agricultural and suburban areas, towns, and cities. Frequently perches on fence posts, utility poles and wires, and in trees. Occupies the same types of habitats during winter as during the breeding season. SPECIAL HABITAT REQUIREMENTS: Open country with low vegetation, cavities in trees with dbh greater than 12 inches, and elevated perches from which to sight prey. NEST: Prefers to nest in natural tree cavities with tight-fitting entrances, or in cavities excavated by flickers. If these are unavailable, nests in a variety of sites including niches in rocky cliffs, under eaves of buildings, in old magpie nests, in cavities in cacti, in unused chimneys, or in nest boxes. Nest sites are usually along roadways, streams, ponds, or forest edges, from 4 to 65 feet above ground, though typically from 10 to 35 feet. FOOD: Hunts from a perch or while hovering over areas with short, open vegetation. Primarily eats insects such as grasshoppers, crickets, and beetles in summer, but also takes mice and other small mammals, birds, lizards, toads, frogs, and small snakes; rarely takes spiders or worms. REFERENCES: Balgooyen 1976, DeGraff et al. 1980, Evans in Farrand 1983a, Heintzelman 1979, McAtee 1935, Smith et al. 1972, Thomas et al. 1979. 93 Merlin Falco columbarius L 12" W 23" RANGE: Breeds from northwestern Alaska and northern Yukon to Labrador and Newfoundland, south to southern Alaska, eastern Oregon, northern Minnesota, southern Quebec, New Brunswick, and Nova Scotia. Winters west of the Rockies from south-central Alaska, southern British Columbia, Wyoming, and Colorado southward, locally across southern Canada, and in the eastern United States from Maryland, the Gulf Coast, and southern Texas to South America. STATUS: Uncommon. HABITAT: Inhabits open areas such as forest edges, bogs, and lakes in boreal and moist Pacific Coastal forests, and prairie-parkland of the northern Great Plains. Some remain in prairie habitat even in winter; others will use almost any habitat type encountered in its winter range. NEST: Generally nests in trees from 5 to 60 feet above ground, often in old stick nests of crows, ravens, magpies, or other raptors, in or near open areas, and generally near water. Occasionally nests on the ground, on bare ledge of a cliff, or in cavities in trees. Prairie birds prefer to nest in isolated groves of trees near water, and in wooded areas along rivers, generally in coniferous trees. FOOD: Sights prey from an inconspicuous perch or during flight. Preys almost entirely on small to medium-sized birds; also takes large insects, scorpions, spiders, crayfish, toads, small snakes, bats, and small mammals. REFERENCES: Evans 1982, Fox 1964, Heintzelman 1979, McAtee 1935, Sprunt 1955, Trimble 1975. 94 Peregrine Falcon RANGE: Breeds from northern Alaska, Banks, Victoria, southern Melville, Somerset and northern Baffin Islands, and Labrador south to Baja California, southern Arizona, New Mexico, western and central Texas, and Colorado; recently re-introduced and re-established as a breeding bird in parts of the northeastern United States. Winters from southern Alaska, the Queen Charlotte Islands, coastal British Columbia, the central and southern United States, and New Brunswick south to South America. STATUS: Rare and endangered; catastrophic decline primarily due to organochlorine pesticides. HABITAT: Usually inhabits open country from tundra and seacoasts, to high mountains and more open forested regions, preferably where there are rocky cliffs with ledges overlooking rivers, lakes, or other water and an abundance of birds. Sometimes breeds in cities. SPECIAL HABITAT REQUIREMENTS: Cliffs or other nesting habitat near water, and an abundance of prey. NEST: Prefers to nest in a shallow depression scraped in gravel and debris on a high cliff ledge, pothole, or small cave that provides sanctuary from disturbance. Bluffs, slopes, pinnacles, cutbanks, and seastacks are also used as nest sites in the far north. Other nest sites include old stick nests of ravens and hawks, ledges of tall buildings, and historically, holes and stubs of large trees. Tends to return to the same nesting cliff. FOOD: Pursues prey, primarily birds, after sighting from perch or while soaring. Small- to medium-sized birds are usually captured in flight; birds too large to be carried are knocked to the ground. Feeds on a wide variety of birds; occasionally takes mammals, some insects, and fishes. REFERENCES: Cade 1960, DeGraff et al. 1980, Evans 1982, Heintzelman 1979, Hickey 1942, Hickey and Anderson 1969, Terres 1980, White and Cade 1971. 95 Prairie Falcon Falco mexicanus RANGE: Breeds from southeastern British Columbia, southern Alberta, southern Saskatchewan, and northern North Dakota south to Baja California, New Mexico, and northern Texas. Winters from the breeding range in southern Canada south to Mexico. STATUS: Locally common. HABITAT: Inhabits prairies, deserts, riverine escarpments, canyons, foothills, and mountains in relatively arid western regions. Occupies open, treeless terrain that accommodates its low-level style of hunting. Wintering birds are found away from the breeding areas in intermontane valleys and on the Great Plains. SPECIAL HABITAT REQUIREMENTS: Suitable nesting sites on cliffs in open country free of human disturbance. NEST: Nests on cliffs, from low rock outcrops of 30 feet to vertical cliffs 400 feet high. Prefers cliffs with a sheltered ledge with loose debris or gravel for a nest scrape, overlooking treeless country for hunting. Also nests in larger caves in cliffs and vertical or columnar cracks with lodged material. Sometimes uses old nests of ravens, hawks, or eagles. FOOD: Typically hunts from perches or in low, rapid, searching flight, usually capturing prey on or near the ground. Feeds on a variety of prey including ducks, prairie chickens, quail, pigeons, doves, sparrows and other small birds, prairie dogs, mice, ground squirrels, young rabbits, grasshoppers, and lizards. REFERENCES: Enderson 1964, Evans 1982, Heintzelman 1979, McAtee 1935, Snow 1974b, Sprunt 1955, Terres 1980. 96 Plain Chachalaca Ortalis vetula (formerly Chachalaca) L RANGE: Resident from the lower Rio Grande Valley to Costa Rica. Introduced on Sapelo, Blackbeard, and Little St. Simons Islands, Georgia. STATUS: Locally common to uncommon; largely restricted to parks and refuges because of clearing of habitat for residential and agricultural purposes. HABITAT: Prefers wooded stream beds with thick growths of ebony blackbead, hackberry, mesquite, and thick, shrubbery undergrowth. Rarely found far from water; also inhabits thick growths of sugarberry, Texas lignumvitae, huisache, cedar elm, and willow. Has adapted well to living in relatively small (1 to 5 acres) tracts of dense woodland vegetation. SPECIAL HABITAT REQUIREMENTS: Dense brushland. NEST: Constructs a small, flimsy nest or uses old nests of birds such as the yellow-billed cuckoo, groove-billed ani, or curve-billed thrasher. Nests in trees, or in vines supported by trees, from 3 to 33 feet above ground. Most commonly uses cedar elm, huisache, sugarberry, anacua, and ebony blackbead. Spanish moss and tangled vines commonly support and conceal the nest. FOOD: Eats berries, especially hackberries, along with the fruit of mesquite, mangoes, junipers, palmettos, persimmons, wild grapes, and figs; also eats green leaves, buds and shoots of plants, grain (especially cracked corn), and some insects. REFERENCES: Johnsgard 1975a, Marion 1974, Marion and Fleetwood 1978, Oberholser 1974a, Terres 1980. 97 Gray Partridge Perdix perdix RANGE: Widely introduced in North America and established locally from southern British Columbia to southwestern Quebec, New Brunswick, and Nova Scotia south to northern Nevada, northern South Dakota, central Indiana, and northern Vermont. Native to Eurasia. STATUS: Locally abundant. HABITAT: Has adapted well to diverse and intensive agricultural land use practices. Prefers cropland areas interspersed with native grassland, but also inhabits brushy canyons and brushy stream bottoms in the West, irrigated agricultural lands, and gently rolling hay fields, grain fields, and pastures. In winter, needs protective woody cover during adverse weather and accessible food resources, such as waste grain and green plant material. SPECIAL HABITAT REQUIREMENTS: Herbaceous cover for nesting, and grit (primarily obtained from gravel roads) for grinding food. NEST: Nests in a shallow scrape in the ground, usually located in grasses, or near the edge of hay fields or grain fields, sometimes in alfalfa fields, or along fencerows or roadsides. FOOD: Feeds on cultivated grains, seeds of grasses and herbs, and on insects. Obtains some of its water from succulent vegetation. REFERENCES: Johnsgard 1975a, Kobriger 1983, McCabe and Hawkins 1946, Terres 1980, Weigand 1980. 98 Chukar Alectoris chukar RANGE: Introduced widely in North America, and established locally from south-central British Columbia to central and eastern Montana south to Baja California, southern Nevada, northwestern New Mexico, and south-central Colorado. Native to Eurasia. STATUS: Locally common. HABITAT: Inhabits open, rocky, sagebrush-grassland habitats from below sea level to as high as 12,000 feet, on dry mountain slopes and canyons. Also inhabits areas with Mormon tea, bitterbrush, currant, and rabbitbrush; in the southern portion of its range it may be found in saltbush-grassland habitat, but generally avoids pinyon-juniper climax habitat. During hot weather, tends to concentrate near water provided by springs, seeps, and small perennial and intermittent streams, dispersing when surrounding vegetation greens up after rains. Moves to lower elevations in heavy snows. Roosts on the ground beneath sagebrush, under junipers, in the shelter of rock outcrops, and in open rocky places, but not in dense cover. SPECIAL HABITAT REQUIREMENTS: Water source during hot weather in summer and early fall. NEST: Nests in a depression scratched in the ground, constructed under shrubs or well concealed by rocks and brush in rocky areas. May occasionally have a dump nest. FOOD: During summer and fall, feeds primarily on seeds of cheatgrass, Russian thistle, rough fiddleneck, and redstem filaree; also takes seeds of Indian ricegrass, curly dock, and mustard. Will also eat grass blades, stems and buds of a variety of plants, wild onion seeds, grasshoppers, and caterpillars. REFERENCES: Christensen 1970, Johnsgard 1975a, Molini 1976. 99 L 27” Ring-necked Pheasant Phasianus colchicus RANGE: Introduced and established widely in North America, from southern British Columbia and central Alberta to northern Minnesota, southwestern Quebec, New Brunswick, and Nova Scotia south, at least locally, to southern interior California, Utah, northern and southeastern Texas, southern Illinois, Pennsylvania, New Jersey, and North Carolina. Native to Asia. STATUS: Common. HABITAT: Inhabits cultivated farmland interspersed with patches of brush or woodlots, especially areas with field border cover. Also inhabits fallow fields, brushy pastures, roadside hedgerows, cutover land, marshes, and meadows. Roosts mostly on the ground in weedy ditches, marshes, cattail swales, weed-grown fence rows, brush heaps, briar patches, and small farm-woodlots. In winter, needs protective cover such as marshes, thickets, shelterbelts, and heavy brush in ravines, along fencerows, and railroad right-of-ways, and an adequate supply of food. SPECIAL HABITAT REQUIREMENTS: Agricultural land interspersed with dense protective cover. NEST: Nests on the ground in a scratched-out depression, preferably among plants that make maximum growth during spring, such as alfalfa or coolseason grasses. Locates nest in fields of alfalfa, sweet clover, winter wheat, or other grasses or grains; along roadsides, fencerows, and hedgerows; or in brushy pastures or wetlands. FOOD: Feeds primarily on plant foods, especially waste grains, but also on seeds of weeds and grasses, acorns, buds and soft parts of herbaceous vegetation, fleshy fruits, insects, snakes, and mice. REFERENCES: Baxter and Wolfe 1973, Dale 1956, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1975a, Ratti in Farrand 1983a, Terres 1980. 100 Spruce Grouse Dendragapus canadensis RANGE: Resident from northern Alaska to northern Quebec, Labrador, New Brunswick, and Nova Scotia south to south-coastal and southeastern Alaska, northern Oregon, southeastern Idaho, northwestern Wyoming, western Montana, and southeastern Alberta to northern Minnesota east to northern Vermont, northern New Hampshire, and eastern Maine (range is generally congruent with that of the northern coniferous forest). STATUS: Uncommon in most of the southern portions of its range. HABITAT: Inhabits short-needled coniferous forests, especially where living branches reach the ground and where there are numerous, scattered forest openings of a few hundred square feet. In the northeast, prefers wet lowland edges; farther west, prefers higher ground. Generally prefers a mixture of jack pine or lodgepole pine and spruce, and a sparse ground cover. Males establish breeding territories in dense pine stands. Shows little fear of humans and often called “fool hen.’’ SPECIAL HABITAT REQUIREMENTS: Large tracts of coniferous forest. NEST: Nests on the ground in well-concealed sites, often under low branches of spruce, jack pine, or white pine; in brush; in deep moss; or adjacent to a tree trunk or stump. FOOD: During winter, consumes diet of nearly 100 percent conifer needles. In other seasons, consumes a variety of foods, including leaves, flowers, berries, seeds, pine needles, and a few insects. Needs a source of fine, mineral-rich gravel. REFERENCES: Johnsgard 1973, Pendergast and Boag 1970, Robinson 1969, 1980, Robinson in Farrand 1983a. 101 Blue Grouse Dendragapus obscurus RANGE: Resident from southeastern Alaska, southern Yukon, and extreme southwestern Mackenzie south along the Pacific Coast to northern California; in mountains to central California, northeastern Nevada, northern Arizona, Colorado, and northern New Mexico; rare in mountains of southern California. Closely associated with Douglas-fir and true firs. STATUS: Common. HABITAT: Occupies a fairly vertical range in the western mountains, ' breeding at lower elevations in open stands of conifers or aspens with a brush understory, in meadows or in stands of mixed brush and herbs interspersed with bare ground adjacent to aspens or conifers. Males display in relatively open stands of trees or shrubs on earth mounds, rocks, logs, cutbanks, and occasionally tree limbs. During autumn, moves up from the open breeding range to stands of conifers or to timberline. SPECIAL HABITAT REQUIREMENTS: Medium to large forest openings; shrubs, grasses, and forbs for nesting. NEST: Nests in a shallow depression on the ground, well-concealed near logs or rocks, at the base of a tree, under sagebrush, or in the shelter of chokecherry, aspen, or cottonwood. FOOD: During winter, eats diet limited to needles and buds of conifers, especially Douglas-fir, but also other firs, hemlock, and lodgepole pine. In other seasons, eats berries, flowers and leaves of herbaceous plants, and insects. In warm weather seldom far from a source of water. Can obtain necessary water from succulent vegetation or berries, if available. REFERENCES: Beer 1943, Bendell and Elliott 1966, Harju in Farrand 1983a, Johnsgard 1973, Rogers 1968, Terres 1980, Verner and Boss 1980. 102 White-tailed Ptarmigan Lagopus leucurus RANGE: Resident from south-central Alaska, central Yukon, and southwestern Mackenzie south to southern Alaska, southern British Columbia, including Vancouver Island, and the Cascade Mountains of Washington, and along the Rocky Mountains from southwestern Alberta to northern New Mexico. Introduced into the high Sierra Nevada of California, Wallowa Mountains in Oregon, and the Uinta Mountains in Utah. Commonly migrates locally during winter to areas slightly below treeline. STATUS: Locally common in alpine tundra. HABITAT: Inhabits rocky tundra areas with sparse vegetation in high mountains. Breeds in territories adjacent to spruce-willow alpine timberline zone (krummholz) and also small windblown areas. Males tend to winter above tree line adjacent to breeding areas where wind prevents complete coverage of woody shrubs; females tend to winter in basins and drainages that are not as windblown and somewhat removed from tundra. SPECIAL HABITAT REQUIREMENTS: Alpine tundra. NEST: Nests on the ground in areas that become snowfree early in June and are somewhat protected from wind, such as under small shrubs or next to rocks larger than 6 inches. Females locate their nests near the fringe of a male’s breeding territory, but more importantly, near brooding areas where vegetation is short and rocks 6 inches or larger cover more than 50 percent of the ground surface. FOOD: During summer, primarily consumes seeds and leaves of smartweeds, sedges, clover, and willow; also takes various green leaves, flowers, and some insects. During winter in Colorado, consumes willow primarily, alder secondarily. In Alaska, consumes alder catkins primarily, willow and birch secondarily. REFERENCES: Braun 1969; Braun in Farrand 1983a; Braun and Rogers 1971; Johnsgard 1973, 1983a; May and Braun 1972; Weeden 1967. 103 Ruffed Grouse Bonasa umbellus 8 gray phase L 14" RANGE: Resident from central Alaska and northern Yukon to southern Labrador south to northwestern California, central and eastern Idaho, central Utah, Wyoming and Montana, central and southeastern Minnesota, Ohio, in the Appalachian Mountains to northern Georgia and northeastern Virginia; locally to western South Dakota; introduced and established in Iowa and Newfoundland. STATUS: Fairly common; population fluctuates. HABITAT: Inhabits successional to subclimax hardwood forests larger than 10 acres that have Betula or Populus present and an understory of small hardwoods, shrubs, and fruit-producing bushes (early successional stages of plant growth on logged-over areas are ideal). Male uses logs, rocks, or other elevated sites for drumming in spring. Frequents hedgerows and brushy patches in early fall; moves into more heavily wooded areas, especially coniferous cover in winter. Roosts in snow when snow is deep and soft, or may roost in trees or on the ground. SPECIAL HABITAT REQUIREMENTS: Hardwood forests with some conifers, dense undergrowth, openings, and drumming sites for males. NEST: Nests on dry ground in the shelter of a fallen log, rock, root, or low-hanging conifer limb, usually near the base of a tree. Commonly nests within 100 feet of a road, path, or clearing, and close to a source of water. FOOD- During winter, feeds primarily on aspen buds, but also on buds of birch, alder, and hazel. In other seasons, consumes an extremely varied diet, including over 600 species of plants (seeds, fruits, leaves, and buds), insects, and other animals, although animal food is only predominate in the diet the first 2 weeks after hatching. REFERENCES: Boag and Sumanik 1969, Bump et al. 1947, DeGraff et al. 1980, Johnsgard 1973, Rue 1973. 104 Sage Grouse Centrocercus urophasianus RANGE: Resident locally from central Washington, southern Idaho, Montana, southeastern Alberta, southwestern Saskatchewan and North Dakota, and western South Dakota south to eastern California, south- central Nevada, southern Utah, Colorado and northern New Mexico. Generally, associated with big sagebrush. STATUS: Common (formerly widespread); range has been reduced, and its population is lower than it was 30 years ago. HABITAT: Inhabits sagebrush-dominated rangelands, from the plains to the mountains, preferably where sagebrush provides 15 to 50 percent of the ground cover. Depends entirely upon forms of sagebrush, primarily big sagebrush, for food from October through May and for cover throughout the year. In spring, males prefer relatively open, rather than dense, sagebrush cover for strutting grounds. May move up to 50 miles or more throughout the year; occupies areas with exposed sagebrush during winter. SPECIAL HABITAT REQUIREMENTS: Sagebrush-dominated rangelands. NEST: Usually nests beneath sagebrush in a shallow depression on the ground. Usually locates nest in drier sites close to strutting grounds where shrub cover is less than 50 percent and vegetation is 10 to 20 inches tall. Moves young broods to moist areas, where there is a plentiful supply of insects and green plant material. FOOD: In winter, feeds entirely on sagebrush leaves. Prefers Wyoming big sagebrush to mountain big sagebrush, thus maximizing proteins and minimizing monoterpenes. Also feeds (to a small extent) on alkali sagebrush and, in Idaho on black sagebrush. In other seasons, also feeds on forbs and some insects. REFERENCES: Braun in Farrand 1983a, Dalke et al. 1963, Johnsgard 1973, 1983a, Patterson 1952, Remington 1983, Tate and Tate 1982. 105 Greater Prairie-Chicken Tympanuchus cupido (includes Attwater’s Prairie-Chicken) RANGE: Resident locally from eastern North Dakota, northwestern and central Minnesota, northern Wisconsin, and northern Michigan south to northeastern Colorado, Kansas, southern and northeastern Oklahoma, central Missouri, and southern Illinois; also in southeastern Texas. STATUS: Endangered in Texas; uncommon and local; range and population reduced because of agriculture, burning, mowing, overgrazing, oil development, drainage, shooting, and urban development. HABITAT: Inhabits stands of natural tall or midgrass prairie, especially where natural grasslands are interspersed with moderate amounts of small-grain cropland. Males favor slightly elevated open areas of short grassland for display grounds, which are approximately 1 acre in size and surrounded by dense grasses with some brush cover. Moderately tall vegetation is used for night roosting, and edges of tallgrass and midgrass for day resting. Requires a stable source of food rather than protective cover or shelter for winter. SPECIAL HABITAT REQUIREMENTS: Shortgrass and tallgrass prairies. NEST: Nests in a slight depression on the ground in a well-drained site that provides good concealment from above, within 1/2 mile of display grounds, in ungrazed meadows, natural prairie stands, or in clumps of prairie grasses, usually near an open area. Chooses brood habitat that is usually heavier than nesting habitat, in old fields, native grasses, or in cultivated pastures, where shade and a plentiful supply of insects and succulent plants are available. FOOD: Consumes mostly insects from May to October, especially grass¬ hoppers. Consumes primarily plant foods, including fruits, leaves, flowers, acorns, seeds of grasses and weeds, and grains the rest of the year. REFERENCES: Johnsgard 1975a, 1983a, Jones 1963, Mackenzie 1977, Terres 1980. 106 Lesser Prairie-Chicken Tympanuchus pallidicinctus RANGE: Resident locally from southeastern Colorado, south-central Kansas, and western Oklahoma to extreme eastern New Mexico and the Texas Panhandle. STATUS: Uncommon and local. HABITAT: Inhabits arid natural grasslands of the southern Great Plains that are interspersed with shrubs 3 feet tall or less. In Colorado, occupies plant communities dominated by sand sagebrush, little bluestem, switchgrass, sideoats grama, and red threeawn. In Oklahoma and New Mexico, occupies a shinnery oak vegetation type. Needs small trees or shrubs such as shin oak, sagebrush, skunkbrush, sand plum, Havard oak, sand sagebrush, and fragrant sumac for shade during warm summer months. In spring and fall males congregate on display grounds which are relatively void of vegetation and have good visibility. Tall, perennial grasses are used for loafing in winter. SPECIAL HABITAT REQUIREMENTS: Shin oak or sand sagebrush rangelands. NEST: Nests on the ground in well-drained sites within 1/2 mile of display grounds. Builds nests that are well concealed from above in ungrazed meadows, natural prairie, or between grass clumps from previous years’ growth, generally in tall, dense, perennial grasses. FOOD: Primarily insectivorous during summer months. Consumes plant foods during the remainder of the year; leaf galls, catkins, leaves and acorns of shin oak can make up 70 percent of the seasonal diet; leaf and flower buds of fragrant sumac and the leaves of sand sagebrush are also important foods. REFERENCES: Copelin 1963; Doerr and Guthery 1980; Hoffman 1963; Johnsgard 1975a, 1983a; Jones 1963; Taylor and Guthery 1980. 107 Sharp-tailed Grouse Tympanuchus phasianellus RANGE: Resident, at least locally, from central Alaska and central Yukon to northern Ontario and west-central Quebec, south to eastern Oregon, central Utah, central Colorado, central Nebraska, central Minnesota, central Wisconsin, northern Michigan, and southern Ontario. STATUS: Locally common; population is down in the northern Great Plains. HABITAT: Inhabits mid to tall grasslands interspersed with scattered woodlands, arid sagebrush, brushy hills, oak savannah, and edges of riparian woodland. Prefers habitats with several small openings, 1 to 10 acres in size, close together or a single large opening of 50 to 100 acres. In late summer and early autumn found mainly in open cover, grassy openings, or in low, scattered brush. Later, moves to thickets and open woods; in winter, to edges of brush, open woods, or swamps. Prefers tops of low to medium hills or ridges with short, sparse vegetation for courtship sites. Roosts in trees or shrubs in brushy cover. SPECIAL HABITAT REQUIREMENTS: Grasslands interspersed with shrubs or trees and grainfields. NEST: Nests on the ground, preferably among tall, rank grasses, but also in brushy or woody areas if grassland quality is poor. May nest in fields of winter wheat, or in residual cover of warm season grasses on north-facing slopes. FOOD: Eats wide variety of foods, primarily the leaves, seeds, or buds of plants, and some insects, but also including fruits, cultivated grains, buds and catkins of birch and aspen, and grasshoppers. REFERENCES: Hamerstrom 1963, Hamerstrom and Hamerstrom 1951, Johnsgard 1973, 1983a, Rogers 1969, Sisson 1976, Tate and Tate 1982. 108 Wild Turkey Meleagris gallopavo (formerly Turkey) RANGE: Resident locally from central Arizona and central Colorado to northern Iowa, central Michigan, southern New Hampshire, and southwestern Maine south to southern Texas, the Gulf Coast, and Florida. Has been reintroduced into much of its former range, and successfully introduced locally in nearly all states outside the historic range. STATUS: Locally fairly common. HABITAT: Inhabits a wide range of forest types from the wooded swamps of the eastern and southeastern states to the sparsely wooded flatlands and river bottoms of the southern Great Plains and coniferous forests of the western mountains. In the East, prefers open, mature hardwood forests containing mast-bearing trees such as oaks; in the Southwest, prefers more arid, grass-dominated habitats having open- topped roosting trees, water, and succulent vegetation. In the West, most often associates with ponderosa or montane forests, scrub oaks, and junipers at altitudes of 6,000 to 12,000 feet. SPECIAL HABITAT REQUIREMENTS: Mast-producing woodlands with forest openings or clearings, large dense conifers or hardwoods for roosting, and water. NEST: Nests in a slight depression on dry ground, usually in dead leaves at the base of a tree, beneath a bush, or under a log. Generally nests close to strutting grounds and near water. In western mountains, it usually nests on north-facing slopes from 7,000 to 9,500 feet in elevation. FOOD: Diet is 90 percent plant foods, including mast of oaks, beeches, and pines; fruits; seeds and grains; and greens of grasses and forbs. Also eats roots, tubers, and insects, especially grasshoppers and walking sticks. REFERENCES: Boeker and Scott 1969, DeGraff et al. 1980, Hillestad 1973, Johnsgard 1975a, Korschgen 1967, Ligon 1946, Lindzey 1967, Markley 1967. 109 Montezuma Quail Cyrtonyx montezumae (formerly Harlequin Quail) RANGE: Resident, at least locally, from central and southeastern Arizona, southern New Mexico, and extreme southwestern Texas south into Mexico. STATUS: Locally fairly common. HABITAT: Inhabits shaded grassy oak canyons, wooded mountain slopes with bunchgrass, and rocky ravines. Seldom goes far from pine-oak woodlands due to dependence on succulent, bulb-producing forbs that grow in pine-oak understory. Heavy grazing tends to reduce bulb- producing forbs, which are replaced by perennials that can provide adequate cover, but not the underground food reserves required during dry seasons. SPECIAL HABITAT REQUIREMENTS: Ungrazed pine-oak woodlands. NEST: Conceals nest in a slight depression on the ground, at the base of trees, in grassy meadows, next to boulders, or under shrubs. FOOD: Coveys typically feed in close groups, digging out bulbs of nutgrasses, wood sorrels, and other plants. Also eats acorns, seeds of legumes, grasses, and pines; fruits of shrubs and trees; and insects. Water is primarily obtained from food. REFERENCES: Johnsgard 1975a, Leopold and McCabe 1957, Phillips et al. 1964, Terres 1980, Wallmo 1954. 110 Northern Bobwhite Colinus virginianus (formerly Bobwhite) 9 RANGE: Resident from southeastern Wyoming and central South Dakota to southern Ontario, southern New Hampshire and southern Maine, south through the central and eastern United States to Florida and southern Arizona into Mexico; introduced and established in western North America. STATUS: Often common to abundant. Masked bobwhite extirpated in southern Arizona but is being reintroduced. HABITAT: Inhabits open pastures, meadows with abundant weedy growth, and cultivated or fallow agricultural lands with hedgerows and dense brush, near open woodlands; avoids deep woods. In the Southwest, found in brushy canyons and hillsides or on dry grasslands with scattered mesquite and cactus. Prefers to winter in coveys (within several miles of breeding areas) in areas where dense cover of brushy thickets, hedgerows, or brush piles provide protection and abundant food resources. SPECIAL HABITAT REQUIREMENTS: Open woodlands adjacent to fields and brushy cover. In winter, brushy cover within 150 feet of feeding areas. NEST: Builds a well-concealed nest on dry ground, usually in a moderately dense stand of herbaceous and grassy vegetation such as goldenrods, panic grasses, cheatgrass, broom sedge, and bluegrass, with scattered shrubs and briars, and patches of bare ground. Prefers areas where standing vegetation is usually less than 20 inches high and upright stems are separate enough for the birds to pass between. FOOD: Consumes seeds, fruits, buds, and other plant parts that contribute 95 percent of the diet in winter and 70 percent in summer. Eats primarily green plant leaves in spring. Also eats insects that account for the remaining diet. REFERENCES: DeGraff et al. 1980, Klimstra and Roseberry 1975, Reid and Goodrum 1979, Robel 1969, Rosene 1969, Tate and Tate 1982, Terres 1980. Ill Scaled Quail Callipepla squamata RANGE: Resident from south-central Arizona, northern New Mexico, east-central Colorado, and southwestern Kansas south through western Oklahoma, western half of Texas, and into Mexico. Introduced and established in central Washington and eastern Nevada. STATUS: Common. HABITAT: Inhabits dry, open country in valleys, plains, or foothills that have a mixture of bare ground, low herbaceous growth, and scattered brushy cover such as mesquite, soapweed, acacias, mimosas, scrub oaks, and other semi-desert shrubs. May be found on rocky, brushy slopes, draws, gullies, canyons, on sand sage grassland, and sometimes on shortgrass plains, pastures, and cultivated fields. If shrub cover is lacking, uses structures such as corrals, feedlots, and buildings for shade, resting areas, and escape cover. Winters in large coveys, usually within 1 1/4 miles from water. SPECIAL HABITAT REQUIREMENTS: Desert grassland or desert scrub with a minimum of 1 loafing covert per 70 acres, and a source of water. NEST: Nests in a slight hollow on the ground in a protected and shaded site. The nest may be under shrubs or among grasses, under old machinery or junk, or under overhanging rocks. FOOD: Feeds primarily on plant seeds, with some fruits and insects. REFERENCES: Ault and Stormer 1983; Goodwin and Hungerford 1977; Johnsgard 1973, 1975a; Kelso 1937; Schemnitz 1961; Stormer 1981. 112 Gambel’s Quail Callipepla gambelii 8V2" RANGE: Resident from east-central California, southern Nevada, southern Utah, western Colorado, and northwestern New Mexico south to Mexico and the Rio Grande Valley of western Texas. STATUS: Common. HABITAT: Inhabits desert scrub dominated by desert hackberry, mesquite, catclaw, buckhorn and cholla cactus; chaparral; and riparian areas; from sea level to elevations above 6,500 feet. Roosts in shrubs or low trees, where shade, brushy escape cover and succulent vegetation are available within about 1500 feet. SPECIAL HABITAT REQUIREMENTS: Open water or succulent vegetation. NEST: Nests on the ground in a scratched-out depression, usually well concealed under tall grass, mesquite, sage, or other shrubs. Occasionally nests above ground in woodpiles, rotted stumps, or abandoned nests of thrashers, roadrunners, or cactus wrens. FOOD: Consumed a diet of 44 percent forb seeds, 14 percent shrub seeds, 5 percent animal foods, 3 percent grass seeds, and 34 percent leafy vegetation in one study. Eats a variety of foods including deer- vetch, filaree, mesquite, paloverde, lupine, white-thorn, mimosa, saguaro, calowrightia, and insects (largely ants.) Eats succulent foods to maintain body moisture. REFERENCES: Goodwin and Hungerford 1977, Gorsuch 1934, Harrison 1979, Hungerford 1962, Johnsgard 1975a. 113 California Quail Callipepla californica RANGE: Resident from southern British Columbia, Washington, and western Idaho south through most of Oregon, California, and Utah to Baja California. Most populations north of southern Oregon and east of California apparently the result of introductions. STATUS: Common over most of its range. HABITAT: Tolerates a relatively broad variety of climates, from cool, wet coastal areas to arid desert. Lives in foothills and valleys where low trees or shrubs have openings of weeds and grass near water, also in coastal scrub, broken chaparral, edges of woodlands, riparian woodland, and on farms, ranches, and parks. Roosts from 15 to 25 feet up in dense growth of trees or shrubs at night; uses brushy thickets during the day for roosting and escape cover. SPECIAL HABITAT REQUIREMENTS: Brushy cover within 50 feet of feeding areas, and near a source of water. NEST: Builds nests in a slight depression on the ground or in tall, dense weedy or grassy cover, well concealed at the base of a tree or shrub, near a fallen tree or under a brush pile. Nests often located along fence rows, in road ditches, or in pastures and close to water. FOOD: Consumes mostly seeds and leafy green vegetation, with only small amounts of animal food in the spring and summer. REFERENCES: Bevier in Farrand 1983a, Browning 1977, Emlen and Glading 1945, Gutierrez 1979-1980, Leopold 1977, Sumner 1935, Terres 1980. 114 Mountain Quail Oreortyx pictus RANGE: Resident from southwestern British Columbia, western and southern Washington, and central Idaho south through the mountains of California and northern and western Nevada to Baja California. STATUS: Common. HABITAT: Inhabits open montane forests with a well-developed brushy understory, steep slopes around edges of mountain meadows, and in logged or burned-over forests, from 1,500 to 10,000 feet. In California, breeds from chaparral up to lodgepole pine forests, preferring areas with much shrubbery and low percent canopy cover. During winter, moves to a lower elevation and occupies a habitat of mixed trees, brush, and herbs that produce mast and seeds. SPECIAL HABITAT REQUIREMENTS: Water during the breeding season. NEST: Nests on the ground, in a well-concealed site, often under fallen pine branches, in weeds or shrubs, at the base of large trees, or beside large rocks. Nests are always near water. FOOD: Forages on the ground and in low shrubs for food. Consumes leaves, flowers, and buds of succulent vegetation; mast; pine seeds; tubers, roots, and bulbs; seeds and fruits; and insects. REFERENCES: Gutierrez 1979-1980, Johnsgard 1975a, Ormiston 1966, Terres 1980, Verner and Boss 1980. 115 Yellow Rail Coturnicops noveboracensis RANGE: Breeds locally from northwestern Alberta and southern Mackenzie to southern Quebec and New Brunswick south to southern Alberta, North Dakota, southern Wisconsin, southern Ontario, Massachusetts, and Connecticut. Winters from coastal North Carolina south to southern Florida, west along the Gulf Coast to central and southeastern Texas; in Mexico; and locally from Oregon south to southern California. STATUS: Depending on locality, common, rare, or casual. HABITAT: Highly secretive; spends most of its time beneath dense, rank vegetation. Inhabits shallow, freshwater, grassy, and sedge marshes and wet meadows. In fall and winter, lives in high margins of fresh and saltwater marshes, savannahs, grainfields, hayfields, and among garden crops. SPECIAL HABITAT REQUIREMENTS: Marshes or wet meadows. NEST: Prefers to nest in drier portions of marsh, usually where ground is damp but there is no standing water. Conceals nest in dense clumps of marsh grasses, with surrounding vegetation forming canopy over the nest. FOOD: Eats small snails, insects, seeds of sedges, grasses, and clover leaves (very little has been reported on food habits). REFERENCES: Anderson 1977, Devitt 1939, Terres 1980, Walkinshaw 1939. 116 Black Rail Laterallus jamaicensis RANGE: Breeds locally along the Atlantic Coast from New York south to central Florida; on the Gulf Coast in eastern Texas and western Florida; in Kansas, and from California to South America. Winters along the coast of California from the breeding range north to Tomales Bay; in the Imperial and lower Colorado River Valleys of southeastern California; along the Gulf Coast from southeastern Texas east to Florida; and in South America. STATUS: Locally common. HABITAT: Inhabits coastal salt marshes, and occasionally inland freshwater marshes and wet meadows. Uses grainfields and hay meadows to some extent. Prefers higher portions of a marsh, where vegetation is rank and dense and the ground is damp. Life history poorly known; spends much of its time under matted grasses in high salt marshes. SPECIAL HABITAT REQUIREMENTS: Marshes with areas of dense, but not necessarily tall, cover and damp soil. NEST: Completely hides nest in clumps of marsh grass or at the base of glasswort, in or along the edge of a marsh. Usually places nest on matted dead grass, but sometimes directly on damp ground. FOOD: Probably feeds mainly on invertebrates. REFERENCES: DeGraff et at. 1980, Pough 1951, Terres 1980, Todd 1977. 117 Clapper Rail Rallus longirostris RANGE: Resident along the Atlantic and Gulf Coasts from Connecticut south to southern Florida and west to southern Texas; locally along the Pacific Coast from central California south to South America; and in the interior of southeastern California and southwestern Arizona at the southern end of the Salton Sea and in the lower Colorado River Valley (northernmost populations tend to be partially migratory). STATUS: Abundant in the east, local in California. Some western subspecies are listed as endangered. HABITAT: Primarily inhabits coastal salt marshes, especially tidal marshes bordered by shallow bodies of salt or brackish water. One subspecies inhabits freshwater marshes along the Colorado River. SPECIAL HABITAT REQUIREMENTS: Dense growths of cordgrass or needlerush, with deep soft soils. NEST: Builds nest 8 to 12 inches above ground and near water in salt marsh cordgrass, in marsh elder, or other marsh vegetation that is more than 2 feet tall, with a canopy of vegetation. Locates nest so it will not be submerged by high tide. FOOD: Prefers to feed on mudflats or along muddy shores of creeks at low tide by probing and gleaning. Eats fiddler crabs, other small crabs, crustaceans, snails, shellfish, aquatic insects, and some seeds. REFERENCES: Adams and Quay 1958, DeGraff et al. 1980, Johnsgard 1975a, Kozicky and Schmidt 1949, Mangold 1977, Stone 1937. 118 King Rail Rallus elegans RANGE: Breeds locally from eastern Nebraska and central Minnesota to Connecticut south through northwestern and central Kansas, central Oklahoma, and most of the eastern United States to western and southern Texas, central Mississippi and Alabama, and southern Florida. Winters primarily from southern Georgia, Florida, the southern portions of the Gulf States, and southern Texas south to Mexico. STATUS: Uncommon; population declining in Midwest prairies, low elsewhere. HABITAT: Inhabits coastal, and inland brackish and freshwater marshes with abundant vegetation (especially sedges, bulrushes, and cattails), roadside ditches, tidal rivers, ricefields, and upland fields near marshes. Forages and nests along waterways made by the muskrat (distribution coincides closely with that of the muskrat). Not known to breed in salt marshes, but wintering birds inhabit coastal brackish, salt (rarely), and freshwater marshes. SPECIAL HABITAT REQUIREMENTS: Wetlands with abundant vegetation and fairly stable water levels during the breeding season. NEST: Conceals nests with a cone-shaped or round canopy of vegetation overhead; usually locates nests 6 to 18 inches above shallow water on grass or sedge tussocks or on hummocks among cattails. FOOD: Feeds on mud flats at low tide, in open roadside ditches, or in very shallow water, 2 to 3 inches deep. Primarily eats crustaceans, especially crayfish and aquatic insects; also takes grasshoppers, crickets, fishes, frogs, grains (especially rice in winter), and seeds of aquatic plants. REFERENCES: Bateman 1977, DeGraff et al. 1980, Johnsgard 1975a, Meanley 1969, Tate and Tate 1982. 119 Virginia Rail Rallus limicola RANGE: Breeds locally from southern British Columbia and northwestern Alberta to southern Quebec, New Brunswick, and southwestern Newfoundland south to Baja California, west-central Texas, Kansas, central Ohio, western Virginia, and along the Atlantic Coast to North Carolina: also in central Louisiana and northern Alabama. Winters from southern British Columbia and western Washington south to Baja California; and from central Texas, the Gulf Coast, and coastal North Carolina south locally to Costa Rica; casual in interior North America. STATUS: Common. HABITAT: Inhabits freshwater marshes, marshy borders of lakes and streams, and occasionally brackish and salt marshes. Prefers areas with shallow water and abundant emergent vegetation. SPECIAL HABITAT REQUIREMENTS: Wetlands with sedge and cattail edge. NEST: Usually constructs nests in cattails or sedges a few inches above shallow water, that are often covered with a loose canopy of vegetation and well attached to surrounding vegetation or on a clump of grass or tussock. FOOD: Consumes a diet that is about two-thirds insects. Also eats small fishes, frogs, mollusks, crustaceans, leeches, and earthworms. Climbs easily on reeds and in bushes to reach seeds and berries, and also eats grass seeds, duckweeds, wild oats, and rice. REFERENCES: Berger 1951, DeGraff et al. 1980, Horak 1970, Johnsgard 1975a, Low and Mansell 1983, Walkinshaw 1937, Zimmerman 1977. 120 Sora Porzana Carolina L 6 3 /V' RANGE: Breeds from southern Yukon and west-central and southwestern Mackenzie to west-central and southern Quebec and southwestern Newfoundland, south locally to Baja California, central Arizona, New Mexico, central Illinois, and Maryland. Winters from central California to the Gulf Coast and southern South Carolina south to South America; occasionally north to southern Canada. STATUS: Common. HABITAT: Prefers freshwater marshes, but also inhabits brackish and salt marshes, ponds, swamps, bogs, wet grassy meadows,and sloughs, especially those with sedges and cattails. In late summer, concentrates in areas where food is plentiful such as in rice fields or other seed- producing areas. SPECIAL HABITAT REQUIREMENTS: Wetlands with abundant, dense vegetation. NEST: Constructs a well-concealed nest that may be fastened to or supported by reed stems on a raised platform of vegetation. Locates nest 6 inches to over a foot above water, or occasionally on the ground. Generally nests over water 6 to 8 inches deep, preferably among sedges. FOOD: Primarily feeds on small mollusks and insects except in the fall; then relies heavily on seeds in freshwater habitats and animal foods in brackish areas. Also eats insects, mollusks, worms, crustaceans, and small tadpoles. Eats seeds and leaves for most of the vegetative portion of the diet. REFERENCES: DeGraff et al. 1980, Horak 1970, Johnsgard 1975a, Low and Mansell 1983, Meanley 1965, Odom 1977, Walkinshaw 1940, Webster 1964. 121 Purple Gallinule Porphyrula martinica L10” RANGE: Breeds along the Atlantic Coast from Maryland and Delaware south through Central America to South America, and in eastern and southern Texas, the Gulf States, and Florida; locally in southern Illinois, western Tennessee, and central Ohio. Winters from southern Texas, Louisiana, and Florida south throughout remainder of breeding range. STATUS: Uncommon throughout breeding range. HABITAT: Characteristically inhabits marshy wetlands with a variety of marsh plants, especially where pickerelweed and plants with floating leaves, such as water lilies, are abundant. Walks freely on lily pads and other floating vegetation, alights readily on bushes, and climbs about in branches over water. SPECIAL HABITAT REQUIREMENTS: Freshwater marshes and swamps, ponds, and channels of slow moving water with well-vegetated edges. NEST: Usually nests on an island of floating water plants. Builds a well- concealed nest, sometimes suspended and woven into marsh vegetation or willow thickets up to 6 feet above shallow water. FOOD: Consumes both plant and animal foods, including the seeds of rice, duckweed, wild millet, and other aquatic plants; also eats mollusks, aquatic insects, snails, and worms. REFERENCES: Cramp and Simmons 1980, Holliman 1977, Johnsgard 1975a, Low and Mansell 1983, Terres 1980. 122 Common Moorhen Gallinula chloropus (formerly Common Gallinule) RANGE: Breeds locally in California, central Arizona, and northern New Mexico, and from central Minnesota and southern Wisconsin to Vermont and Massachusetts, south to South America. Winters in eastern North America from South Carolina and the Gulf Coast southward, elsewhere throughout the breeding range, occasionally north to Utah, Minnesota, southern Ontario, and New England. STATUS: Locally common. HABITAT: Inhabits freshwater marshes, lakes, ponds, slow-flowing streams and rivers, and nearly any body of water with emergent vegetation such as cattails, bulrushes, reeds, sedges, and burreeds growing in water over one foot deep. Also inhabits rice fields in the South. SPECIAL HABITAT REQUIREMENTS: Emergent vegetation growing in water 1 to 3 feet deep and areas of open water. NEST: Builds nests in emergent vegetation 2 to 6 feet high, on a hummock or other clump of vegetation over water that is 1 to 3 feet deep. Locates nest at water level or up to 2 feet above water, concealing it with a canopy of surrounding plants. Occasionally nests in shrubs such as willow or alder. Builds brood platforms or uses muskrat houses or platforms built by coots. FOOD: Dives, dabbles, or wades while foraging; eats mainly leaves and stems of underwater plants, duckweed, leaves of grasses and herbs, and seeds and berries. Occasionally eats insects, earthworms, slugs, snails, and rarely aquatic vertebrates such as tadpoles or small fishes. REFERENCES: DeGraff et al. 1980, Fredrickson 1971, Johnsgard 1975a, Krauth 1972, Low and Mansell 1983, Strohmeyer 1977. 123 American Coot Fulica americana RANGE: Breeds from east-central Alaska and southern Yukon to southwestern Quebec, southern New Brunswick, and Nova Scotia south locally to Central America. Winters from southeastern Alaska and British Columbia south through the Pacific States, and from Colorado and northern Arizona to the lower Mississippi and Ohio Valleys, and Maryland south to the southeastern United States and Central America. STATUS: Common. HABITAT: Inhabits marshes, lakes, ponds, sloughs, potholes, and marshy borders of creeks and rivers, or ephemeral habitats when conditions are suitable. Prefers wetlands with a good interspersion of emergent vegetation, especially cattails and bulrushes. During migration, found on rivers, lakes, ponds, reservoirs, or sewage lagoons, and on freshwater or brackishwaters in winter. SPECIAL HABITAT REQUIREMENTS: Freshwater wetlands, with shallow water 1 to 4 feet deep and emergent vegetation interspersed with areas of open water. NEST: Constructs floating display platforms, egg nests, and brood nests located in and anchored to emergent vegetation, usually within 4 feet of open water and over water 1 to 4 feet deep. May also nest on top of a muskrat house. FOOD: Primarily dabbles and grazes, but also dives. Eats a variety of plant foods, which make up 89 percent of the diet. During winter often grazes on pastures and cultivated fields. Eats mostly pondweeds, sedges, algae, grasses (especially wild rice), and other plants; also eats fishes, tadpoles, worms, crustaceans, spiders, beetles, dragonflies, and bugs. REFERENCES: Cramp and Simmons 1980; DeGraff et al. 1980; Fjeldsa 1977; Fredrickson 1970, 1977; Gullion 1954; Johnsgard 1975a; Jones 1940; Kiel 1955. 124 Limpkin Aramus guarauna RANGE: Resident from southeastern Georgia and northern Florida (east of the Apalachicola River) south through peninsular Florida, Central America, and South America. STATUS: Locally common. HABITAT: Inhabits freshwater cypress swamps, marshes of saw grass or bulrushes, bayous, sloughs, and wooded marshy rivers or streams. Sometimes perches in tops of the tallest trees or on shrubs. SPECIAL HABITAT REQUIREMENTS: Freshwater marshes with dense vegetation or trees and shrubs for nesting. NEST: Commonly nests among clumps of tall grasses or other aquatic plants above shallow water. Sometimes nests 5 to 8 feet above the ground in bushes, vines, or trees along banks of streams, or on platforms just above water in dense vegetation. FOOD: Probes for food in shallow water or in soft mud on mud flats for freshwater snails, especially Pomacea; also eats mussels, frogs, lizards, worms, insects, and a few seeds. REFERENCES: Bent 1926, Harrison 1975, Oberholser 1974a, Sykes in Farrand 1983a, Terres 1980. 125 Grus canadensis Sandhill Crane RANGE: Breeds from western and central Alaska and northern Yukon to Baffin Island, south locally to southern Alaska, northeastern California, Colorado, southern Minnesota, southern Michigan, and western Quebec. Resident from southern Mississippi, southern Alabama, and southern Georgia south to Florida. Winters from central California and southeastern Arizona to the Gulf Coast and southern Georgia south to Mexico. STATUS: Northern subspecies (Lesser, Canadian, and Greater) are migratory and locally common, while nonmigratory subspecies (Cuban, Florida, and Mississippi) are threatened or endangered. HABITAT: Inhabits prairies, tundra, open pinewood flats, and other open areas. Breeds in or near shallow marshes, bogs, sloughs, margins of lakes, ponds, and river deltas. In mountainous regions, inhabits isolated, well-watered river valleys, marshes, and meadows. Occasionally inhabits relatively small marshes and patches of prairie in forested country. During winter, roosts in flocks at night on low damp ground or in shallow water, and flies to feeding grounds at dawn. SPECIAL HABITAT REQUIREMENTS: Shallow wetlands adjacent to a meadow, cultivated fields, or open woodlands and free of human disturbance. NEST: Usually nests in or near shallow wetlands adjacent to feeding grounds. Nests are located on a mound of emergent vegetation, sticks, grass, moss, or mud among rushes, sedges, grasses or other tall, dense vegetation. Pairs mate for life. FOOD: Feeds in cultivated fields, pastures, wet meadows, and marshes. Eats wheat, corn, alfalfa, sorghum, barley, roots and tubers, berries, small mammals, snakes, frogs, lizards, crickets, and grasshoppers. REFERENCES: Archibald in Farrand 1983a, Cramp and Simmons 1980, Johnsgard 1975a, Lewis 1977, Terres 1980, Walkinshaw 1949. 126 Whooping Crane Grus americana RANGE: Breeds in the vicinity of Wood Buffalo National Park in south-central Mackenzie and in northern Alberta. Introduced at Grays Lake National Wildlife Refuge, Idaho. Winters in the vicinity of Aransas National Wildlife Refuge on the Gulf Coast of Texas, occasionally northeast to southern Louisiana. STATUS: Endangered, numbering fewer than 150 individuals in the traditional wild flock, about 13 in the introduced Rocky Mountain flock, and about 55 in captivity. HABITAT: Inhabits marshy areas interspersed with shallow potholes having soft marly bottoms and a pH range of 7.6 to 8.3. Primarily inhabits aspen parkland, but also in northern coniferous forest, shortgrass plains, northern mixed forest, river deltas, and tundra. Winters on tallgrass prairies, salt flats, coastal marshes, lagoons, and brackishwater areas. SPECIAL HABITAT REQUIREMENTS: Large, shallow wetlands that provide visibility over a wide area and are free of human disturbance. NEST: Nests on a mound of bulrushes in shallow water, on islands or along shores of large wetlands where there is a heavy cover of bulrushes. Pairs mate for life, and return to the same general area each year but not to the same nest site. FOOD: In winter, primarily feeds on blue crabs, but also eats aquatic insects, freshwater minnows, shrimps, crayfishes, marine worms, snails, clams, sprouting corn, acorns, grasses, sedges, and other plants. (Foods of other seasons have not been well documented.) REFERENCES: Allen 1952, Archibald in Farrand 1983a, Mackenzie 1977, McNulty 1966, Novakowski 1966, Terres 1980. 127 Black-bellied Plover Pluvialis squatarola RANGE: Breeds from northern Alaska south to western Alaska, and from northwestern Mackenzie and Banks Island, southern Melville, Devon, and western and southern Baffin Islands south to the Yukon River, north- central Mackenzie, northern Keewatin, and Southampton and Coats Islands. Winters primarily in coastal areas from southern British Columbia and New Jersey south along both coasts of the United States to South America. STATUS: Common. HABITAT: Breeds on moist to dry upland rolling tundra. In other seasons frequents mudflats, beaches, shores of ponds and lakes, flooded fields, and salt marshes. Commonly associates with other shorebirds, especially willets, golden plovers, knots, and curlews. SPECIAL HABITAT REQUIREMENTS: Tundra during breeding season. NEST: Nests in a depression on the ground in relatively dry sites on or near a ridge, often in a prominent area affording a wide view. Usually locates nest on gravelly ground, sometimes with large boulders or with sparse vegetation of lichens, dryad, saxifrage, willows, sedges, or grasses. FOOD: Feeds along seacoasts on broad tidal sand flats and mud flats and in salt marshes, or inland around lakeshores, in meadows and upland pastures, or in plowed fields. Diet includes marine worms, small mollusks, crustaceans, marine insects, grasshoppers, locusts, cutworms, grubs, beetles, earthworms, and some seeds and berries. REFERENCES: Bent 1929, Hussell and Page 1976, Palmer 1967, Terres 1980. 128 Lesser Golden-Plover p luvialis dominica [formerly American Golden Plover) RANGE: Breeds from northern Alaska and northern Yukon to Banks, Devon, and northern Baffin Islands south to central Alaska, northwestern British Columbia, southern Keewatin, northern Ontario and Southampton, and southern Baffin Islands. Winters in southern South America. STATUS: Common, once abundant; population decline due to hunting at the end of the 19th century but is recovering. HABITAT: Breeds on arctic and subarctic tundra beyond treelimit, usually where the ground cover is lichens and mosses. In migration, occupies short-grass pastures, plowed fields, and burned-over meadows, or beaches and mudflats in coastal areas. SPECIAL HABITAT REQUIREMENTS: Dry, grassy tundra. NEST: Nests in a depression on dry ground, preferably on higher sites such as banks of gullies or streams but not necessarily near water. Eggs are difficult to see against tundra. Young birds quickly move to wetter areas such as sphagnum swamps. FOOD: Feeds in rolling pasturelands with short, scanty grass, on prairies, plains, plowed land, open sand, or mud flats. Mostly eats insects, especially grasshoppers and crickets. Also eats grubs, cutworms, caterpillars, beetles, spiders, mollusks, small crustaceans, small fishes, crowberries, and blueberries. REFERENCES: Bent 1929, Cramp and Simmons 1983, Palmer 1967, Terres 1980. 129 Snowy Plover Charadrius alexandrinus im. L5‘/2” RANGE: Breeds along the Pacific Coast from southern Washington to Baja California, and locally from southern Oregon, western Nevada, southwestern Montana, central Kansas, and north-central Oklahoma south to southeastern California and north-central Texas; also along the Gulf Coast from Florida west to Texas. Winters on islands and in coastal areas from northern Oregon and the Gulf Coast south to Costa Rica. STATUS: Locally common; suffering a serious decline in the southern and middle Pacific Coast regions. HABITAT: Inhabits dry sandy coastal beaches above the wash of the tides, sand spits or bars separating the ocean from coastal wetlands, estuarine margins, alkali flats, dry lake beds, or the shores of salt ponds and alkali lakes. Prefers open habitats; avoids thick vegetation and narrow beaches littered with driftwood or backed by bluffs where it might be trapped by high water. SPECIAL HABITAT REQUIREMENTS: Open nesting habitat, preferably near water. NEST: Nests singly or sometimes in loose colonies in flat areas devoid of, or sparsely covered with, vegetation or driftwood. Generally nests near water, but occasionally farther away if no formidable barrier is between the nest and water, in a scrape on the ground, usually among small rocks, kelp, or other objects. FOOD: Forages on wet sand of beaches, at the surf line, or along muddy or alkaline shores of ponds and lakes inland. Food is mostly small crustaceans, marine worms, other minute marine animals, beetles, flies, and other insects. REFERENCES: Bent 1929, Harrison 1979, Page and Stenzel 1981, Palmer 1967, Tate and Tate 1982, Terres 1980, Wilds in Farrand 1983a. 130 Semipalmated Plover Charadrius semipalmatus winter summer L5 3 /r RANGE: Breeds from northern Alaska, northern Yukon, Banks, Victoria and central Baffin Islands to the northern Labrador Coast, south to western Alaska, southwestern and central British Columbia, northern Manitoba, central Quebec, and southern Nova Scotia. Nonbreeding birds often summer in the wintering areas south to Panama. Winters primarily in coastal areas from central California, the Gulf Coast, and South Carolina south to South America. STATUS: Common. HABITAT: Found during spring and fall migration on beaches, mudflats, lakeshores, riverbanks, freshly plowed fields, shallow marshes, and peat banks. It breeds in dry Arctic tundra, sometimes quite far from water, and prefers lichen-grown gravelly tundra or areas of rubble and patches of stranded debris. Avoids grassy areas. SPECIAL HABITAT REQUIREMENTS: Level, well-drained gravelly tundra. NEST: Nests in unsheltered scrapes on level ground in gravelly or sandy soil, or in moss or lichens and above the high-water line in loose colonies. FOOD: Prefers to feed in shallow streams or mudbars in tundra ponds, but also forages on tidal flats. Diet includes marine worms, mollusks, small crustaceans, larvae of midges and other insects, ants, beetles, grasshoppers, and spiders. REFERENCES: Cottam and Hanson 1938, Palmer 1967, Sutton and Parmelee 1955, Terres 1980. 131 Piping Plover Charadrius melodus summer winter L5‘/2 RANGE: Breeds locally from south-central Alberta to south-central Manitoba, south to eastern Montana and central and eastern Nebraska; in the Great Lakes region from northern Michigan and southern Ontario south to the southern shores of Lake Michigan and Lake Ontario; and in coastal areas from Newfoundland south to Virginia. Winters on the coast from South Carolina south to Florida and west to eastern Texas, sparsely in Bahamas and Greater Antilles. STATUS: Endangered; numbers are slowly declining. HABITAT: Inhabits exposed, sparsely vegetated sandy shores and islands of shallow lakes and ponds, dry sandy ocean beaches, higher portions of strand near dunes, and large open sandy areas, especially where scattered grass tufts are present. In winter it is found on beaches, margins of lagoons, and areas of rubble. SPECIAL HABITAT REQUIREMENTS: Unspoiled, undeveloped beaches with little vegetation. NEST: Nests in a hollow in sand, well beyond high tide on ocean beaches, on raised sandspits, or on the lower slopes of dunes. Generally nests on narrow beaches as little as 6 feet wide. May sometimes nest under tufts of grass. Adults tend to return to the same breeding area year after year. FOOD: Forages on beaches, along margins of watercourses, and on tidal sandflats for marine worms, insects such as fly larvae and beetles, crustaceans, and mollusks. REFERENCES: Bent 1929, Cairnes 1982, DeGraff et al. 1980, Harrison 1979, Palmer 1967, Tate and Tate 1982, Terres 1980, Wilcox 1959. 132 Killdeer Charadrius vociferus RANGE: Breeds from east-central and southeastern Alaska and southern Yukon to central Quebec and western Newfoundland, south to Mexico. Winters from southern British Columbia across the central United States to New England and south throughout the remainder of North America to South America. STATUS: Common throughout range. HABITAT: Indiscriminately occupies open areas, but favors open dry uplands, meadows, pastures, and disturbed or heavily grazed areas where grass is short, sparse, or absent. In winter, inhabits open fields, beaches, along watercourses, and on mudflats. SPECIAL HABITAT REQUIREMENTS: Open areas with closely cropped or sparse vegetation. NEST: Nests in a scrape on gravelly or bare ground, occasionally on a flat or gently sloping roof of a building, often with a few pebbles, grasses, or weeds in the scrape. FOOD: Prey is snatched or gleaned from the surface of dry or moist ground or from very shallow water. Eats a diet that is mainly insects, especially grasshoppers, beetles, and dragonflies, but also includes centipedes, spiders, worms, snails, crayfish, and weed seeds. REFERENCES: Cramp and Simmons 1983, DeGraff et al. 1980, Palmer 1967, Terres 1980. 133 Mountain Plover Charadrius montanus winter UVi" RANGE: Breeds from extreme southern Alberta and northern Montana south to central and southeastern New Mexico, western Texas, western Oklahoma, and western Missouri. Winters from central California, southern Arizona, and central and coastal Texas south to Mexico. STATUS: Common but declining. HABITAT: A species of the high plains and arid regions of western valleys and hills, usually found far from water. Generally avoids mountainous areas and prefers areas dominated by blue grama grass and buffalo grass. In winter, congregates in flocks of 15 to several hundred on alkali flats, plowed ground, grazed pastures, or other open arid habitats. SPECIAL HABITAT REQUIREMENTS: Shortgrass prairie and arid plains. NEST: Nests in scrapes on flat ground, preferably in areas of blue grama-buffalo grass with scattered clumps of cacti and western wheat grass. Avoids tall vegetation. FOOD: Consumes mostly, if not entirely, insects caught on the dry plains and prairies, primarily grasshoppers, crickets, beetles, and flies. REFERENCES: Bent 1929, Garrett in Farrand 1983a, Graul 1975, Palmer 1967, Terres 1980. 134 Black-necked Stilt Himantopus mexicanus RANGE: Breeds locally on the Atlantic Coast from southern New Jersey south to southern Florida, and from southern Oregon, southern Colorado, central Kansas, the Gulf Coast of Texas, and southern Louisiana south to South America. Winters from central California, the Gulf Coast of Texas and Louisiana, and southern Florida south to South America. STATUS: Common. HABITAT: Inhabits shallow freshwater and brackish ponds, alkaline lakes, wet meadows, open marshes, and flooded fields and pastures. Commonly associates with other shorebirds, especially avocets, godwits, and curlews. SPECIAL HABITAT REQUIREMENTS: Shallow wetlands. NEST: Nests in slight depressions on the ground, on sandy or gravelly shores, or along drier margins of ponds and lakes, or on deep, well-built, floating platforms over shallow water in loose colonies. Also nests on hummocks, on small islands, or under clumps of vegetation and may be well concealed or in the open. FOOD: Forages along muddy shores and in shallow waters for dragonfly nymphs, caddis flies, mayfly nymphs, flies, billbugs, mosquito larvae, grasshoppers, crayfish, and snails. Also consumes small fishes and seeds of aquatic and marsh plants. REFERENCES: Bent 1927, Palmer 1967, Terres 1980, Wilds in Farrand 1983a. 135 American Avocet Recurvirostra americana RANGE: Breeds from southeastern British Columbia and central Alberta to Minnesota, south locally to southern California, northern Utah, and southern New Mexico, and east to central Kansas and coastal Texas. Winters mostly in coastal lowlands from northern California and southern Texas south to Mexico, and locally in southern Florida. STATUS: Common. HABITAT: Inhabits the borders of muddy saline, alkaline, and freshwater ponds, lakes, and marshes, particularly favoring shallow alkaline lakes, wet meadows and pastures with scattered open pools. Commonly associates with black-necked stilts, godwits, and lesser yellowlegs. SPECIAL HABITAT REQUIREMENTS: Wetlands bordered by open flats or areas with scattered tufts of grass. NEST: Eggs are laid in a scrape on the ground. Nests in colonies on dry, sun-baked mudflats, near water, on low, gravelly or sandy islands with scant vegetation, or in marshes bordering shallow lakes. If the water level rises to flood the nest, adds vegetation to raise the nest above the level of water. FOOD: Gathers food by stirring up water for seeds, aquatic insects, and small crustaceans, and by probing in soft mud. Consumes dragonfly nymphs, back swimmers, water boatmen, beetles, grasshoppers, crickets, weevils, flies and their larvae, centipedes, small snails, small crustaceans, small fishes, and seeds of marsh and aquatic plants. REFERENCES: Bent 1927, Gibson 1971, Harrison 1979, Low and Mansell 1983, Palmer 1967, Terres 1980, Wilds in Farrand 1983a. 136 Greater Yellowlegs Tringa melanoleuca RANGE: Breeds from southern Alaska, southwestern Mackenzie, and south-central British Columbia east across the northern and central portions of the Canadian Provinces to central and southern Labrador, Newfoundland, and northeastern Nova Scotia. Nonbreeding birds sometimes summer on the wintering grounds, especially along the coasts of the United States. Winters from Oregon and southern Nevada to southern Texas, the Gulf Coast, and coastal South Carolina south to South America. STATUS: Common. HABITAT: Found in the Nearctic boreal region to the edge of subarctic coniferous forest zone, where it inhabits swampy muskegs or bogs with scattered trees, wet clearings and pools, or tundra. Perches freely when breeding, often alighting on tops of trees, bushes, or dead stubs. Outside the breeding season, frequents shallow fresh, brackish, and salt waters, mudflats, river bars, tidal marshes and pools, rain pools in fields, and damp grassy meadows. SPECIAL HABITAT REQUIREMENTS: Muskeg and tundra. NEST: Nests in a depression on the ground, usually near trees, logs, or stumps, on a dry wooded ridge or on recently burned ground, and normally near water. FOOD: Feeds by picking, snatching, skimming, and sweeping, but not by probing. Favors mudflats and shallow borders of lakes and streams for feeding sites, where it finds small fishes, tadpoles, worms, mollusks, snails, crabs, and insects and their larvae. REFERENCES: Cramp and Simmons 1983, Low and Mansell 1983, Palmer 1967. 137 Lesser Yellowlegs Tringa flavipes RANGE: Breeds from central Alaska and central Yukon to northern Manitoba, northern Ontario, and extreme west-central Quebec south to east-central British Columbia, across to southeastern Manitoba. Winters along the Pacific and Atlantic Coasts from southern California and South Carolina south to South America. STATUS: Common. HABITAT: Inhabits the Nearctic coniferous forest zone, from boreal and subarctic regions into the low Arctic; occurs mainly inland, and to some extent upland. Prefers grassy meadows and bogs, natural clearings, or burned areas in forest with scattered stumps and fallen logs, often far from open water. Outside the breeding season, inhabits shallow prairie sloughs in open country, muddy shores of lakes and marshy ponds, sewage beds, river margins, and inland and coastal marshes. SPECIAL HABITAT REQUIREMENTS: Tundra and muskeg. NEST: Nests in a depression on the ground, singly or in loose colonies. Locates nest on a dry sloping bank, ridge, or level plateau, in open high woodland with sparse, fairly low undergrowth, in swampy muskeg, or on undrained land surrounded by farmland. FOOD: Forages by picking and snatching food from shallow water, especially in wet, shortgrass marshes, or in shallow ponds, wet cultivated fields, or on mudflats. Eats ants, bugs, flies, grasshoppers, insect larvae, small fishes, crustaceans, and worms. REFERENCES: Bent 1927, Cramp and Simmons 1983, Low and Mansell 1983, Palmer 1967. 138 Solitary Sandpiper Tringa solitaria RANGE: Breeds from central and south-coastal Alaska and northern Yukon to northern and central Ontario, east through central Quebec to central and southern Labrador, and south to northwestern and central British Columbia across to southern Manitoba and northern Minnesota. Winters from the Gulf Coast, southeastern Georgia, and Florida south to South America. STATUS: Common. HABITAT: Inhabits inland muskeg with scattered mature trees or clumps of trees near freshwater lakes and ponds in the coniferous forest belt of boreal and subarctic regions. On its breeding range it perches freely on treetops, twigs, limbs, and stumps. Outside of the breeding season it occurs inland along shallow freshwater woodland streams, ponds, bogs, flooded marshes, stagnant pools, mudflats, and barnyard puddles. SPECIAL HABITAT REQUIREMENTS: Muskegs. NEST: Nests up to 43 feet above ground in the old nests of American robins, waxwings, rusty blackbirds, and eastern kingbirds. Usually uses nests in coniferous trees that border muskeg or open bogs, or beside a lake. FOOD: Forages while walking about on stranded vegetation, in soft mud, or in very shallow water. Eats aquatic insects, especially larvae, also worms, grubs, dragonfly nymphs, water-scavenger beetles, water boatmen, grasshoppers, caterpillars, spiders, small crustaceans, and small frogs. REFERENCES: Bent 1929, Cramp and Simmons 1983, Palmer 1967, Pough 1951. 139 Willet Catoptrophorus semipalmatus RANGE: Breeds locally from eastern Oregon, central Alberta, and southwestern Manitoba, south to northeastern and east-central California, across to western and northern Nebraska and eastern South Dakota; locally along the Atlantic and Gulf Coasts from Prince Edward Island south to southern Florida and west to southern Texas. Occurs sporadically (nonbreeding birds) in summer as far south as northern South America. Winters along the Pacific and Atlantic Coasts from northern California and Virginia south to South America. STATUS: Common. HABITAT: Inhabits tidal and coastal marshes, beaches, sandy islands with tall and thick grasses, open pastures, dry uplands near water, and along dunes in the East. Prefers inland prairies and plains, alkali flats, and grassy dikes, usually near water in the West. Associates freely with godwits, curlews, large plovers, and some shorebirds. Often perches on bushes, trees, fences, posts, rocks, and buildings. SPECIAL HABITAT REQUIREMENTS: Moist plains and prairies in western North America, coastal marshes and nearby grassy areas in the East. NEST: Nests semi-colonially in depressions on the ground or in a thick clump of vegetation, sometimes far from water in the West. Locates nest in open areas on a sandy beach to well hidden in low grasses. FOOD: Forages by probing or by snatching food from the ground on tidal flats, in salt or brackish marshes, water-soaked pastures, along muddy creek banks, and on saline flats. Eats fiddler crabs, mollusks, marine worms, small fishes, adult insects, and some seeds, leaves, and roots of grasses. REFERENCES: Bent 1927, Low and Mansell 1983, Palmer 1967, Stenzel et al. 1976, Terres 1980, Wilds in Farrand 1983a. 140 Spotted Sandpiper Actitis macularia RANGE: Breeds from central Alaska and central Yukon to Labrador and Newfoundland, south to southern Alaska, southern California, and central Arizona across to the northern portions of the Gulf States, North Carolina, Virginia, and eastern Maryland. Occasionally nonbreeding birds remain on the wintering ground in summer. Winters from southwestern British Columbia, southern Arizona, southern New Mexico, southern Texas, the southern portions of the Gulf States and coastal South Carolina south to South America. STATUS: Common. HABITAT: Inhabits the edges of ponds, lakes, rivers, and streams and open terrain with temporary pools, up to 14,000 feet elevation. It is sometimes found far from water in dry fields, pastures, and weedy shoulders of roads, occasionally on coastal beaches and dunes. Roosts on stumps, stranded logs, or rocks affording a clear view. In winter, frequents watercourses shaded by trees, and prefers shallow, muddy lagoons, creeks, canals, and higher mudflats. SPECIAL HABITAT REQUIREMENTS: Margins of freshwater bodies. NEST: Builds solitary or loosely colonial nests on the ground, among thick, tall grasses, occasionally under a bush or log, and usually near water. FOOD: Forages ashore or in shallow water, picking up insects and other small invertebrates. Eats fly larvae, pupae, and adults; mayflies; grasshoppers; crickets; mole crickets; worms; mollusks; crustaceans; and spiders. REFERENCES: Cramp and Simmons 1983, DeGraff et al. 1980, Knowles 1942, Palmer 1967. 141 Upland Sandpiper Bartramia longicauda RANGE: Breeds locally from north-central Alaska, northern Yukon, and northern Alberta to southern Quebec, central Maine, and southern New Brunswick south to northeastern Oregon, central Colorado, north-central Texas, central Missouri, West Virginia, and Maryland. Winters in South America. STATUS: Uncommon; once abundant, numbers have been reduced due to past hunting pressure and agricultural practices. HABITAT: Inhabits grassy open areas, ranging from sandy, sparsely vegetated flats to open, grassy bogs and muskeg. Most often found in rich pastureland, hayfields, and alfalfa fields. During the breeding season alights freely fenceposts, telephone poles, and other elevated sites. During migration, frequents alfalfa fields, pastures, prairie dog towns, and rarely shores and mudflats. SPECIAL HABITAT REQUIREMENTS: Open grasslands. NEST: Nests in depressions on the ground among rank grasses, along sloughs in prairies, or in clearings of spruce muskeg, in loosely spaced colonies. Conceals nests are by covering them with nearby vegetation. FOOD: Prefers to forage where the grasses are low and open enough to provide good visibility, and where grasshoppers and crickets are most abundant. Also eats weevils, ants, berries, waste grain, and seeds of grasses and weeds. REFERENCES: Cramp and Simmons 1983, DeGraff et al. 1983, Palmer 1967, Tate and Tate 1982. L 142 RANGE: Breeds from south-central British Columbia to southern Manitoba, south to northeastern California, central Utah, central New Mexico, and northern Texas, and east to southwestern Kansas. Winters from central California, southern Texas, southern Louisiana, and coastal South Carolina south to Mexico. STATUS: Fairly common; once fairly abundant, lower population is due to past hunting pressure and loss of habitat due to grazing and agriculture. HABITAT: Inhabits grasslands ranging from moist meadowland to very dry prairie. When at or near water, often loosely associates with godwits, willets, and yellowlegs. During the breeding season, commonly perches on bushes, low trees, dirt mounds, rocks, stumps, fenceposts, utility poles, or on other elevated sites. In other seasons, frequents wet habitats such as the shallow margins of inland and coastal waters, open areas of marshes, intertidal zones, or on sandbars. SPECIAL HABITAT REQUIREMENTS: Prairies or grassy meadows. NEST: Nests in a slight hollow on the ground, usually in flat areas among short grasses such as cheatgrass and bluegrass. Locates nests in moist or arid areas far from water. FOOD: Forages in areas that have a large variety of plants. Feeds by probing and pecking in wet sand or mud, under shallow water, or in uplands. Eats beetles, grasshoppers, caterpillars, other insect larvae, mud crabs, fiddler crabs, ghost shrimps, occasionally small fishes, and berries. REFERENCES: Allen 1980, Palmer 1967, Stenzel et al. 1976, Tate and Tate 1982. 143 Hudsonian Godwit Limosa haemastica RANGE: Breeds locally in south-coastal and western Alaska, Mackenzie, northwestern British Columbia, and around Hudson Bay. Winters in South America. STATUS: Locally common. HABITAT: Inhabits wet bogs and marshes in open expanses along the northern edge of boreal forests and moist tundra, near tidal or fluvial shorelines. In other seasons, frequents fresh, brackish, and salt waters, on beaches, mudflats, marshes, flooded fields, and shallow ponds. SPECIAL HABITAT REQUIREMENTS: Extensive sedge marshes and meadows near tidal flats. NEST: Nests in a depression on top of a hummock in a sedge marsh or meadow with water up to several inches deep. Frequently locates nest under a dwarf birch, occasionally on a sedge or grass tussock, under or among small willows, or near a fallen spruce tree, but usually where the vegetation is at least 5 to 6 inches high and thick enough to conceal the nest from the sides. FOOD: Feeds on marine worms, horseflies, mosquitoes, and other insects, mollusks, and crustaceans. Prefers to feed in water about as deep as its bill is long. REFERENCES: Bent 1927, Hagar 1966, Mackenzie 1977, Palmer 1967, Wilds in Farrand 1983a. 144 Marbled Godwit Limosa fedoa RANGE: Breeds from central Alberta to southern Manitoba and northern Ontario south to central Montana, northeastern South Dakota, and northwestern Minnesota. Some nonbreeding birds occur on the winter range during summer. Winters from central California, western Nevada, the Gulf Coast and coastal South Carolina south to South America. STATUS: Common. HABITAT: Inhabits grassy plains, broad flat wet meadows, and prairie sloughs, usually near lakes, rivers, or streams. During migration, frequents open coastal beaches and along lake shores; in winter, inhabits tidal flats in sheltered bays, next to inlets, and on open beaches. Gregarious and often in the company of willets and avocets. SPECIAL HABITAT REQUIREMENTS: Grassy prairies near water. NEST: Builds single, or possibly semi-colonial, nest in slight hollow in short grasses, often in plain sight and usually near water. FOOD: Forages by probing in shallow water and soft mud of sloughs and lake shores for aquatic insects and mollusks, and by capturing grasshoppers and other insects on meadows and low-lying prairies. Also eats snails, small crustaceans, insect larvae, worms, and leeches. REFERENCES: Bent 1927, Palmer 1967, Wilds in Farrand 1983a. 145 Ruddy Turnstone Arenaria interpres winter L 7" RANGE: Breeds from northern Alaska and the Canadian Arctic islands south to western Alaska, and Southampton, Coats, and Mansel Islands, probably also the northern portions of Mackenzie and Keewatin. Nonbreeding birds may be found through the winter range in summer. Winters along the Pacific and Atlantic Coasts from central California and New York south to South America. STATUS: Common. HABITAT: Inhabits flat, lichen-covered, mossy, or gravelly tundra near the seacoast in a variety of boreal habitats. Often perches on boulders, stakes, pilings, piers, and boats during breeding season. In other seasons, frequents rocks, reefs, and mussel beds of the intertidal zone, sandy beaches, and solid mudflats. SPECIAL HABITAT REQUIREMENTS: Dry, dwarf-shrub tundra near the coast. NEST: Nests in a depression in tundra in exposed or sheltered sites such as beside a rock or clump of vegetation. Uses a wet area near the nest site for brood rearing. FOOD: Forages among seaweeds, rocks and shells, and roots in wet sand. Eats mollusks, crustaceans, worms, flies and their larvae picked from the carcasses of seals and whales, eggs of gulls and terns, grasshoppers, soft parts of barnacles, sand fleas, and fiddler crabs. REFERENCES: Bent 1929, Palmer 1967, Terres 1980, Wilds in Farrand 1983a. 146 Red Knot Calidris canutus winter RANGE: Breeds in northwestern and northern Alaska and the Canadian Arctic islands east to Ellesmere Island and south to southern Victoria and Southampton Islands. Nonbreeding birds occasionally summer in the wintering range. Winters along the Pacific and Atlantic Coasts from southern California and Massachusetts south to South America. STATUS: Locally common; once abundant, its numbers were reduced by hunting in the late 19th century. HABITAT: Inhabits high inland plains, plateaus, and elevated slopes covered with glacial gravel and frost-riven rocks and shales, sometimes several miles from the coast. In migration and in winter, occurs mainly along the coast on exposed mudflats, sand spits, beaches, matted salt marshes, and river deltas. SPECIAL HABITAT REQUIREMENTS: Barren or stony tundra and dry nesting sites. NEST: Nests in a shallow depression among dryads, lichens, and other tundra vegetation, rubble, and gravel, and usually on high, dry hills and plateaus. FOOD: Flies to fresh waters or marine shores from nesting areas up to several miles away to feed during the breeding season. Mainly feeds on the surface, but also probes for food in mud and sand. Eats insects, mollusks, crustaceans, king crab eggs, and some seeds, buds and shoots of grasses, sedges, pondweeds, and bulrushes. REFERENCES: Bent 1927, Davis in Farrand 1983a, Palmer 1967, Sperry 1940, Terres 1980. 147 Sanderling Calidris alba summer RANGE: Breeds in northern Alaska, and from Prince Patrick Island to northern Ellesmere Island, south to northern Mackenzie, northern Keewatin, and Southampton, and northern Baffin Islands. Nonbreeding birds occur in summer in the winter range. Winters in the Aleutian Islands (locally), and from southern Alaska, the Gulf Coast, and Massachusetts south along the Atlantic and Pacific Coasts to South America. STATUS: Common. HABITAT: Inhabits high Arctic tundra, particularly dry clay-mixed stony plains sparsely covered with willows, dryad, and saxifrage, on islands, peninsulas, and along the coast. Outside the breeding season, frequents sandy coastal beaches and tidal flats. Inland migrants inhabit sandbars along rivers and on lake beaches. SPECIAL HABITAT REQUIREMENTS: Dry tundra. NEST: Nests in a slight hollow or depression on the ground on stony, well-drained ridge tops, gentle slopes, or on level alluvial plains. Usually locates nest at the edge of, or in, clumps of low plants and within several hundred yards of a marshy pond, but occasionally nests up to a mile from wet tundra. FOOD: Probes in wet sand, picks up food washed on shore by the tide, and snatches up insects. In winter and on migration, feeds on sand fleas, shrimp, and other small crustaceans, small mollusks, and marine worms. On the breeding grounds, feeds on flies and their larvae, and other insects. REFERENCES: Bent 1927, Cramp and Simmons 1983, Davis in Farrand 1983a, Parmelee 1970, Palmer 1967. 148 Semipalmated Sandpiper Calidris pusilla RANGE: Breeds from the Arctic Coast of western and northern Alaska north to Victoria and central Baffin Islands, east to northern Labrador, and south to western Alaska and east-central Mackenzie across to northern Ontario, northern Quebec, and coastal Labrador. Nonbreeding birds may summer in coastal North America south to the Gulf Coast and Panama. Winters from southern Florida south to South America. STATUS: Abundant. HABITAT: Inhabits subarctic and low to high Arctic tundra from coasts, dunes, borders of tidal inlets and deltas to damp grassy flats in interior and wet riverside tundra. Often occurs near lakes or pools, shifting from first areas uncovered by melting snow and surface ice to others becoming clear shortly afterwards, including upland tundra. In other seasons, frequents mudflats, sandy beaches, and wet meadows, favoring the vicinity of water on tidal flats, lagoons, and ponds. SPECIAL HABITAT REQUIREMENTS: Grassy or hummocky tundra. NEST: Nests in a slight depression on the ground, amid short herbage, sometimes in sand on grassy dunes or in low wet tundra near small lakes. FOOD: Forages by snatching food from surface and probing in soft mud on mudflats or in wet sand exposed by ebbing tide. Eats beetles, flies, fly larvae, mosquitoes, small mollusks, marine worms, small crustaceans, and bits of seaweeds. REFERENCES: Cramp and Simmons 1983, Davis in Farrand 1983a, Palmer 1967, Terres 1980, Townsend in Bent 1927. 149 Western Sandpiper Calidris mauri winter L 5'4" RANGE: Breeds on islands in the Bering Sea and along the coasts of western and northern Alaska. Winters from California and North Carolina south along both the Pacific and Atlantic Coasts to South America. STATUS: Common. HABITAT: Inhabits a complex mosaic of wet low-lying grass and sedge marshes dotted with small pools and lakes, and relatively well-drained heath-covered tundra such as hummocks, ridges, and better drained slopes of hills that are vegetated with mosses, lichens, dwarf shrub heath, dwarf birch, willows, and some herbs, grasses, and sedges. Outside of the breeding season, frequents mudflats, beaches, shores of lakes and ponds, and flooded fields. SPECIAL HABITAT REQUIREMENTS: Patches of dwarf shrub-heath tundra (at least 1/2 acre) interspersed among wet marshes for nesting. NEST: Nests on the ground in loose colonies or singly in nests that are usually well camouflaged under low vegetation. Locates nests in dry or moist areas, from the upper slopes of hills down to the marsh edge, or within a marsh on a patch of heath tundra. After fledging, moves into the marshes, frequenting margins of lakes and rivers. FOOD: Feeds by snatching or probing for food close to or in water; eats small invertebrates, including insect larvae and aquatic beetles, bugs, marine worms, and small snails. REFERENCES: Bent 1927, Cramp and Simmons 1983, Holmes 1971, Palmer 1967. 150 Least Sandpiper Calidris minutilla winter L W RANGE: Breeds from western Alaska and northern Yukon to southern Keewatin, northern Quebec, and northern Labrador south to the Alaska Peninsula, southeastern Alaska, and northwestern British Columbia across to northern Ontario, eastern Quebec, Nova Scotia, and Newfoundland; isolated breeding in Massachusetts. Nonbreeding birds summer in the wintering range, primarily in North America south to California and the Gulf Coast. Winters from coastal Oregon and southern Nevada to central Texas, the Gulf States, and North Carolina south to South America. STATUS: Very common. HABITAT: Inhabits open grass or sedge bogs and marshes in the northern spruce forest just south of treeless tundra, or among complexes of poois and water channels with scattered knolls and hummocks. Outside of the breeding season, prefers wet, muddy, or grassy areas such as muddy shores of grass fringed marshes or estuaries, grassy wet meadows, and grass-bordered mudflats of lakes, ponds, or rivers; found less frequently on sandy beaches. SPECIAL HABITAT REQUIREMENTS: Wetlands of subarctic boreal forests and tundra. NEST: Nests in a depression in a mossy hummock, a plant tuft, in a clump of grass, or sometimes on the ground, usually in marshy cover but sometimes in drier upland near water. FOOD: Prefers to feed in marshes, where it snatches insects or probes for food in soft mud or in shallow water. Eats midges and other dipterans, ground beetles, grasshoppers, insect larvae, small crustaceans, small mollusks, and worms. REFERENCES: Cramp and Simmons 1983, Cottam and Hanson 1938, Low and Mansell 1983, Palmer 1967, Pough 1951, Terres 1980. 151 White-rumped Sandpiper Calidris fuscicollis winter L 6Vt" RANGE: Breeds from northern Alaska to northern Bylot Island, south to the mainland coasts of Mackenzie and Keewatin, northwestern Hudson Bay, Southampton Island, and southern Baffin Island. Migrates along the Atlantic Coast and winters in South America. STATUS: Uncommon. HABITAT: Inhabits both lowland and upland tundra, frequently around bog pools, on dry ridges, or among grassy tussocks near rivers or lakes. Usually closely associated with moist, open terrain, but tolerates occasional freezing and snow cover and a wide range of temperatures. In migration, prefers shallow grassy pools, wet meadows, and marshes but also occurs on sandbars, mudflats, and beaches. SPECIAL HABITAT REQUIREMENTS: Mossy or grassy tundra. NEST: Nests in mossy depressions in clumps of grasses and sedges in the uplands, or in mossy hummocks on well-vegetated tundra that is persistently wet, often near marshy ponds and lake shores. Conceals nest, usually among grasslike plants, including narrow-leaved cotton- grass, grass rush, water sedges, and mosses. FOOD: Forages by snatching prey and by probing deeply in soft mud. Eats cranefly larvae, beetles, grasshoppers, clover-root curculio, and other insects, tiny mollusks, marine worms, and a few seeds. REFERENCES: Bent 1927, Cramp and Simons 1983, Drury 1961, Palmer 1967, Parmelee et al. 1968, Terres 1980. 152 Baird’s Sandpiper Zalidris bairdii L 6" RANGE: Breeds from western and northern Alaska to Ellesmere Island, south to central Alaska, northern Mackenzie, Keewatin, Southampton Island and south-central Baffin Islands. Winters in South America. STATUS: Uncommon. HABITAT: Inhabits dry coastal and alpine tundra, particularly barren, exposed ridges, terrace banks, and raised beaches that are sparsely covered with low matted vegetation. Prefers sheltered places and frequents muddy, sandy, and grassy areas near water, including irrigated fields, shores of lakes and ponds, alpine tundra, and marshes. During migration, prefers inland to coastal habitats. SPECIAL HABITAT REQUIREMENTS: Dry tundra. NEST: Nests are in shallow depressions on the ground, usually in wind¬ blown and lichen-strewn areas with large patches of bare soil. Sometimes nests in grassy areas in a tuft of vegetation or among lichen- covered rocks. FOOD: Forages by snatching food, primarily in dry areas or along wet edges of lakes and ponds. Eats small crustaceans, spiders, beetles and their larvae, gnats, craneflies, and other insects, moss, leaves and stems, and other plant parts. REFERENCES: Cramp and Simmons 1983, Davis in Farrand 1983a, Drury 1961, Palmer 1967, Terres 1980. 153 Pectoral Sandpiper Calidris melanotos L TVi" RANGE: Breeds from western and northern Alaska to Bathurst and Devon Islands, south to western Alaska, central Mackenzie, and southeastern Keewatin Districts, and the south coast of Hudson Bay. Winters in South America, casually north to the Gulf Coast and Florida. STATUS: Common. HABITAT: Inhabits dry and moss-lichen tundra, tundra grassland, tussocky tundra with sedge, and peat tundra with hummocks up to 10 to 15 feet. In Alaska, found along the coast and in the foothills, frequenting a variety of tundra habitats on flat terrain that are poorly drained, usually wet, and characterized by low grasses and sedges, dwarf shrubs, and cottongrass tussocks. Outside of the breeding season, prefers grassy terrain bordering moving or still waters; only rarely found on open mudflats. SPECIAL HABITAT REQUIREMENTS: Dry nesting sites on arctic tundra NEST: Nests in a depression on dry ground in areas with a continuous cover of grasses and sedges. Hides nest well, usually under a tree or bush. FOOD: Forages mainly in dry, grassy meadows on insects, especially flies and their larvae. Also eats beetles, crickets, grasshoppers, amphipods, other tiny crustaceans, mites, spiders, algae, and a few seeds of grasses, lupines, and violets. REFERENCES: Bent 1927, Cramp and Simons 1983, Low and Mansell 1983, Palmer 1967, Pitelka 1959. 154 Stilt Sandpiper Oalidris himantopus L 7W RANGE: Breeds from northern Alaska, northern Yukon, northern Mackenzie, and southern Victoria Island southeast to southeastern Keewatin, northeastern Manitoba, and northern Ontario, probably also south locally in Canada to borders of the taiga. Winters primarily in South America, but casually northward to southeastern California, the Gulf Coast, and Florida. STATUS: Uncommon. HABITAT: Inhabits sedge meadows interrupted by old beach ridges, eskers, or other elevated areas dominated by dwarf birch, heaths, willows, crowberries, and dryads. Sometimes occurs in wet tundra areas with fairly high willows, or on much drier slopes with moderate vegetative cover, avoiding truly barren ridgetops. SPECIAL HABITAT REQUIREMENTS: Well-drained sedge meadows in arctic tundra with elevated sites for nesting. NEST: Nests in a depression on the ground in relatively open areas of dry tundra, usually atop a hummock or on a low, well-drained gravel ridge; occasionally nests next to a shrub. Locates nest sites independent of standing water. May reuse old nests. Moves chicks from drying sedge meadows to wet areas. FOOD: Forages on dry ridgetops, around clumps of sedges, at the edges of tiny depressions filled with water, in marshes at tundra pond margins, by probing in soft mud on mudflats or while wading in water. Feeds opportunistically, on a relatively small spectrum of food including larval and adult beetles, larvae of other insects, flies and other flying insects, water bugs, small snails, and small seeds. REFERENCES: Cramp and Simmons 1983, Jehl 1973, Palmer 1967, Terres 1980. 155 Buff-breasted Sandpiper Tryngites subruficollis L 6Vi" RANGE: Breeds from northern Alaska to Banks, Melville, Bathurst, and Devon islands, south to northwestern Mackenzie, and Jenny Lind, and King William Islands. Winters in South America. STATUS: Uncommon; once abundant, its numbers were reduced by several decades of hunting pressure. HABITAT: Prefers raised and grassy terrain, sometimes by streams, but avoids marshy areas. Predominately a ground bird, it only occasionally occurs on beaches and along shores in migration, favoring short-grass prairies, burned-over grasslands, cotton fields, recently plowed fields, sun-baked stubble, and barren, recently inundated lands. SPECIAL HABITAT REQUIREMENTS: Dry, grassy tundra. NEST: Nests in a shallow cavity in dry, mossy or grassy tundra, sometimes near water or on high and dry banks of black tundra. FOOD: Forages primarily on insects gleaned from the surface. Eats adu and larval beetles, larvae and pupae of flies, some spiders, and seeds o smartweeds, pondweeds, and spikerushes. REFERENCES: Bent 1927, Cramp and Simmons 1983, Palmer 1967. 156 Short-billed Dowitcher L imnodromus griseus RANGE: Breeds from southern Yukon to northeastern Mackenzie, south to east-central British Columbia, amd across to central Saskatchewan; from the interior of the Ungava Peninsula south to northern Ontario, and in coastal regions of southern Alaska. Nonbreeding birds often occur south to wintering grounds in summer. Winters from central California, southern Arizona, the Gulf Coast, and coastal South Carolina south to South America. STATUS: Common. HABITAT: Primarily inhabits coniferous forest and muskeg with thin crusts of moss and long grass floating on a liquid morass, but also inhabits swampy coastal tundra. Low scrub of willow, alder, and birch are often present, with a few taller larch and spruce, sometimes flanked by dry ridges of dense coniferous forest or burned-over forests. After breeding, moves to open prairie lakes and sloughs. During migration and winter, occurs on mud and sand flats in sheltered bays and estuaries, on the borders of shallow pools in salt marshes, on sandy beaches, and in flooded fields. SPECIAL HABITAT REQUIREMENTS: Swampy coastal tundra or muskeg. NEST: Nests in a hollow in mosses, in a clump of grasses, or on dry ground in wet areas. Sometimes nests in a small clearing in coniferous forest near muskeg, but not under trees or in broken terrain. FOOD: Feeds by probing deeply in wet sand, mud, in shallow water, or in seaweeds. Eats flies, beetles, dragonfly nymphs, and other insects, as well as marine worms, snails, crabs, shrimps, and seeds of marsh and aquatic plants. REFERENCES: Cramp and Simmons 1983, Palmer 1967, Sperry 1940. 157 Long-billed Dowitcher Limnodromus scolopaceus RANGE: Breeds in coastal western and northern Alaska, northern Yukon, and northwestern Mackenzie. Winters from central California, southern Arizona, southern New Mexico, central Texas, the Gulf Coast, and southern Florida south to Panama. STATUS: Common. HABITAT: Found on arctic continental coastal belts and marginally within the subarctic just beyond treeline. Inhabits grassy and sedgy tundra, with or without scattered low woody vegetation and usually near shallow fresh water. In migration and winter, prefers grassy margins of shallow, muddy freshwater pools and, occasionally saltwater habitats. Associates freely with other shorebirds, including the larger plovers. SPECIAL HABITAT REQUIREMENTS: Grassy tundra and wet meadows. NEST: Nests in a shallow depression in a tuft of grass or in moss, on dry or moist ground, usually near freshwater. FOOD: Feeds in open unvegetated tracts of mud, as well as patches surrounded by tundra. Eats flies, beetles, small crustaceans and mollusks, marine worms, spiders, and seeds of aquatic plants. REFERENCES: Bent 1927, Cramp and Simmons 1983, Palmer 1967, Sperry 1940. 158 Common Snipe 3 allinago gallinago L9 RANGE: Breeds from northern Alaska and northern Yukon to southern Keewatin, northern Quebec, and central Labrador, south to central California, east-central Arizona, and northern Colorado across to northern West Virginia, New England, and the Maritime Provinces. Winters from southeastern Alaska, southern British Columbia, the central United States, and Virginia south to South America. STATUS: Common. HABITAT: Inhabits wetlands, especially fens, bogs, swamps, and marshes, primarily in peatlands scattered within the spruce, fir, and larch boreal forest. Occupies areas with fairly dense, low woody growth such as willows and alders, and with a ground cover of sphagnum, sedges, and grasses, preferably near open pastures or other clearings. Also inhabits areas of decomposed wet plant litter along ponds, meandering rivers and brooks, and other marshy sites. In winter it occupies wet, marshy habitats, wet meadows, flooded fields, and stream edges. SPECIAL HABITAT REQUIREMENTS: Bogs, fens, and swamps with moist organic soils near open areas free of obstacles or high vegetation that might interfere with display activities. NEST: Nests in a scrape on fairly dry ground or in a tussock of grass or sedge, usually in wet habitats but occasionally at the edge of wetlands. Conceals nest, sometimes covering it with an arch of dry vegetation. FOOD: Forages by probing in soft mud and shallow water, and by gleaning grasses and the surfaces of marsh plants. Diet consists primarily of animal foods including insects, crayfish, crabs, earthworms, and mollusks. Also eats some seeds. REFERENCES: DeGraff et al. 1980, Fogarty and Arnold 1977, Palmer 1967, Sperry 1940, Tuck 1972. 159 American Woodcock Scolopax minor RANGE: Breeds from southern Manitoba, northern Minnesota and south- central and southern Ontario to northern New Brunswick and Newfoundland, south throughout eastern North America to the Gulf States, and southern Florida, and west. Winters from eastern Oklahoma, southern Missouri, Tennessee, and Virginia south to east-central Texas, the Gulf Coast, and southern Florida. STATUS: Common; may be declining in parts of the East. HABITAT: Inhabits moist woodlands in early stages of succession especially, those with birch, aspen, red maple, alder, or willows under 25 feet tall and having an understory of conifers. Also found in alder swales with surrounding pockets of second-growth mixed hardwoods, old agricultural fields, burned or recently logged areas, areas too wet to support forest growth, hardwoods adjacent to streams and ponds, and brushy edges of woods. Uses open fields, cultivated land, pastures, and clearings at least 1/4 acre in size on relatively flat ground, and with a slight amount of ground cover for singing grounds. SPECIAL HABITAT REQUIREMENTS: Dense brushy swales with nearby fields or small forest openings for courtship activities and roost sites, and fertile, generally poorly drained soils containing an abundance of earthworms for feeding. NEST: Nests in a slight depression on dry ground, usually within a few yards of a brushy edge. Also nests on a hummock in wet areas, in open fields, or in young to middle-aged hardwoods of low to medium density. FOOD: Feeds in open pastures, cultivated fields, and along stream banks, probing in soft mud and leaf litter for earthworms, which make up 50 to 90 percent of the diet. Also eats insect larvae and adults, crustaceans, spiders, seeds, and berries. REFERENCES: DeGraff et al. 1980, Gregg and Hale 1977, Mendall and Aldous 1943, Owen 1977, Palmer 1967, Sepik et al. 1981, Sperry 1940. 160 Wilson’s Phalarope Phalaropus tricolor winter L 71 / 2 " RANGE: Breeds from southern Yukon and northern Alberta to southern Michigan and southwestern Quebec south to south-central California, east-central Arizona, west-central New Mexico, northern Texas, eastern South Dakota, northern Illinois, northern Indiana, and northern Ohio; isolated breeding in Massachusetts. Winters primarily in South America, casually as far north as southern California and southern Texas. STATUS: Uncommon. HABITAT: Once inhabited natural prairies but now found mainly on highly disturbed mixed-grass prairies dotted with small glacial potholes. Also found in taiga broken by moist, grassy muskeg and many small lakes and pools, and in farming country of aspen-grove parklands. Inhabits rolling uplands as high as 6,900 feet in elevation. Outside the breeding season, mainly found on inland wetlands but sometimes on saline or alkaline depressions. SPECIAL HABITAT REQUIREMENTS: Shallow water bordered by low grasses or sedges. NEST: Nests semi-colonially in a scrapes on the ground around damp meadows with marsh grasses and sedges or rushes. Also nests by shallow sloughs, morasses fringed with short grasses or sedges, lake shores, and in hay meadows or pastures, often 50 to 100 yards from water. FOOD: Forages on muddy shores and in shallow water; stirs up food by whirling its body around in water. Consumes larvae of mosquitoes and craneflies; predaceous diving beetles, aquatic bugs, brine shrimp, amphipods, eggs of water fleas; and seeds of aquatic plants. REFERENCES: Bent 1927, Cramp and Simmons 1983, Hohn 1967, Low and Mansell 1983, Palmer 1967. 161 Red-necked Phalarope Phalaropus lobatus winter L 6" summer RANGE: Breeds from northern Alaska and southern Victoria Island to central Keewatin and southern Baffin Island, south to northwestern British Columbia and northern Alberta across to northern Quebec, and locally along coast of Labrador. Winters mainly at sea in the Southern Hemisphere, largely in tropical and subtropical oceans. STATUS: Uncommon. HABITAT: Inhabits the wetter portions of flat alluvial plains, sedge-grass marshlands, clearings in alder and willow scrub, and heath covered slopes above alder and willow scrub. In winter, occurs near upwellings or where other local conditions produce a high biomass of accessible food organisms. SPECIAL HABITAT REQUIREMENTS: Wet grassy or sedgy terrain interspersed with pools, boreal clearings, or tundra. NEST: Nests in a small hollow in moss or among sedges, usually atop a small hummock surrounded by water or near a marshy pond or small stream. FOOD: Forages while swimming, wading, and walking, chiefly on invertebrates. During the breeding season and in migration, eats predominately small insects, especially adult flies and larvae. Also consumes mollusks, crustaceans, spiders, mites, worms, and rarely small fishes and tadpoles, as well as some seeds and algae. REFERENCES: Bent 1927, Clapp et al. 1983, Cramp and Simmons 1983, Hohn 1968, Palmer 1967. 162 Franklin’s Gull Larus pipixcan RANGE: Breeds from eastern Alberta and central Saskatchewan to western Minnesota, south locally to east-central Oregon, northwestern Wyoming, and northwestern Iowa. Winters primarily in South America, rarely in southern coastal California, and casually along the Gulf Coast of Texas and Louisiana. STATUS: Common. HABITAT: Breeds exclusively in shallow freshwater marshes and sloughs in the temperate prairie belt. Favors shallow wetlands up to 6 inches deep with bulrushes, cattails, whitetop, and common reeds, preferably near cultivated lands. In migration and in winter, inhabits sandy beaches, sandbars, fields, and pastures. SPECIAL HABITAT REQUIREMENTS: Marshes and sloughs with sparse emergent vegetation no denser than one plant less than 3 feet tall per square foot. NEST: Nests in colonies from a few hundred up to 50,000 pairs. Builds nests on masses of marsh vegetation, often floating on water and anchored to surrounding vegetation. Nests usually near open water. FOOD: During the breeding season, forages in marshes and fields up to 30 miles from the nesting site. Feeds on a wide variety of foods, especially insects, also taking some amphibians; seeds of wheat, oats, and barley; a few small mammals; and in winter, fish and crustaceans. REFERENCES: Clapp et al. 1983, Cramp and Simmons 1983, Guay 1968, Low and Mansell 1983, McAtee and Beal 1912. 163 Bonaparte’s Gull Larus Philadelphia Lit” RANGE: Breeds from western and central Alaska and central Yukon to northern Manitoba, south to southern British Columbia, central Saskatchewan, southern Manitoba, and southern James Bay. Occurs in summer (nonbreeding birds) south to coastal areas in California and New England, and in interior to the Great Lakes. Winters from Washington south along the Pacific Coast into Mexico and from the Great Lakes south through the Ohio and Mississippi Valleys to the Gulf Coast. STATUS: Common. HABITAT: Inhabits coastal and interior lowlands, primarily black fly- infested muskeg swamps in taiga up to treeline. Outside the breeding season it occurs on freshwater lakes, rivers, and sloughs, wet meadows, flooded fields, estuaries, shallow coastal waters, bays, and inlets. SPECIAL HABITAT REQUIREMENTS: Ponds or lakes in swampy muskeg flanked by short to medium conifers. NEST: Nests in dispersed colonies or noncolonially, from 4 to 20 feet above ground in branches or stumps of spruce, fir, and tamarack, and near water. May also nest in reeds, on mudflats of temporary potholes, and in clumps of bulrushes. FOOD: Forages in a variety of habitats associated with water, such as marshy ponds, freshwater marshes, rivers, lakes, estuaries, salt marshes, beaches, bays, and open ocean. Eats small fishes, insects, spiders, snails, crustaceans, and marine worms. REFERENCES: Bent 1921, Clapp et al. 1983, Cramp and Simmons 1983. 164 Ring-billed Gull Larus delawarensis S' -- L 16 ” ' RANGE: Breeds in the West from southern interior British Columbia and northeastern Alberta to north-central Manitoba south to northeastern California, south-central Colorado and northeastern South Dakota; in the East from north-central Ontario to southern Labrador south to eastern Wisconsin and northern Illinois across to central New Hampshire and New Brunswick. Winters along the Pacific Coast from southern British Columbia south to Mexico; in the interior from the Great Lakes to Mexico and the Gulf Coast; and along the Atlantic Coast from the Gulf of St. Lawrence to the Greater Antilles. STATUS: Common. HABITAT: From boreal regions to temperate prairies, inhabits small to moderately sized rocky islands and occasionally peninsulas in large freshwater lakes, rivers, or ponds (a few colonies are on oceanic islands or coasts). Usually avoids densely settled areas. Outside the breeding season, frequents harbors, refuse dumps, sewage outlets, reservoirs, lakes, ponds, streams, coastal bays, estuaries, beaches, and mudflats. Roosts on exposed sandbars and islands. SPECIAL HABITAT REQUIREMENTS: Islands and peninsulas covered with low vegetation 6 to 54 inches high. NEST: Frequently nests in mixed colonies with other Laridae, including herring and California gulls. Usually nests on the ground in flat, elevated, sparsely vegetated areas, but sometimes in low trees. FOOD: Forages in plowed fields, pastures, tidal flats in salt marshes, along the shore, along beaches, and in shallow waters. During the breeding season, feeds extensively on insects, grains, and small fishes. Also eats small rodents, earthworms, and refuse. REFERENCES: Clapp et al. 1983, Cramp and Simmons 1983, Jarvis and Southern 1976, Vermeer 1970. 165 California Gull Larus californicus LI 7' RANGE: Breeds from southern Mackenzie south to northern Utah and north-central Colorado, and west to southern interior British Columbia and northeastern California. Winters from southern Washington and eastern Idaho south, mostly along the Pacific Coast to Mexico. STATUS: Common. HABITAT: Inhabits barren islands on fresh, brackish, or alkaline lakes, shores of lakes or ponds, and marshes. Favors sites with low, sparse vegetation. Outside the breeding season it occurs on seacoasts, bays, estuaries, mudflats, irrigated fields, and other agricultural lands. SPECIAL HABITAT REQUIREMENTS: Open sandy or gravelly lakeshores or islands. NEST: A colonial nester, often nesting in mixed colonies with the ring¬ billed gull. Avoids dense herbaceous cover, and constructs its nest in a scrape on the ground in elevated, boulder-strewn areas. FOOD: Forages opportunistically on shortgrass plains and cultivated lands for a wide variety of foods, including insects, carrion and garbage, earthworms, young birds, bird eggs, and rodents. REFERENCES: Greenhalgh 1952, Johnsgard 1979, Vermeer 1970. 166 Herring Gull Larus argentatus RANGE: Breeds from northern Alaska and northern Yukon to central Keewatin, western Baffin Island, and northern Labrador south to south- central British Columbia, central Alberta, northern Minnesota, northeastern Illinois, northern Ohio, northern New York, and along the Atlantic Coast to northeastern South Carolina. Winters from southern Alaska, the Great Lakes region, and Newfoundland south, mostly at sea and along coasts, large rivers and lakes, to Panama. STATUS: Abundant. HABITAT: Uses a wide variety of habitats, including sandy, rocky, or wooded islands, stabilized sand dunes, margins of tundra lakes, Spartina marshes, cliffs, grass meadows, and buildings. In winter, occurs primarily along the shore of the ocean or other bodies of water, concentrating on beaches and in areas where food is likely to be abundant. SPECIAL HABITAT REQUIREMENTS: Nesting sites must be free of terrestrial predators and within 25 miles of a dependable source of food. NEST: Usually nests in exposed sites on the ground in small to large colonies, but occasionally in trees. Prefers to nest in low sites; depending on habitat may nest at the base of boulders, stumps, or bushes on grassy slopes, near large, tall clumps of vegetation, on drift adjacent to salt marshes, or on rock or grassy substrates. FOOD: Feeds opportunistically in garbage dumps, around seafood¬ processing operations, in pastures and cultivated fields, on lawns, tundra, and beaches, and at sea. Consumes largely animal matter, including small mammals, birds and bird eggs, amphibians, fishes, shellfishes, and a great variety of invertebrates, plus berries and some fruit, as well as carrion and, at times, garbage. REFERENCES: Burger and Shisler 1978, Clapp et al. 1983, Cramp and Simmons 1983, Forbush and May 1955. 167 Caspian Tern Sterna caspia A L20 RANGE: Breeds locally in the West from coastal and eastern Washington, eastern Oregon, northern Utah and northwestern Wyoming south to southern California and western Nevada; in the interior from southern Mackenzie to southern James Bay south to North Dakota, northeastern Illinois, and southern Ontario; at scattered localities along the Atlantic Coast from Newfoundland to South Carolina; and along the Gulf Coast from Texas east to Florida. Nonbreeding birds often summer in the James Bay and Great Lakes region, and along both coasts of the United States. Winters primarily in coastal areas from California and North Carolina south to Mexico. STATUS: Common. HABITAT: Usually found near the coastline on sandy, stony, or shell beaches, barrier or spoil islands, islands with sand-gravel substrate with little or no vegetation, or on a shell berm in a salt marsh. Tends to occupy less-developed and less-polluted segments of the coast, but is also found inland along shorelines of large lakes. Wintering terns generally are found along beaches, and on isolated spits, often roosting with other larids. In migration, occurs along water courses or in large marshes. SPECIAL HABITAT REQUIREMENTS: Sparsely vegetated islets or shorelines. NEST: Usually found in compact colonies, but occasionally nests singly in the vicinity of other tern species in shallow depressions in the ground on bare sandy or rocky soil. FOOD: Feeds almost entirely on fish 3 to 10 inches long, foraging on species and sizes that are most readily available; also takes crayfish, insects, nestlings and eggs of other birds, and rarely, carrion. REFERENCES: Clapp et al. 1983, Johnsgard 1979, Ludwig 1965. 168 Common Tern Sterna hirundo RANGE: Breeds from south-central Mackenzie to southern Quebec and southern Labrador south to eastern Washington and southeastern Alberta, across to central Minnesota, northern Ohio, and northwestern Vermont, and locally along the Atlantic Coast to North Carolina; locally on the Gulf Coast in Texas, Mississippi, and western Florida. Winters in South America, rarely along the coasts of southern California, South Carolina, Florida, and the Gulf Coast. STATUS: Common; of special concern on the blue list; declining on East Coast. HABITAT: Uses a variety of habitats, mainly near water, often on islets, and usually in areas with little or no vegetation. Inhabits sparsely vegetated sandy islands, barrier beaches, marshy islands, small islands in salt marshes, or low, small, rocky islands in lakes and rivers. After nesting, typically found along shorelines, on exposed rocks and old pilings, and inshore over shallow coastal waters. SPECIAL HABITAT REQUIREMENTS: Nesting areas with scant vegetation, isolated from disturbance and predation, and in close proximity to a source of food. NEST: Builds nests in colonies; nests may vary from a slight hollow in sand or among pebbles to a well-built hollowed mound of grasses and seaweeds, may be in the open or near weeds, grasses, or bushes. Generally prefers sparse cover around the nest. FOOD: Forages in shallow waters, margins of lakes, or along the coast, but tends to avoid muddy waters. Consumes a diet that varies with locality, including fish less than 6 inches, shrimp and other crustaceans, aquatic worms, insects, and some waste material. REFERENCES: Burger and Lesser 1978, Forbush and May 1955, Johnsgard 1979, Palmer 1941, Tate and Tate 1982. 169 Forster’s Tern Sterna forsteri C — LI 4” RANGE: Breeds from southeastern British Columbia and central Alberta to central Manitoba, south to southern California and south-central Idaho, across to central Kansas, northern Iowa, and northwestern Indiana; along the Atlantic Coast from southern New York south locally to North Carolina; and along the Gulf Coast from Texas east to Louisiana. Winters along the Pacific and Atlantic Coasts from central California and Virginia south to Central America. STATUS: Common. HABITAT: Primarily inhabits large saltwater and freshwater marshes; also found on marshy bays, marshy parts of islands, marshy edges of streams and lakes, sloughs, dikes in evaporation ponds, estuarine islands, marshes adjacent to barrier beaches, and dredge-spoil islets. In winter, occurs in harbors, marshy bays, estuaries, lagoons, and inlets along coastal areas, occasionally occurring inland along lakes and ponds. SPECIAL HABITAT REQUIREMENTS: Extensive marshy areas with vegetated nest sites partly open to water. NEST: Usually found in small colonies, nesting on mats of floating dead vegetation, flattened reeds and cattails, large muskrat houses near the edges of open pools of water, floating rootstalks of cattails, or sometimes in a shallow depression in sand or mud. May also locate nests on sand or gravel bars, beaches, or grassy islands. Sometimes uses old or abandoned nests of western and pied-billed grebes. FOOD: Feeds over or near the marshes in which it nests. Eats small fish, insects, crustaceans, and frogs. REFERENCES: Bergman et al. 1970, Clapp et al. 1983, Forbush and May 1955, Johnsgard 1979, Low and Mansell 1983, McNicholl 1971. 170 Least Tern Sterna antillarum L8'/2” RANGE: Breeds along the Pacific Coast from central California south to southern Baja California, inland along the Colorado, Red, Missouri, and Mississippi River systems from southern South Dakota, western Iowa, Southwestern Missouri, northwestern Indiana, central Louisiana, northeastern Texas, central Oklahoma, western Kansas, and central New Mexico, and along the Atlantic Coast from Maine to Florida and west to Texas. Winters along the Pacific Coast from Baja California and along the Gulf Coast to South America. STATUS: Threatened in the Great Plains; endangered along California Coast; stable along Atlantic Coast, but is listed on New Jersey and Maine’s threatened and endangered lists. HABITAT: Inhabits river sandbars, inland islands, broad areas of sand or gravel beaches, and newly cleared land along the coast. Frequents salt plains in Oklahoma. SPECIAL HABITAT REQUIREMENTS: Open, sandy coastal beaches, and river sandbars for nesting. NEST: Nests solitarily or in scattered colonies. Nests in scrapes (with little or no lining) in sand or gravel (gravel or pebble substrates are preferred) above ordinary tides. Shares habitat with piping plover in the Great Plains. FOOD: Skims the surface of the water or hovers and dives for food, sometimes spears fish with closed bill. Eats sand eels, shrimp, and small fish. REFERENCES: Bent 1921, Tompkins 1959, Johnsgard 1979, Terres 1980. 171 Black Tern Chlidonias niger RANGE: Breeds from southwestern and east-central British Columbia and south-central Mackenzie to southern Quebec and southern New Brunswick, south locally to south-central California and northern Utah, across to Nebraska, south-central Illinois, Pennsylvania, and Maine. Nonbreeding birds occur in summer south on the Pacific Coast to Panama, and in eastern North America to the Gulf Coast. Winters in South America. STATUS: Common; overall population is stable or decreasing slightly. HABITAT: Found in taiga and on the plains and prairies, where it inhabits shallow marshes, open areas of deeper marshes, reed-bordered sloughs, natural ponds, lakes, fish and stock ponds, shallow river impoundments, wet meadows, river oxbows, ditches, edges of streams, and swampy grasslands. In migration, frequents freshwater and saltwater, occurring along the coast and along marshes, rivers, lakes and nearby cultivated fields. SPECIAL HABITAT REQUIREMENTS: Aquatic habitats with extensive stands of emergent vegetation and large areas of open water. NEST: Often nests in small colonies, but occasionally nests singly. Usually places nest on a floating mass of vegetation such as cattails and bulrushes and anchored to surrounding vegetation, on floating pieces of wood, or in a slight hollow atop a muskrat house. Prefers areas of emergent vegetation over water up to 3 feet deep or near open water. Sometimes uses abandoned nests of other birds including grebes, Forster’s terns, and American coots. FOOD: Does not compete strongly with fish-eating species; consumes a diet that includes aquatic and land insects, worms, small mollusks, crustaceans, and a few small fishes and grubs. REFERENCES: Clapp et al. 1983, Forbush and May 1955, Johnsgard 1979, Low and Mansell 1983, McNicholl 1971, Tate and Tate 1982. 172 Marbled Murrelet Brachyramphus marmoratus summer winter L 8" RANGE: Breeds from Alaska south to central California. Winters, off shore, from southern Alaska to central California (casually to southern California). STATUS: Locally common. HABITAT: Inhabits coniferous forests, coastal islands, and inland lakes, usually within 12 miles from ocean. SPECIAL HABITAT REQUIREMENTS: Old growth coniferous forests. NEST: Nests solitarily on bare rocks, below ledges, in rock cavities, on the ground, and in coniferous trees. Usually nests within 4 miles of the ocean but occasionally farther inland. FOOD: Dives for food and consumes mostly small fish and various crustaceans but diet varies regionally. REFERENCES: Terres 1980, Day et al. 1983, Carter and Sealy 1986. 173 Rock Dove Columba livia RANGE: Resident from southern Alaska and southern Canada south throughout North America. Introduced; originally an Eurasian species. STATUS: Common. HABITAT: May inhabit narrow, steep-walled canyons and rocky cliffs, but far more commonly found near human habitations. NEST: Prefers to nest on or in structures that provide narrow ledges similar to cliff ledges. Constructs a flimsy nest on stone, brick, and concrete buildings in cities, high upon ledges, under windows, under bridges, on monuments, in barns or other man-made structures, preferably in semi-dark cavities. May nest singly or in colonies. FOOD: Commonly feeds on parkland, sidewalks, and parking lots in cities; cultivated fields; feedlots; and wastelands. Gleans seeds of weeds, grasses, and grains, takes human handouts, and eats a few berries and tender roots of grasses. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Terres 1980. 174 White-crowned Pigeon Columba leucocephala RANGE: Breeds in southern Florida on islets in the Florida Keys, throughout Florida Bay, the Lesser Antilles, and islets of Central America. Nonbreeding birds may occur in summer in southern peninsular Florida. Winters from southern peninsular Florida and the Florida Keys throughout most of the breeding range to central America. STATUS: Common. HABITAT: Generally gregarious: breeds and roosts in large concentrations on brushy, small, low islands and keys, among coastal mangroves and pines. NEST: Nests colonially, but not with or near other colonial species. Builds nest from a few feet to 15 feet or more above ground, on top of cactus or bushes, or high in mangroves; occasionally low and over water. FOOD: Forages in open forests, woodlands, and scrubs and eats seeds, berries, and a few insects. REFERENCES: Cottam and Knappen 1939, Harrison 1975, Sykes in Farrand 1983b, Terres 1980, Wiley and Wiley 1979. 175 Red-billed Pigeon Columba flavirostris RANGE: Breeds in the lower and middle Rio Grande Valley in southern Texas south through Mexico into Central America; mostly absent in winter. STATUS: Local and uncommon. HABITAT: Inhabits semiarid woodlands near water. In Texas, it is found in river thickets containing tall timber and a thick undergrowth of thorny shrubs including ebony blackbead, huisache, mesquite, baldcypress, great leucaena, Mexican ash, elm, black willow, and hackberry. Often perches on exposed bare branches high in trees. SPECIAL HABITAT REQUIREMENTS: Tall dense brush with small patches of tall trees. NEST: Nests 8 to 30 feet above ground on a horizontal tree branch, in a clump of small branches, or in a tree concealed by a tangle of vines or brush. FOOD: Forages high in the crowns of trees and in stubble fields on a diet that includes fruits of hackberries, condalia, and wild grape; acorns; and waste grains. REFERENCES: Bent 1932, Oberholser 1974a, Terres 1980. 176 Band-tailed Pigeon Columba fasciata L13” RANGE: Breeds from southeastern Alaska and southwestern British Columbia south through the mountains of Washington, Oregon, California, and extreme western Nevada to Baja California: and from southern Nevada, Arizona, central Utah, north-central Colorado, New Mexico, and western Texas south to Honduras. Winters from central California, central Arizona, and western Texas south to Honduras. STATUS: Locally common. HABITAT: Along the Pacific Coast, inhabits a variety of forest lands with western hemlock, western redcedar, and Douglas-fir to ponderosa pine, white fir, or incense cedar. In Oregon and Washington, prefers forest land with a good interspersion of serai stages and openings; in California, prefers forests, woodlands, or chaparral with an abundance of oak. In the interior, occupies habitats ranging from montane oak woodlands to arid woodlands of pinyon pine and oaks, and from agricultural areas near forests to berry-producing areas at 11,000 feet elevation. Occasionally found in spruce-fir associations characterized by Engelmann spruce, subalpine fir, lodgepole pine, limber pine, and aspen, but prefers sites dominated by ponderosa pine and Gambel oak. SPECIAL HABITAT REQUIREMENTS: Mature conifers or broad leaved trees at least 8 feet tall for nesting and a source of mineral water or salt deposits in early fall and winter. NEST: Nests in coniferous or deciduous trees usually located near a clearing and on a moderate to steep slope or precipice. Conceals nest on horizontal branches typically 15 to 40 feet high, rarely on the ground. FOOD: Gleans sometimes exclusively on one species or source of food as long as the supply lasts. Mainly eats mast and berries, but also eats acorns, pine nuts, pinyon nuts, blossoms, green and ripe fruits, and some waste grains. REFERENCES: Jeffrey 1977, Johnsgard 1975a, Neff 1947, Peeters 1962. 177 White-winged Dove Zenaida asiatica RANGE: Breeds from southeastern California and southern Nevada to southwestern Texas south, through Central America into western South America. Introduced and established in southern Florida. Winters generally in the breeding range, but northern birds are mostly migratory, casually ranging north to northern California and Colorado and occurring regularly along the Gulf Coast east to Florida. STATUS: Locally abundant. HABITAT: Occupies a variety of habitats in semiarid woodlands. Prefers tall, dense, or brushy riparian woodlands, with trees from 15 to 25 feet tall and an understory of thorny shrubs. Also occupies desert scrub, desert grassland, oak woodland, chaparral, valleys of desert mountains, and shade and fruit trees of agricultural areas, country roadsides, and suburban residential areas. SPECIAL HABITAT REQUIREMENTS: Nest sites with trees of low to medium height having dense foliage and fairly open ground cover. NEST: Builds nests on relatively horizontal branches of a variety of trees and large shrubs in shaded sites, preferably in the interior of woodlands and thickets. FOOD: Forages in shelter of thickets, weeds, or fields for seeds. Also eats mast, fruit, and some insects. REFERENCES: Brown 1977, Cottam and Trefethen 1968, Neff 1940, Oberholser 1974a. 178 Mourning Dove Zenaida macroura RANGE: Breeds from southern and central Alberta to southern New Brunswick and Nova Scotia south to Mexico. Winters primarily from northern California east across the central United States to Iowa, southern Michigan, southern Ontario, New York, and New England south throughout the breeding range to central Panama. STATUS: Abundant. HABITAT: Occupies a broad range of habitats from desert areas close to water to a variety of wooded habitats, but avoids densely forested regions. Inhabits open country, especially fields, orchards, or generally weedy areas having an abundance of grains and seeds. Also inhabits open mixed woodlands and wood edges, shelterbelts, church and cemetery sites, evergreen plantations, suburbs, and cities. SPECIAL HABITAT REQUIREMENTS: Open country with some bare ground and adequate foods. NEST: Loosely colonial or solitary. Generally nests on horizontal branches in shrubs and trees, especially conifers up to 100 feet, but typically 10 to 25 feet above ground, and occasionally on the ground. FOOD: Feeds primarily on weed seeds and waste grains. Also eats a few insects, small snails, fruits, and nuts. REFERENCES: Davison and Sullivan 1963, DeGraff et al. 1980, Forbush and May 1955, Hanson and Kosack 1963, Johnsgard 1975a, Keeler 1977. 179 Inca Dove Columbina inca RANGE: Resident from extreme southeastern California, central Arizona, southern New Mexico, and central Texas south to Costa Rica. STATUS: Common to abundant. HABITAT: Primarily found in the vicinity of human habitations, especially around introduced broad-leaved deciduous trees, exotic conifers, and native live oaks in areas with little or no understory. Inhabits poultry and livestock feedlots, yards, gardens, orchards, school grounds, city parks, and roads through brushy mesquite pastures, usually near a source of water. Roosts in evergreen trees. NEST: Nests on a horizontal fork or flattened tree limb or in a bush, 4 to 25 feet above ground in a brushy pasture, or more often in the immediate vicinity of houses. Builds nests in native shrubs (including catclaw and chollas), in baldcypress, in shade trees, on top of utility poles, in hanging baskets near a house, and sometimes in nests of other species including mourning doves, mockingbirds, and cactus wrens. Generally uses nests for 2 or more consecutive nestings. FOOD: Feeds almost entirely on the ground, taking seeds of a wide variety of native plants. Eats wheat, cracked corn, oats, and milo readily if available. REFERENCES: Anderson and Anderson 1948, Harrison 1979, Johnston 1960, Oberholser 1974a. 180 Common Ground-Dove Columbina passerina (formerly Ground Dove) L5‘/ 2 ” RANGE: Resident from southern California, central Arizona, southern New Mexico, central Texas, the Gulf Coast, and South Carolina south to South America. STATUS: Common, locally abundant. HABITAT: Inhabits sparsely wooded areas with low undergrowth, roadsides, fields, orchards, sandy reefs, and open sandy areas in forest and savannah; over much of its range, is now primarily around farms and towns. In Texas, occurs in grassy mesquite-live oak-cactus savannah and to some extent scrubby juniper-oak areas; in Georgia and South Carolina, occurs mainly near beaches and sea islands with tall beach grass. In Arizona, inhabits river bottomlands with mesquite or tamarisk woods. SPECIAL HABITAT REQUIREMENTS: Open areas with plants that produce small seeds. NEST: Uses a wide range of sites for nesting. May nest in a slight hollow on the ground, in a low bush, or in a tree, up to 25 feet above the ground. FOOD: Feeds primarily on small seeds gathered from gardens and lawns, along roadsides, in fields, weed patches, or grassy areas. Also takes a few berries and some insects. REFERENCES: Bent 1932, Forbush and May 1955, Goodwin 1967, Oberholser 1974a, Phillips et al. 1964. 181 Black-billed Cuckoo Coccyzus erythropthalmus RANGE: Breeds from east-central and southeastern Alberta and southern Saskatchewan to New Brunswick and Nova Scotia south, at least locally, to eastern Colorado, north-central Texas, northern Arkansas, northern Alabama, and the Carolinas. Winters in South America. STATUS: Common. HABITAT: Prefers extensive areas of upland woods that provide a variety of trees, bushes, and vines. Also occurs in brushy pastures, hedgerows, open woodlands, orchards, thickets, and along wooded roadsides. SPECIAL HABITAT REQUIREMENTS: Low, dense, shrubby vegetation. NEST: Selects nest sites that are well concealed by overhanging branches and leaf clusters. Usually nests in shrubs or on a low tree branch, typically 4 to 6 feet above ground. Occasionally lays eggs in the nests of other birds. FOOD: Feeds primarily on caterpillars, especially tent caterpillars; also other insects, spiders, and a few tiny mollusks, fishes, and wild fruits and berries. REFERENCES: Beal 1904, Bent 1940a, DeGraff et al. 1980, Herrick 1910, Johnsgard 1979, Spencer 1943. 182 Yellow-billed Cuckoo Coccyzus americanus RANGE: Breeds from interior California and northern Utah to southwestern Quebec and southern New Brunswick, south to southern Arizona and into Mexico. Winters in South America. STATUS: Common. HABITAT: Favors moderately dense thickets near watercourses, second- growth woodlands, deserted farmlands overgrown with shrubs and brush, and brushy orchards for habitat. Also inhabits open woods, avoiding extremely dense woods and high elevations. SPECIAL HABITAT REQUIREMENTS: Low, dense, shrubby vegetation. NEST: Prefers to nest in thick bushes overgrown with vines or in trees on horizontal limbs, typically 4 to 8 feet above ground. Conceals nest with surrounding foliage. FOOD: Forages among leaves for food, which consists mainly of caterpillars, especially tent caterpillars and fall webworms. Also eats other insects and spiders, some small wild fruits, some frogs, and occasionally a small lizard. REFERENCES: Beal 1904, Bent 1940a, DeGraff et al. 1980, Johnsgard 1979, Preble 1957, Tate and Tate 1982. 183 Mangrove Cuckoo Coccyzus minor RANGE: Breeds in southern Florida from Tampa Bay and Miami southward in coastal areas, including the Florida Keys through Mexico to South America. Winters throughout the breeding range, but mostly south of Florida. STATUS: Rare and local. HABITAT: Inhabits red and black mangrove thickets and swamps near saltwater, and upland hardwood hummocks. NEST: Typically builds nest on a horizontal mangrove branch, that is indistinguishable from the nest of the yellow-billed cuckoo. FOOD: Often feeds in open fields and clearings near thickets on caterpillars and other insects, spiders, and a few small fruits and wild berries. REFERENCES: Bent 1940a, Harrison 1975, Sykes in Farrand 1983b, Terres 1980. 184 L 22" Greater Roadrunner Geococcyx californianus (formerly Roadrunner) RANGE: Resident from northern California (rarely) and western and central Nevada east to southern Kansas and north-central Louisiana south to Mexico. STATUS: Common. HABITAT: Typically associated with desert regions, but also found in chaparral, grasslands, open woodlands of pine and oak, agricultural areas, and moist woodlands. Frequents edge habitats provided by a mixture of open land, brush, and forest; also at home among tall pines and magnolias or mesquite and cactus. SPECIAL HABITAT REQUIREMENTS: Bare areas with some scattered trees and bushes. NEST: Usually nests in a low tree, bush, thicket, or clump of cactus from 3 to 15 feet above ground, rarely on the ground. In desert regions, builds nest so that bands of shade cross the nest during the extreme heat of the day; prefers staghorn cholla for nesting site in arid regions. FOOD: Feeds primarily on animals, with about 10 percent of the diet consisting of vegetable matter. Eats a great variety of insects, spiders, scorpions, snails, lizards, and small snakes; also eats some young birds and small mammals, and fruits and seeds. REFERENCES: Bryant 1916, Johnsgard 1979, Oberholser 1974a, Ohmart 1973, Terrill in Farrand 1983b. 185 Smooth-billed Ani Crotophaga ani L RANGE: Resident in central and southern Florida from Tampa Bay and Merritt Island region southward, especially from Lake Okeechobee area to Dade County into Central and South America. STATUS: Locally common. HABITAT: Inhabits dense brush or hedgerows near fields, pastures, and marshes, roadside thickets, gardens and thick woods in cities, towns, and around farms. NEST: May nest singly or communally, with several females laying their eggs in a single nest. The nest is constructed in trees or dense shrubs 6 to 30 feet above ground, commonly near human habitation. FOOD: Forages in moist open meadows for grasshoppers, crickets, moths, green tree frogs, and chameleons. REFERENCES: Harrison 1975, Kaufman in Farrand 1983b, Merritt 1951. 186 Groove-billed Ani Crotophaga sulcirostris L 14" RANGE: Resident in central and southern Texas and southern Louisiana south through Mexico and Central America to South America. STATUS: Common to uncommon. HABITAT: Tolerant of a wide range of ecological conditions, inhabiting brushy pastures, orchards, light second growth, lawns and clearings near homes, marshes, and moist thickets along rivers. Also found in semi-desert areas with scattered cacti and acacias and thickets of ebony blackbead, mesquite, and Jerusalem-thorn. Frequently accompanies livestock, which stir insects from the grass. SPECIAL HABITAT REQUIREMENTS: Open areas with low vegetation for foraging, and trees and bushes for roosting and nesting. NEST: Builds nest from 2 to 25 feet up in densely foliated trees or, in arid regions, in opuntias and other cacti. May nest singly or communally, with several females laying their eggs in a single nest. FOOD: Forages in pastures and farm fields, feeding on insects stirred up by cattle. Eats mainly insects, including grasshoppers, termites, and roaches, plus spiders, small lizards, and berries. REFERENCES: Kaufman in Farrand 1983b, Oberholser 1974a, Skutch 1959, Terres 1980. 187 Common Barn-Owl Tyto alba (formerly Barn Owl) L14" RANGE: Resident from southwestern British Columbia, southern Idaho, and Montana east to southern Vermont and Massachusetts south through the United States to South America. Northernmost populations are partially migratory, wintering south to southern Mexico and the West Indies. STATUS: Uncommon; overall population level is low, but stable. HABITAT: Found in open to semiopen habitats such as prairie, farmland, savannah, marshland, and desert, but prefers the vicinity of farms and towns. Avoids woodlands and higher elevations. SPECIAL HABITAT REQUIREMENTS: Abundant supply of small mammals for food, and hollow trees, old buildings, barns, cavities, or caves for nesting and roosting. NEST: Nests in a variety of sites. Favors natural tree hollows, especially in live oaks near a marshy meadow. Typically nests in old barns, church and school steeples, silos, or abandoned buildings. Also uses protected ledges along cliff faces, abandoned underground burrows of badgers, woodchucks, or other mammals, caves, cavities in high stream banks (8 to 10 feet above water level), abandoned nests of crows, hawks, or magpies, and artificial nest sites. Will return to the same nest site year after year if undisturbed. FOOD: Flunts by night over marshes, meadows, fields, barnyards, brushy areas, pastures, and other open areas for small mammals, especially mice, and occasionally small birds and large insects. Also eats some frogs, snakes, lizards, and crayfish. REFERENCES: Coats in Farrand 1983b, DeGraff et al. 1980, Hawbecker 1945, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Tate and Tate 1982. 188 Flammulated Owl Otus flammeolus RANGE: Breeds locally from southern British Columbia, southern Idaho, and northern Colorado south to southern California, southern Arizona, southern New Mexico, western Texas and Mexico. Winters in Mexico, casually north to southern California. STATUS: Rare to locally common. HABITAT: Inhabits forests of the western mountains, mostly from 4,500 to 7,800 feet but as high as 10,000 feet elevation. Prefers woods with dense, thicketlike cover close to relatively open areas. Favors ponderosa pine forests but also occurs in forests of spruce-fir, Douglas-fir, lodgepole pine, aspen, and pinyon-juniper. SPECIAL HABITAT REQUIREMENTS: Some undergrowth or inter¬ mixture of oaks in the forest. NEST: Usually nests in abandoned flicker or other woodpecker nest cavities from 7 to 25 feet above ground in aspen, oaks, or pines. Will forceably evict a flicker if an abandoned cavity is not available; rarely nests in holes constructed by bank swallows. FOOD: Consumes a diet of insects and other invertebrates such as spiders, scorpions, and centipedes; prefers moths, beetles, crickets, and grasshoppers and will sometimes eat small birds and small mammals. REFERENCES: Coats in Farrand 1983b, Heintzelman 1979, Karalus and Eckert 1974, Oberholser 1974a, Phillips et al. 1964. 189 L8” red phase Eastern Screech-Owl Otus asio Western Screech-Owl Otus kennicottii (formerly Screech Owl) 1 RANGE: Eastern Screech-Owl; Resident from southern Manitoba and northern Minnesota to southwestern Quebec and Maine, south through the eastern United States, eastern Montana, eastern Colorado, and western Oklahoma to southern Texas, and across to southern Florida. Western Screech-Owl; Resident from south-coastal and southeastern Alaska, coastal and southern British Columbia, and northern Idaho to southeastern Colorado and extreme western Oklahoma, south to Mexico and western Texas. STATUS: Common; populations are declining in the West, but are stable in the East. HABITAT: Found in a variety of habitats, favors oak and riparian woodlands in the West, and open woodlands adjacent to meadows, marshes, or fields in the East. Also inhabits orchards, shade trees in towns and cities, small woodlots, and deciduous forests. Prefers areas with widely spaced trees interspersed with grassy open spaces for hunting. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting and roosting in trees with a minimum dbh of 12 inches. NEST: Nests in natural cavities in trees or in old woodpecker holes, especially those of the northern flicker and pileated woodpecker. Chooses cavities with openings 3 to 5 inches in diameter that are typically 5 to 30 feet (but up to 50 feet) above the ground. Many use same cavity for many years; will use artificial cavities. FOOD: Hunts for its food in grassy openings, fields, meadows, or along wooded field margins or streams. Primarily takes rodents, especially meadow voles, but also eats insects, scorpions, spiders, centipedes, crayfish, amphibians, reptiles, fishes, and small birds. REFERENCES: DeGraff et al. 1980, Earhart and Johnson 1970, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Scott et al. 1977, Tate and Tate 1982, Thomas et al. 1979, Van Camp and Henny1975. 190 Whiskered Screech-Owl Otus trichopsis (formerly Whiskered Owl) L6!4” RANGE: Resident from southeastern Arizona south to Nicaragua. STATUS: Common. HABITAT: Inhabits scattered to dense woodlands on the slopes of valleys and in canyons, from 4,000 to 7,000 feet (usually between 5,500 and 6,500) feet in elevation. Occurs in dense oak, oak-pine, and sycamore woodlands, avoiding forests of pure pines or firs. SPECIAL HABITAT REQUIREMENTS: Cavities in trees for nesting. NEST: Nests in natural cavities in trees or in abandoned nest cavities of northern flicker and other woodpeckers. Prefers cavities in large branches or stubs to cavities in tree trunks. Favors relatively deep holes with bottoms 14 to 16 inches below the cavity entrance. Usually chooses a nest site that is 10 to 20 feet above the ground, generally in white oaks, but also uses other species. FOOD: Consumes a diet mostly of insects and other invertebrates, including crickets, large moths, beetles, grasshoppers, large caterpillars, scorpions, centipedes, some small mammals, and rarely, small birds. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974. 191 Great Horned Owl Bubo virginianus RANGE: Breeds from western and central Alaska and central Yukon to Labrador and Newfoundland, south throughout the Americas to Tierra del Fuego. Winters generally throughout the breeding range, with the northernmost populations being partially migratory. STATUS: Common. HABITAT: Occurs in a wide variety of forested habitats. Inhabits open coniferous, deciduous, or mixed woodlands, orchards, second-growth forests, marshes, swamps, riverine forests, partially wooded slopes, brushy hillsides, farm woodlots, large city parks, and rocky canyons well away from forest cover. In the South, prefers baldcypress hummocks and expansive dense palmetto woodlands interspersed with scattered pine. SPECIAL HABITAT REQUIREMENTS: Large abandoned bird nests or large cavities for nesting. NEST: Uses a wide variety of nest sites up to 70 feet aboveground; frequently abandoned nests of hawks, herons, or crows. Also uses large tree cavities, crotches, stumps, caves, and ledges. Occasionally, constructs a nest but most often uses abandoned nests. FOOD: Prefers open areas near woodlands such as marshes or meadows for hunting. Consumes an extremely varied diet; will attempt to kill animals up to the size of a turkey or porcupine, but prefers small to medium-sized mammals and birds. Also preys upon reptiles, amphibians, large insects, and fishes; rarely consumes carrion. REFERENCES: DeGraff et al. 1980, Earhart and Johnson 1970, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Sprunt 1955. 192 Snowy Owl Nyctea scandiaca L20” RANGE: Breeds from northern Alaska and northern Yukon to Prince Patrick and northern Ellesmere islands, south to coastal western Alaska, northern Mackenzie and southern Keewatin across to northern Quebec and northern Labrador. Winters irregularly south to southern Canada, Minnesota, and New York, and occasionally further south. STATUS: Common in the far North, rare and local in the conterminous United States. HABITAT: Found north of the tree line in arctic tundra, where it inhabits moss- and lichen-covered flatlands, lowlands, and valleys well dotted with frost-heave mounds, hillocks, or rocks. Avoids very marshy areas with no raised sites. During years of low lemming abundance, moves southward into the conterminous United States, onto open fields, sandy beaches, barrier islands, and marshes. SPECIAL HABITAT REQUIREMENTS: Raised sites such as frost-heave mounds several yards in diameter and as much as 2 to 3 feet above the ground, large rocks, or other abrupt rises in the ground for roosting and nesting. NEST: Generally nests at elevations less than 650 feet. Nests in a shallow depression on top of a frost-heave mound or other raised site, occasionally on a gravel bank, tidal flat, or on slopes above a marsh or lake. Rarely, nests in an abandoned eagle nest high in a tree. FOOD: Kills a wide variety of mammals and birds, especially lemmings and mice. Throughout its summer and winter range, also preys on a variety of rodents, rabbits, waterfowl, shorebirds, songbirds, fish—almost any small animal that can be caught. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974, Udvardy 1977, Watson 1957. 193 Northern Hawk-Owl Surnia ulula L14” RANGE: Breeds from tree-line in western and central Alaska to southern Keewatin, central Labrador south to southern Alaska, northern Minnesota, northern Michigan, and New Brunswick. Winters from the breeding range southward irregularly to southern Canada and northern Minnesota, and casually the northern United States. HABITAT: Inhabits much of the northern open coniferous or mixed forests, forest edges, clearings, old burns, dense brushy areas (especially tamarack), scrubby second-growth woodlands, and muskeg. SPECIAL HABITAT REQUIREMENTS: Open woodlands with cavities for nesting. NEST: Usually nests in natural cavities or in enlarged holes of pileated woodpeckers and flickers, but also in birch, spruce, and poplar snags. Occasionally nests on cliffs or in crow’s nests. FOOD: Hunts extensively during the day, preying on small mammals (especially mice), birds, and insects. In summer, consumes primarily small mammals and insects. Preys on ptarmigan during winter when other foods are scarce. REFERENCES: Bent 1938, Henderson 1919, 1925, Mendall 1944. 194 Northern Pygmy-Owl Glaucidium gnoma (formerly Pygmy-Owl) L6” RANGE: Resident from southeastern Alaska, British Columbia, southwestern Alberta, and western Montana south, mostly in mountainous regions through southern California and interior Mexico to Central America, and extending east as far as central Colorado, central New Mexico, and extreme western Texas. STATUS: Common. HABITAT: Inhabits deciduous, coniferous, and mixed forests in the West. In Arizona, prefers mixed oak-pine forests on south-facing slopes from 4,000 to 13,000 feet in elevation, but tends to frequent coniferous forests at the higher elevations; in California, occurs up to 6,000 feet, primarily in mixed scattered hardwoods and conifers. In the Rocky Mountains, occurs from 5,000 to 10,000 feet in dense pine forests or open areas with scattered trees; along the Pacific Coast, prefers dense damp forests of firs, redwoods, and cedars. SPECIAL HABITAT REQUIREMENTS: Natural cavities or old woodpecker holes for nesting. NEST: Nests in abandoned cavities of the hairy woodpecker or northern flicker or natural cavities. Chooses cavities from 8 to 100 feet above the ground; in Rocky Mountains uses cavities up to 24 feet above the ground; in California, from 40 to 75 feet, and along the coast from 50 to 60 feet (but up to 100 feet) above the ground. May use the same nest site for several years. FOOD: Primarily preys upon mice and large insects; also eats other small mammals, small birds, spiders, scorpions, centipedes, small snakes, lizards, and toads. REFERENCES: Earhart and Johnson 1970, Heintzelman 1979, Karalus and Eckert 1974. 195 Ferruginous Pygmy-Owl Glaucidium brasilianum (formerly Ferruginous Owl) RANGE: Resident from south-central Arizona and southern Texas south through Mexico to South America. STATUS: Rare and local. HABITAT: Inhabits wooded river bottoms of cottonwoods and mesquite, but prefers densely vegetated desert areas with saguaro and cholla cacti and mesquite thickets. It is found from sea level to 4,000 feet. SPECIAL HABITAT REQUIREMENTS: Abandoned woodpecker holes for nesting. NEST: Nests in abandoned woodpecker cavities from 10 to 40 feet above the ground in cottonwoods, mesquite, and most often in saguaro cactus. May use same nest sites year after year. FOOD: Hunts from a perch, preying primarily on crickets, other large insects, and lizards. Also eats scorpions, caterpillars, and small birds and mammals. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974, Oberholser 1974a, Terres 1980, Terrill in Farrand 1983b. 196 Elf Owl Micrathene whitneyi L5!4” RANGE: Breeds from southeastern California, extreme southern Nevada, central Arizona, southwestern New Mexico, and western and southern Texas south to Mexico. Winters in Mexico. STATUS: Common. HABITAT: Prefers arid, low elevation desert areas overgrown with cacti, mesquite, and creosote bush, or with agave, ocotillo, and cactus desert scrub. Also inhabits riparian cottonwood and willow groves; mesquite floodplains; walnut, sycamore, and oak woodlands; and juniper, pinyon pine, and oak woodlands up to 7,000 feet in elevation. Avoids pure stands of pine, but will inhabit almost every other type of dry, woody vegetation. SPECIAL HABITAT REQUIREMENTS: Abandoned woodpecker holes for nesting. NEST: Nests in abandoned woodpecker holes, especially in saguaro cactus but also in agave bloom stalks, tree stumps, cottonwoods, mesquite, sycamore, pines, walnut oaks, or willows growing on mesas and desert slopes and in canyons. Nests generally 10 to 30 feet above the ground. FOOD: Preys primarily on large insects but also eats scorpions and rarely lizards, snakes, and small birds. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974, Ligon 1968, Oberholser 1974a. 197 Burrowing Owl Athene cunicularia L8 RANGE: Breeds from southern interior British Columbia to southern Manitoba south through eastern Washington, central Oregon, and California to Baja California, east to western Minnesota, western Missouri, Oklahoma, eastern Louisiana, and south to Mexico and Central America; also in Florida. Winters throughout breeding range except for the northern portions in the Great Basin and Great Plains regions. STATUS: Locally common; populations declining due to widespread elimination of burrowing rodents, notably prairie dogs and ground squirrels. HABITAT: Prefers nonforested plains, grasslands, deserts, and sometimes open areas such as vacant lots near human habitations or airports. Largely depends on mammals that dig burrows in open areas with short vegetation for nesting, roosting, and for escape. Commonly perches on fence posts, bushes, utility wires, roadside billboards, and burrow mounds. SPECIAL HABITAT REQUIREMENTS: Burrows of colonial burrowing mammals, especially prairie dogs and ground squirrels in open spaces. NEST: In the West, often nests in colonies in abandoned burrows of prairie dogs, and ground squirrels; also nests in burrows of woodchucks, foxes, badgers, coyotes, and armadillos. In Florida, nests in gopher tortoise burrows. Can excavate own burrow but usually enlarges burrows started by mammals and uses same burrow for years if not disturbed. FOOD: Hunts by ground foraging, hovering, from a perch, or by flycatching. Primarily eats insects and small mammals, but also takes some birds, fishes, and frogs. REFERENCES: Butts 1973, Errington and Bennett 1935, Evans 1982, Heintzelman 1979, Karalus and Eckert 1974, Tate and Tate 1982, Terres 1980, Zarn 1974b. 198 Spotted Owl Strix occidentalis RANGE: Resident from southwestern British Columbia south through western Washington and western Oregon to southern California; also in the mountains of southern Utah, central Colorado, Arizona, New Mexico, and extreme western Texas south into northern Mexico. STATUS: Rare; habitat is limited due to harvesting of old-growth forests. The Northern Spotted Owl subspecies is federally listed as a threatened species. HABITAT: Inhabits dense coniferous forests with crown closures of at least 80 percent or mixed woodlands and deeply shaded canyons in coastal and mountainous areas of the West. In California, prefers dense fir or Douglas-fir forests on sheer, heavily wooded cliffsides or in narrow canyons, but also inhabits stream valleys well grown with oaks, sycamores, willows, cottonwoods, and alder tangles. In Southwest, favors forested mountain tablelands and canyons from 5,500 to 9,000 feet with dense aspen clumps and creek fringe maples. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting and at least 600 acres of dense, old-growth forest or deep, narrow, well-wooded canyons per pair. NEST: Generally nests in cool, shaded areas with well-developed understory and near water. Prefers natural cavities in the old-growth trees, especially Douglas-fir or oaks, with broken tops and infested with mistletoe. Also will nest in cliff cavities, cave floors, occasionally abandoned hawk or raven nests, and hollow logs on the ground. Rarely builds its own nest in the crotch of a tall tree. FOOD: Preys on a wide variety of animals, but mainly takes small mammals; also eats small birds and large insects. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974, Marshall 1942, Tate and Tate 1982, Zarn 1974c. 199 Barred Owl Strix varia RANGE: Resident from southern and eastern British Columbia, northern Washington, and extreme northwestern Montana east to central Saskatchewan, and from southern Manitoba and central Ontario to New Brunswick and Nova Scotia, south to central and southern Texas, the Gulf Coast, southern Florida, and northern Mexico. Northernmost populations are partially migratory. STATUS: Common to uncommon. HABITAT: Prefers dense woodlands bordering lakes, streams, swamps, marshes, or low meadows. Favors oak woodlands or mixed forests free of a dense understory but also inhabits deciduous, coniferous, and mixed forests. May also inhabit isolated woodlots with numerous mature trees. SPECIAL HABITAT REQUIREMENTS: Cool, damp lowlands with large cavity trees 20 inches dbh or greater for nesting. NEST: Typically nests in a large cavity in a dead tree; may nest in abandoned hawk, crow, or squirrel nests if cavities are scarce. Generally chooses tall, old trees with cavities at least 25 feet above the ground, and in the forest interior. May use the same nest site for many years. FOOD: Hunts for prey over open fields, clearings, and wetlands near woodlands. Feeds on a wide variety of animals, especially mice and other small mammals; also eats birds (from warblers to grouse and other species of owls), fishes, frogs, salamanders, lizards, snakes, crayfish, scorpions, snails, spiders, and large insects. REFERENCES: DeGraff et al. 1980, Dunstan and Sample 1972, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Nicholls and Warner 1972. 200 RANGE: Breeds from central Alaska and northern Yukon to northern Manitoba and northern Ontario, south locally to central California, northern Idaho, northwestern Wyoming, central Saskatchewan, northern Minnesota, and south-central Ontario. Winters generally through the breeding range, wandering south irregularly to the northern tier of States. STATUS: Locally common to rare. HABITAT: Inhabits dense coniferous forests in Canada, and montane coniferous forests of the western States. Usually prefers pine and fir forests, rarely straying far out onto tundra barrens and muskeg marshes. Nests in mature poplar woodlands, preferably near muskeg areas, well secluded from human activities, and in spruce stands with islands of tamarack. In winter, may inhabit forests, sparse woodland edges bordering open fields, weedy fields with posts or scattered low trees or bushes, or brackish tidal meadows. SPECIAL HABITAT REQUIREMENTS: Old hawk or crow nests high in trees. NEST: Does not build its own nest but uses old nests of goshawks, red¬ tailed hawks, other large hawks, crows, ravens, or artificial nests. Locates nests 10 to 100 feet high in tamarack, balsam poplar, aspen, and spruce trees. FOOD: Preys primarily on small mammals but also takes some birds up to the size of a grouse. REFERENCES: Godfrey 1967, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Nero 1980. 201 Long-eared Owl Asio otus RANGE: Breeds from northern Yukon, southwestern Mackenzie, northern Saskatchewan and Nova Scotia, south to northern Baja California, southern Arizona, western and central Texas, Arkansas, northern Ohio, western Virginia, and New England. Winters from southern Canada south to Baja California, Mexico, southern Texas, the Gulf Coast, and Georgia; casually to Florida. STATUS: Locally common. HABITAT: Often inhabits coniferous woodlands but also deciduous forests and forested areas near open country. Also will inhabit open or dense woodlands, parks, orchards, woodlots, wooded swamps, streams, and reservoir shorelines, even low-growing scrub if it is in the form of dense, tangled thickets. Occurs up to 10,000 feet. SPECIAL HABITAT REQUIREMENTS: Dense vegetation for nesting and roosting cover. NEST: Most often uses old nests of large birds such as crows, hawks, ravens, herons, or magpies, but will use squirrel nests and natural tree cavities. Usually locates nest 15 to 30 feet above the ground, but may nest on the ground or on ledges. Rarely, will construct own nest. FOOD: Forages over wooded and open country, preying primarily on mice and other small mammals. Also eats some bats, cottontails, small birds, frogs, small snakes, and insects. REFERENCES: Armstrong 1958, DeGraff et al. 1980, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974. 202 Short-eared Owl Asio flammeus RANGE: Breeds from northern Alaska and northern Yukon to northern Quebec and Labrador, south to central California, northern Nevada, Utah, Kansas, Missouri, northern Ohio, northern Virginia, and New Jersey. Winters generally in the breeding range from southern Canada south to Mexico. STATUS: Locally common; population is declining across southern portions of its range. HABITAT: Primarily inhabits marshland and open grasslands, but also tundra, open fields, forest clearings, sagelands, deserts, pastures, prairies, lower mountain slopes, canyons, arroyos, dunes, meadows, and other open habitats. In winter, prefers open areas with little or no snow. SPECIAL HABITAT REQUIREMENTS: Extensive open grasslands with an abundance of rodents. NEST: Nests are sometimes in small loose colonies, placed in slight depressions on the ground, either in exposed situations or in grassy cover among clumps of weeds or grasses. Rarely, will nest in an excavated burrow. FOOD: Preys primarily on small mammals, especially voles; also eats birds, bats, and large insects. REFERENCES: Clark 1975, DeGraff et al. 1980, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974, Low and Mansell 1983, Tate and Tate 1982. 203 Boreal Owl Aegolius funereus L10" RANGE: Breeds from central Alaska and central Yukon to central Quebec and Labrador, south to northern British Columbia and central Alberta, across to northeastern Minnesota, western and central Ontario, southern Quebec and New Brunswick; also to central Colorado and northeastern Wyoming in the Rocky Mountains. STATUS: Local and uncommon in United States. HABITAT: Occurs in mixed coniferous-hardwood forests, but prefers extensive growth of stunted spruce in close proximity to open grasslands. Also inhabits dense alder thickets and forest edges. SPECIAL HABITAT REQUIREMENTS: Abandoned woodpecker holes in dead or live trees for nesting. NEST: Prefers abandoned northern flicker or pileated woodpecker cavities in conifers, but will also nest in woodpecker holes in deciduous trees. Usually locates nest 10 to 25 feet above the ground. Sometimes nests in natural cavities and rarely in abandoned bird nests. FOOD: Preys primarily on small mammals such as lemmings, voles, and mice. Also eats insects, bats, some frogs, salamanders, small snakes and lizards, and during the nesting season, a few birds. REFERENCES: Heintzelman 1979, Karalus and Eckert 1974. 204 Northern Saw-whet Owl Aegolius acadicus (formerly Saw-whet Owl) RANGE: Breeds from southern Alaska, central British Columbia, and central Alberta to southern Quebec and northern New Brunswick, south to southern California, central Mexico, extreme western Texas, central Missouri, southern Wisconsin, central Ohio, West Virginia, and New York; also in the mountains of eastern Tennessee and western North Carolina. Winters generally throughout the breeding range, south irregularly to southern Arizona, the Gulf Coast, and central Florida. STATUS: Uncommon. HABITAT: Favors dense woods, especially swampy areas of coniferous or hardwood forests. Also inhabits tamarack bogs, alder thickets, cedar groves, woodlots, and roadside shade trees; may take up temporary residence in or around a barn. Prefers cedar groves and vine clusters for roosting. SPECIAL HABITAT REQUIREMENTS: Tree cavities large enough for nesting and roosting. NEST: Usually nests in abandoned nest holes of northern flickers, hairy woodpeckers, or other woodpeckers but will use natural cavities of suitable size. Usually nests 20 to 40 (range 14 to 60) feet above the ground. Occasionally uses nest boxes with a layer of straw or sawdust. FOOD: Mostly eats small mammals; also preys on small birds, some insects, and frogs. REFERENCES: DeGraff et al. 1980, Heintzelman 1979, Johnsgard 1979, Karalus and Eckert 1974. 205 Lesser Nighthawk Chordeiles acutipennis L 8" W 21" RANGE: Breeds from central interior California, southern Nevada, extreme southwestern Utah, central Arizona, central New Mexico, and central and southeastern Texas south to South America. Winters from Mexico south to South America. STATUS: Common. HABITAT: Inhabits bare or somewhat brushy country in low deserts of the Southwest. Occurs around dry fields, dry washes and riverbeds, sandy flats, and broad, rocky, sparsely vegetated valleys. NEST: Generally lays eggs on bare ground in open sandy or gravelly areas but also in brushy areas, lowlands, hills, canyons, and dry rocky slopes and mesas, or on flat gravel and asphalt roofs. FOOD: Favors areas with concentrations of flying insects, near trees and brush along springs and streams; catches a variety of insects in flight, including winged ants, mosquitoes, June bugs, beetles, moths, and grasshoppers. REFERENCES: Bent 1940b, Harrison 1979, Oberholser 1974a, Terres 1980. 206 Common Nighthawk Chordeiles minor L 9" W 23" RANGE: Breeds from southern Yukon and southern Mackenzie to central Quebec and southern Labrador, south to southern California, southern Nevada, southern Arizona, Texas, the Gulf Coast, Florida, Mexico, and Central America. Winters in South America. STATUS: Common; population is declining in some regions. HABITAT: Inhabits varied habitats throughout most of North America. Prefers open habitats such as grasslands, sparse woods, or towns and cities. Also inhabits areas with plowed fields, and gravel beaches, as well as railroad right-of-ways and barren areas with rocky soils. NEST: Lays eggs on flat substrates such as gravelly ground, burned-over areas, gravel and asphalt rooftops, dry barren plains, bare rock, and partially vegetated soil, but always in the open. FOOD: Mainly crepuscular and nocturnal; sweeps flying insects, from tiny gnats to large moths, out of the air. Eats large quantities of mosquitoes and flying ants as well as beetles, plant lice, grasshoppers, locusts, horseflies, and other insects. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Tate and Tate 1982, Terres 1980. 207 Common Pauraque Nyctidromus albicollis (formerly Pauraque) L 11" RANGE: Resident in southern Texas south to South America. STATUS: Common. HABITAT: During the breeding season inhabits brushy woods, open chaparral, rivers that are lined by trees and brush. Prefers patches of mesquite and ebony blackbead woodlands. In winter, prefers dense bottomland thickets. NEST: Lays eggs on the ground in open woodlands with a leaf-covered floor, in brush, or on the edges of fields. Does not build a nest, but in more open areas may conceal eggs by a bush. FOOD: Usually perches on ground and flies up to snatch airborne insects; sometimes perches on low dead limbs or on top of a bush to hunt for insects. Eats low-flying moths, beetles, locusts, bugs, bees, wasps, butterflies, and other insects. REFERENCES: Bent 1940a, Harrison 1979, Oberholser 1974a. 208 Common Poorwill Phalaenoptilus nuttallii (formerly Poor-will) L 7" RANGE: Breeds from southern interior British Columbia, Montana, southeastern Alberta, and southwestern South Dakota south to Baja California, central Mexico, and central Texas and east to eastern Kansas. Winters in southern portion of breeding range. STATUS: Common. HABITAT: Inhabits a variety of arid and semiarid habitats in the West, from lowlands up to 8,000 to 10,000 feet on mountain slopes. Inhabits sparse desert brushland, open prairies, open pinyon-juniper woodlands, mixed chaparral-grassland, brushy rocky canyons, mountain scrub, and pine-oak woodlands. Seems to prefer rocky habitats with scrubby cover or xeric woodlands. NEST: Lays eggs on gravelly ground or on a flat rock, or in a slight hollow scraped in the ground. Often lays eggs so that they are partially shaded by a bush, weeds, or a tuft of grass. FOOD: Catches insects such as moths, beetles, chinch bugs, locusts, and grasshoppers by leaping from the ground or a perch, or by picking them up from the ground. REFERENCES: Bent 1940a, Bevier in Farrand 1983b, Oberholser 1974a, Terres 1980. 209 Chuck-will’s-widow Caprimulgus carolinensis l li- RANGE: Breeds from eastern Kansas, southern Iowa, and central Illinois to New Jersey and southern New York, south to south-central and southeastern Texas, the Gulf Coast, and southern Florida. Winters from southeastern Texas, Louisiana, and northern Florida to South America. STATUS: Locally common. HABITAT: Prefers mixed oak and pine forests, but also inhabits evergreen oak groves, forest edges, and woodlands along river courses. During migration and in winter, frequents open woodland and scrub and palmetto thickets. NEST: Lays eggs on the ground on dead leaves, usually at the edges of forests, near roads or other clearings, usually with little or no undergrowth around the eggs. FOOD: Forages at night by flying along edges of woods and fields, catching night-flying moths, beetles, and other insects. Rarely captures small birds such as hummingbirds, warblers, and sparrows and swallows them whole. REFERENCES: Bent 1940a, Forbush and May 1955, Johnsgard 1979, Sykes in Farrand 1983b, Terres 1980. 210 Whip-poor-will Caprimulgus vociferus RANGE: Breeds from north-central Saskatchewan and southern Manitoba to southern Quebec and Nova Scotia, south to eastern Kansas, northeastern Texas, and northern Louisiana across to central Georgia; and from southern California, southern Nevada, central Arizona, and extreme western Texas south to Mexico. Winters from southern Texas, the Gulf Coast, and east-central South Carolina south to Central America. STATUS: Common; population is declining slightly throughout the breeding range. HABITAT: In the East, prefers open hardwood or mixed woodlands of pine, oak, and beech, particularly younger stands in fairly dry habitats, also favors stands with scattered clearings. In the Southwest, frequents densely wooded slopes of oak and pine in canyons and mountains. NEST: Lays eggs on dead leaves on well-drained ground, usually in areas of partial shade where there is no undergrowth. Often nests among trees at the edge of a clearing or path, sometimes laying eggs in the shade of a small bush. FOOD: Feeds nocturnally, often pursuing insects attracted to lights near buildings in rural areas. Feeds in flight on moths, beetles, mosquitoes, ants, grasshoppers, June bugs, gnats, and other insects. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Raynor 1941, Tate and Tate 1982, Terres 1980. 211 Black Swift Cypseloides niger RANGE: Breeds locally from southeastern Alaska, northwestern and central British Columbia, and southwestern Alberta south through the Pacific states and Mexico to Central America; also in northwestern Montana, Colorado, and central Utah. Winters in Mexico and Central America. STATUS: Rare or uncommon. HABITAT: Found in areas with rocky cliffs available for nesting, varying from ocean cliffs to mountain ledges, at elevations from sea level to 11,000 feet. SPECIAL HABITAT REQUIREMENTS: Crevices or ledges on rocky cliffs for nesting, preferably near or behind a waterfall. NEST: Nests in small colonies, from 5 to 15 pairs, on a sea cliff, ledge, or cave, or in a crevice or ledge on a sheer, high, moist cliff face near or behind a waterfall, or over a pool. FOOD: Feeds exclusively on insects captured, usually high in the air, during long-distance foraging flights over all types of terrain. It eats various flies, midges, beetles, termites, flying ants, aphids, bees, wasps, and some spiders. REFERENCES: Bailey and Niedrach 1965, Lack 1956, Terres 1980, Verner and Boss 1980. 212 Chimney Swift Chaetura pelagica W 12Vi" RANGE: Breeds east of the Rocky Mountains from east-central Saskatchewan and southern Manitoba to southern Quebec and New Brunswick, south to eastern New Mexico, south-central and southern Texas, the Gulf Coast and south-central Florida. Winters in South America. STATUS: Common. HABITAT: Not confined to any single habitat, as breeding range is largely dependent on suitable nesting sites. Formerly found in habitats with mature, hollow trees, now occurs primarily in the vicinity of buildings in towns, cities, and around farms. SPECIAL HABITAT REQUIREMENTS: Hollow trees or structures such as chimneys, silos, cisterns, wells, rafters, and airshafts for nest sites. NEST: Originally adapted to nesting in caves and tree hollows, now mostly nests in man-made structures. Prefers dark areas and sheltered sites high above the ground. Constructs nest of twigs glued together with saliva; attaches nest to a vertical wall anywhere from near the top of a structure to more than 20 feet below the top. FOOD: Forages almost entirely on flying insects including beetles, flies, ants, termites, and bugs but will sometimes take small caterpillars hanging from tree branches or leaves. REFERENCES: Bailey and Niedrach 1965, DeGraff et al. 1980, Fischer 1958, Forbush and May 1955, Johnsgard 1979, Terres 1980. 213 Vaux’s Swift Chaetura vauxi f RANGE: Breeds from southeastern Alaska, northwestern and southern British Columbia, northern Idaho, and western Montana south, chiefly from the Cascades and Sierra Nevada westward, to central California south to northern Mexico. Winters in Central America. STATUS: Uncommon. HABITAT: Inhabits forested regions with large trees. In Montana, it occurs in mixed forests of Douglas-fir, ponderosa pine, lowland fir, western larch, yellow birch, aspen, and cottonwoods; in California, inhabits ponderosa pine, mixed conifer, and Jeffrey pine forests, and possibly black oak woodlands. Also inhabits river valleys in dense Douglas-fir and redwood forests. SPECIAL HABITAT REQUIREMENTS: Hollow stubs or snags for nesting. NEST: Nests are usually in a tall, hollow, dead tree, or inside burned-out stumps, generally within 20 inches but up to 6 feet up from the bottom of the cavity. Also nests in chimneys, but is not dependent upon them. FOOD: Feeds exclusively on flying insects captured in mid-air in forest openings, especially over lakeshores and streams. REFERENCES: Baldwin and Hunter 1963, Baldwin and Zaczkowski 1963, Bent 1940b, Scott et al. 1977, Terres 1980, Verner and Boss 1980. 214 White-throated Swift Aeronutes saxatalis RANGE: Breeds from southern British Columbia, Idaho, Montana, and southwestern South Dakota south through the Pacific and southwestern States to Baja California, Mexico, and Central America, and east to western Nebraska and western Texas. Winters from central California, central Arizona and, rarely, southern New Mexico south to Central America. STATUS: Common. HABITAT: Inhabits areas with steep cliffs and deep canyons at elevations from near sea level to about 13,000 feet. Inhabits primarily mountainous country but also coastal cliffs, rugged foothills, and desert canyons; ranges over adjacent valleys while foraging. SPECIAL HABITAT REQUIREMENTS: Crevices in cliffs for nesting. NEST: Places nests in deep cracks and crevices in steep, rocky, often inaccessible cliff faces or canyons, from 10 to 200 feet or more above the base. Sometimes nests in cracks of building walls. FOOD: Feeds on flying insects captured over any terrain while flying swiftly, usually high above the ground. Eats flies, beetles, bees, wasps, ants, bugs, leafhoppers, and other insects. REFERENCES: Bailey and Niedrach 1965, Bent 1940b, Terres 1980, Verner and Boss 1980. 215 Broad-billed Hummingbird Cynanthus latirostris L 3W’ RANGE: Breeds from southeastern Arizona, southwestern New Mexico, and very locally in western Texas south through Mexico. Winters in Mexico, casually north to southern Arizona. STATUS: Common. HABITAT: Prefers desert mountain canyons, riparian woodlands, and higher desert washes, especially where sycamores, cottonwoods, willows, and mesquite are present. SPECIAL HABITAT REQUIREMENTS: Red or red and yellow flowers for nectar. NEST: Places nest on a branch of a small tree, or a stalk of a vine or shrub, usually 4 to 7 feet above the ground. FOOD: Prefers to feed on nectar from red, or red and yellow flowers such as ocotillo, paintbrushes, and others. Also gleans small insects and spiders from the undersides of branches and leaves. REFERENCES: Cottam and Knappen 1939, Johnsgard 1983b, Moore 1939, Terres 1980, Terrill in Farrand 1983b. 216 White-eared Hummingbird Hylocharis leucotis RANGE: Resident from Sonora, Chihuahua, Coahuila, Nuevo Leon, and Tamaulipas south through the highlands of Mexico, Guatemala, El Salvador, and Honduras to north-central Nicaragua. Irregularly, in summer, in the mountains of southern Arizona, southwestern New Mexico, and western Texas; northernmost populations are partially migratory. STATUS: Rare and irregular in United States. HABITAT: Prefers the undergrowth of oak forests but also occurs in pine woods, dense pine-oak forests, high mountain fir forests, partially open mountain country with scattered trees and shrubs, suburban gardens, and vacant lots with scattered shrubs and flowers. (Little is known about its habits in the United States, and there is no good evidence that it has ever nested in Arizona.) SPECIAL HABITAT REQUIREMENTS: Flowers, especially blue flowers such as salvia, for nectar. NEST: Nest sites are nearly always in shrubs or fairly low trees. FOOD: Feeds on flies and other insects found in honeysuckle and other flowers. Is easily attracted to hummingbird feeders. REFERENCES: Cottam and Knappen 1939, Johnsgard 1983b, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b. 217 Buff-bellied Hummingbird Amazilia yucatanensis L 3 3 A" RANGE: Resident from southern Texas in the lower Rio Grande Valley south to Honduras. Casual north to central and eastern Texas, and southern Louisiana. STATUS: Uncommon; once common, but has largely disappeared north of the border, probably as a result of habitat destruction and the spraying of insecticides. HABITAT: Inhabits semiarid lowlands dominated by woods or scrubby growth, preferring dense thickets, flowering bushes, and tangled vines along the banks of streams and ponds, resacas, and gullies. Also occurs in semiarid coastal scrub in open woods, chaparral thickets, farms and gardens, and in citrus groves. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar. NEST: Places nest 3 to 8 feet above the ground on a small drooping limb or horizontal fork of a twig in a small tree or bush such as anacahuita, ebony blackbead, or hackberry, but sometimes in willow. Often choses a nest site that is near a road, trail, or other clearing. FOOD: Feeds on the nectar of native flowers and eats insects. REFERENCES: Johnsgard 1983b, Oberholser 1974a, Terres 1980. 218 Violet-crowned Hummingbird mazilia violiceps RANGE: Breeds in southern Arizona in the Huachuca and Chiricahua mountains, and in southwestern New Mexico in Guadalupe Canyon south to central Mexico. Casual in southern California. With a few exceptions, withdraws into Mexico in winter. STATUS: Rare and local; first discovered breeding in the United States in 1959. HABITAT: Prefers riparian sycamore groves in desert mountain canyons. In the United States, generally associated with streamside plant life in the deserts and foothills of mountains. NEST: Builds a nest that is saddled to a horizontal limb in sycamores, 25 to 40 feet above the ground. FOOD: Probably consumes nectar and insects but no definitive reports have been published. REFERENCES: Johnsgard 1983b, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b, Zimmerman and Levy 1960. 219 Blue-throated Hummingbird Lampornis clemenciae L 5Va" RANGE: Breeds in southeastern Arizona in the Huachuca and Chiricahua Mountains, and from western Texas south to Oaxaca. Casual north to Colorado. Winters in Mexico, but occurs casually at Arizona feeders. STATUS: Fairly common to uncommon. HABITAT: Inhabits lush vegetation along wooded streamsides in mountain canyons. In Texas, found among baldcypress, pines, oaks, and bigtooth maples from 4,900 to 7,500 feet in elevation. SPECIAL HABITAT REQUIREMENTS: Nest sites that are sheltered from rain and sunlight, near an abundance of flowers for feeding, and within a few feet of a stream. NEST: Builds nest in sites that are completely covered from above, such as in vertical-walled canyons, rock overhangs, on plant stalks, under eaves of buildings, bridges, water towers, and inside buildings. May use the same nest site for several years, adding to the nest each time it is used. FOOD: Forages on drier canyon slopes to feed on nectar of agave and honeysuckle flowers; also frequents flower gardens. Also eats flies, bugs, small beetles, wasps, spiders, daddy-longlegs, and pollen. REFERENCES: Cottam and Knappen 1939, Johnsgard 1983b, Oberholser 1974a, Terres 1980, Terrill in Farrand 1983b. 220 Magnificent Hummingbird Eugenes fulgens (formerly Rivoli’s Hummingbird) 8 L 5" RANGE: Breeds in western Colorado, and from southeastern Arizona, southwestern New Mexico, and western Texas south to Panama. Winters in Mexico and Central America, though a few remain at Arizona feeders. STATUS: Rare in Colorado, elsewhere uncommon to common. HABITAT: Found above 5,000 feet in deciduous woods along streams, and in pine or oak woods on mountain slopes and ridges. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar. NEST: Builds nest on a horizontal branch from 20 to 55 feet above the ground. Uses a variety of trees for nesting, including cottonwoods, mountain maples, sycamores, alders, walnuts, pines, and Douglas-fir. FOOD: Visits flowers for nectar, including those of agave, iris, and bright red salvia. Also eats leaf bugs, aphids, leafhoppers, parasitic wasps, beetles, flies, moths, and spiders. REFERENCES: Cottam and Knappen 1939, Johnsgard 1983b, Kaufman in Farrand 1983b, Terres 1980. 221 Lucifer Hummingbird Calothorax lucifer L3W RANGE: Breeds from southern Arizona and the Chisos Mountains of western Texas south to the highlands of Mexico. Winters in Mexico. STATUS: Rare and local. HABITAT: Inhabits cactus-covered slopes, dry canyons, arid mesas, and foothills and semi-desert habitats from 4,000 to 7,400 feet, often where century-plant agaves are abundant. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar, particularly agaves. NEST: Places nest in a shrub or on a seed pod of a dead stalk of an agave, 4 to 6 feet above the ground. FOOD: Forages on Chisos bluebonnet, ocotillo, tree tobacco, and yellow- green blossoms of agave for nectar; probably consumes a high proportion of insects as well. REFERENCES: Garrett in Farrand 1983b, Johnsgard 1983b, Oberholser 1974a, Terres 1980. 222 Ruby-throated Hummingbird Archilochus colubris L 3" im. a” RANGE: Breeds from central Alberta and central Saskatchewan to southern Quebec and New Brunswick south, east of the Rocky Mountains, to southern Texas, the Gulf Coast, and Florida. Winters from southern Texas south to Central America; also in southern Florida. STATUS: Common. HABITAT: Occurs in a variety of wooded habitats, ranging from rather dense to open coniferous and deciduous woodlands, orchards, and shade trees in yards. Also inhabits mixed woodlands, parks, and gardens, often breeding in woodlands near streams or wooded swamps. SPECIAL HABITAT REQUIREMENTS: Plants that provide tubular nectar¬ bearing flowers such as honeysuckle, lantana, gilia, and trumpet vine, especially red flowers. NEST: Places nest 6 to 50 feet, typically 10 to 20 feet, above the ground, on a fairly level or downward slanting twig or branch protected from above by larger branches or a leafy canopy. Often locates nest near or sometimes directly over water, or near a woodland trail. Uses a variety of trees for nesting, but appears to favor hardwoods over conifers, especially those with rough, lichen-covered bark. May use the same nest site year after year. FOOD: Consumes nectar from flowers, especially red, orange, and pink ones. Also eats small insects, spiders, and tree sap. REFERENCES: Beal and McAtee 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, 1983b, Terres 1980. 223 Black-chinned Hummingbird Archilochus alexandri o* L 3" RANGE: Breeds from southwestern British Columbia and northwestern Montana south to Baja California, southern Texas, and northern Mexico, and east to western Wyoming, eastern Colorado, eastern New Mexico, and central Texas. Winters in Mexico, casually to southern Texas, southern Louisiana, northwestern Florida, and southern California. STATUS: Common. HABITAT: Found most frequently in arid regions, where it inhabits oak and riparian woodlands of canyons and lowlands, small patches of willows along dry washes, chaparral, pine-oak woodlands, orchards, and parks. Perfers sites with a low percentage of canopy cover. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar. NEST: Usually places nest 4 to 8 feet (but up to 30 feet) above the ground, on a small drooping branch or in a fork of a small tree or shrub, near or overhanging a stream, spring, or dry creek bed. Prefers to nest in oaks, but also nests in alders, cottonwoods, sycamores, laurel, willows, apple, and orange trees in orchards, and in woody vines and tall herbaceous weeds. FOOD: Feeds on nectar from flowers. Also gleans insects from foliage, and hawks for flying insects. REFERENCES: Bent 1940b, Harrison 1979, Johnsgard 1983b, Phillips et al. 1964, Terres 1980, Verner and Boss 1980. 224 Anna’s Hummingbird Calypte anna RANGE- Breeds in western Washington, western Oregon, California, and southern Arizona. Nonbreeding birds occur regularly north to southern British Columbia and, rarely, east to western Texas. Winters from southwestern Oregon south to Mexico, and east to southern Arizona. STATUS: Common. HABITAT: Typically in habitats such as chaparral-covered hillsides, canyons, and mountain slopes and oak or sycamore woodlands in canyon bottoms. Also inhabits open mixed woodlands and chaparral, parks, and gardens from sea level to 5,900 feet, preferring timbered sites with sparse canopy cover. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar near the nest site, and forest openings. NEST: Places nest on a variety of substrates, including trees such as citrus, eucalyptus, and oaks, as well as chaparral bushes and vines at heights ranging from less than 3 to 30 feet. FOOD: Obtains nectar from many flowering plants. Also eats flies, bees, wasps, bugs, and spiders gleaned from flowers and foliage and hawked from the air. REFERENCES: Beal and McAtee 1912, Grinnell and Miller 1944, Johnsgard 1983b, Stiles 1973, Verner and Boss 1980. 225 Costa’s Hummingbird Calypte costae c f RANGE: Breeds from central California, southern Nevada, and southwestern Utah south to southern Arizona and Mexico. Winters from southern California and southern Arizona south to Mexico. STATUS: Common. HABITAT: In southwestern deserts, frequents arid washes and hillsides, dry chaparral, and suburban areas where exotic plants have been introduced. In California, inhabits washes, mesas, and hillsides, particularly where sages, ocotillo, yuccas, and cholla cacti are abundant. Relatively independent of water during the breeding season, and thus occupies drier habitats than other hummingbirds. SPECIAL HABITAT REQUIREMENTS: Flowering plants for nectar. NEST: Builds nests in a variety of sites, usually from 1 to 9 feet above the ground, on twigs or limbs of oaks, alders, hackberry, willow, palm, citrus trees in open orchards, or other trees; in sage, dead yuccas, branching cacti, and paloverde; or on vines clinging to rock faces. FOOD: Obtains nectar and insects from a variety of flowering desert plants; also eats spiders. REFERENCES: Garrett in Farrand 1983b, Grinned and Miller 1944, Johnsgard 1983b, Terres 1980. 226 Calliope Hummingbird Stellula calliope RANGE: Breeds in the mountains from central interior British Columbia and southwestern Alberta south to Baja California, and east to northern Wyoming and western Colorado. Winters in Mexico. STATUS: Uncommon. HABITAT: Frequents meadows and canyons; riparian aspen, willow, and alder thickets; and other brushy areas within the coniferous forests of western mountains. Occupies a broad vertical range during the breeding season, from 600 feet in the northern portions of its range to 11,500 feet in the Sierra Nevada. Prefers timbered stands near water with a low to intermediate canopy cover. SPECIAL HABITAT REQUIREMENTS: Flowers, preferably red, for nectar. NEST: Typically locates nest below a larger branch or canopy of foliage, usually on a branch that has small knots of dead, black or gray mistletoe or pine cones which nest strongly resembles. Locates nest 2 to 70 feet above ground, frequently in a riparian area. May use the same site in subsequent years. FOOD: Obtains nectar from a variety of flowers, preferring red ones. Hawks for small flying insects, and eats spiders. REFERENCES: Bent 1940b, Calder 1971, Garrett in Farrand 1983b, Johnsgard 1983b, Terres 1980, Verner and Boss 1980. 227 L 3W Broad-tailed Hummingbird Selasphorus platycercus RANGE: Breeds in the mountains from north-central Idaho, northern Utah, and northern Wyoming south to southeastern California, northern Mexico, and western Texas. Winters in Mexico. STATUS: Common. HABITAT: Inhabits meadows and patches of flowers within pine, spruce, fir, and sometimes aspen forests from 4,000 to 11,000 feet. Also inhabits gardens in towns and cities, and sometimes ranges eastward onto the Plains. SPECIAL HABITAT REQUIREMENTS: Flowers for nectar. NEST: Nests are saddled on large horizontal limbs or small twigs in shrubs along moist canyon walls, in Douglas-fir, ponderosa pine, subalpine fir, or other conifer, oak, aspen, alder, willow, or cottonwood. Usually locates nest 4 to 15 feet above the ground, and frequently uses the same location for several consecutive summers. FOOD: Obtains nectar from a variety of flowers. Comes to hummingbird feeders for sugar water and eats small insects and spiders. REFERENCES: Bailey and Niedrach 1965, Bent 1940b, Johnsgard 1979, 1983b, Phillips et al. 1964, Terres 1980. 228 Rufous Hummingbird RANGE: Breeds from southern Alaska, southern Yukon, and western and southern British Columbia to western Montana south, primarily in the mountains, to northwestern California, eastern Oregon, and central Idaho. Winters in Mexico, in small numbers to southern Texas and the Gulf Coast and, rarely, in coastal southern California. STATUS: Common. HABITAT: Occurs in northwestern parks and gardens, in chaparral, and in meadows, forest edges, and riparian thickets of coniferous woodlands. During migration, may be found in high mountain meadows as well as in the Pacific lowlands in open areas where flowers are present. SPECIAL HABITAT REQUIREMENTS: Flowers (especially red) for nectar. NEST: In northern latitudes, builds nest close to the ground where it is sheltered from wind and cold; otherwise, builds nest 5 to 50 feet above ground. May nest in a variety of sites, sometimes in colonies with up to 20 nests in a small area. Favors the drooping branches of conifers, but also nests in bushes and among vines. Sometimes builds a new nest on top of the previous year’s nest. FOOD: Consumes nectar from flowers, especially red flowers. Also eats insects. REFERENCES: Bent 1940b, Garrett in Farrand 1983b, Johnsgard 1983b, Terres 1980. 229 Allen’s Hummingbird Selasphorus sasin 9 L 3" RANGE: Breeds from southwestern Oregon south through coastal California to Santa Barbara County. Resident in southern California in the Channel Islands and on the Palos Verdes Peninsula. Winters in Mexico. STATUS: Common. HABITAT: Found within the Pacific coastal fog belt, inhabiting meadows, moist canyon bottoms, humid woody or brushy ravines, brushy edges of coniferous forest, coastal chaparral, and parks. SPECIAL HABITAT REQUIREMENTS: Shade, preferably patchy, over the nest site, and flowers for nectar. NEST: Usually builds nest on a site with several separate supports such as a dense tangle of vines. Less frequently, attaches nest to the side of a drooping twig or limb from 1 to 90 feet above the ground in trees such as oaks, eucalyptus, and Monterey cypress, or in shrubs in streamside thickets. FOOD: Obtains nectar from a variety of flowers. Also hawks for small insects. REFERENCES: Aldrich 1945, Bent 1940b, Harrison 1979, Terres 1980. 230 Elegant Trogon Trogon elegans (formerly Coppery-tailed Trogon) RANGE: Resident from southern Arizona, primarily in Chiricahua, Huachuca and Atascosa mountains south to Costa Rica. Mostly migratory in northernmost part of range; casual in Arizona, southwestern New Mexico, and southern Texas in winter. STATUS: Locally fairly common. HABITAT: Occurs in oak and pine-oak forests in mountain canyons, and in sycamore, walnut, and cottonwood groves along canyon streams. SPECIAL HABITAT REQUIREMENTS: Natural cavities in trees or large, deserted woodpecker holes. NEST: Builds nest inside cavities of large streamside trees such as sycamores or cottonwoods, 12 to 40 feet above the ground. FOOD: Primarily eats insects and some fruits. REFERENCES: Bent 1940a, Cottam and Knappen 1939, Oberholser 1974a, Terres 1980, Terrill in Farrand 1983b. 231 Belted Kingfisher Ceryle alcyon L 12" RANGE: Breeds from western and central Alaska, central Yukon, and western and south-central Mackenzie to central Quebec and east-central Labrador south to southern California, southern Texas, the Gulf Coast, and central Florida. Winters from south-coastal and southeastern Alaska, central and southern British Columbia, and western Montana across to Nebraska, the southern Great Lakes and New England south to South America. STATUS: Common. HABITAT: Occurs in the vicinity of ponds, lakes, rivers, and streams, even rocky seacoasts near areas of exposed vertical ground such as bluffs, road cuts, gravel pits, or sandbanks. Prefers small, clear bodies of water to large lakes. In winter, frequents ice-free waters that allow access to food. SPECIAL HABITAT REQUIREMENTS: Nests preferably within 1 mile of water with low turbidity supporting adequate aquatic animal populations, and perches near water to sight prey. NEST: Typically excavates a nest burrow 3 to 6 feet, up to 15 feet, deep in a bank with sandy, gravelly, or clay soil. Constructs burrow at least 5 feet above level ground or water, and usually within 3 feet of the top of a bank. Occasionally locates burrow far from water, and at times may have to forage up to 5 miles from the nest site. Builds a nest cavity that is an enlarged area at the end of the burrow, often lined with disgorged food pellets. FOOD: Feeds primarily on fish averaging 3 to 4 inches, caught by diving into water. Forages from a perch or while hovering over water. Also may eat crayfish, mollusks, frogs, tadpoles, lizards, newts, mice, large insects, and occasionally fleshy fruits. REFERENCES: Cornwall 1963, DeGraff et al. 1980, Johnsgard 1979, Terres 1980, White 1953. 232 Green Kingfisher RANGE: Resident in south-central and southern Texas, occurring up the Rio Grande at least as far as Del Rio, north onto Edwards Plateau, and south through Mexico to South America. STATUS: Fairly common. HABITAT: Prefers small, shaded, clear streams and quiet backwaters, but also found around larger bodies of water with dense, low vegetation along the banks. SPECIAL HABITAT REQUIREMENTS: High banks along clear streams with an abundant supply of small fishes. NEST: Digs burrow 2 to 3 feet deep near the top of a high bank over water with a small entrance, about 2 to 3 inches in diameter, usually well concealed by trailing plants and vines or dead vegetation draping the top of the bank. FOOD: Sights small fishes from a low perch of overhanging branches, roots, or rocks, often just inches above water; plunges into the water from the perch to catch the fishes, rarely hovering. REFERENCES: Bent 1940a, Kaufman in Farrand 1983b, Oberholser 1974a, Terres 1980. 233 Lewis’ Woodpecker Melanerpes lewis RANGE: Breeds from southern British Columbia to southwestern South Dakota and northwestern Nebraska south to south-central California, central Arizona, southern New Mexico, and eastern Colorado. Winters from northern Oregon, southern Idaho, central Colorado, and south- central Nebraska south irregularly to northern Baja California, northern Mexico, southern New Mexico, and west Texas. STATUS: Population has been declining; has been placed on the blue list. HABITAT: Inhabits open country with scattered trees rather than dense forests; open or parklike ponderosa pine forests are probably the major breeding habitat. Is attracted to burned-over stands of Douglas-fir, mixed conifer, pinyon-juniper, riparian, and oak woodlands but is also found in fringes of pine and juniper tree stands and in deciduous forests, especially riparian cottonwoods. Prefers areas with a good understory of grasses and shrubs to support insect prey populations. Winters over a wide range of habitats, especially where oaks are present. SPECIAL HABITAT REQUIREMENTS: Dead trees or tall stumps for cavity nests. NEST: Generally excavates its own nest cavity in dead trees or tall stumps, but will use natural cavities or old excavated nest sites. Nests about equally in coniferous and deciduous trees, but favors ponderosa pine, cottonwood, and sycamore. FOOD: Primarily eats insects during spring and summer. Catches flying insects by hawking from perches in dead trees or stumps. Mostly eats fruits and berries during fall. Gathers and stores winter food, mostly acorns, in crevices of dead trees, power poles, or oak bark. REFERENCES: Beal and McAtee 1912, Bock 1970, Johnsgard 1979, MacRoberts and MacRoberts 1976, Oberholser 1974a, Tate and Tate 1982. 234 Red-headed Woodpecker Melanerpes erythrocephalus L 7VF RANGE: Breeds from southern Saskatchewan, southern Ontario, southern New Hampshire, and southern New Brunswick south to central Texas, the Gulf Coast and Florida, extending west to central Montana, eastern Wyoming, eastern Colorado, and central New Mexico, rarely to northeastern Utah. Winters regularly through the southern two-thirds of the breeding range, rarely or casually north to the limits of the breeding range. STATUS: Common, but declining in the Southeast. HABITAT: Inhabits relatively open forests or woodlots with low stem density, preferring savannahlike grasslands with scattered trees and forest edges. Attracted to areas with many dead trees which provide nesting and roosting sites, and lush herbaceous ground cover that produces abundant insect populations. Tends to avoid forests with closed canopies, but will move from forest edges to the interior during winter. SPECIAL HABITAT REQUIREMENTS: Relatively open forests with dead and dying trees for cavities and feeding perches. NEST: Nests generally in the trunk of a dead tree but sometimes in a dead limb. Tends to select isolated snags for nesting, especially those without bark. FOOD: In summer, mostly eats insects caught by hawking from perches in dead trees; stores mast, mainly acorns, beechnuts, and corn, under bark, in cracks, knotholes, and tree cavities for winter use. REFERENCES: Beal 1911, Conner and Adkisson 1977, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, MacRoberts and MacRoberts 1976, Reller 1972, Tate and Tate 1982. 235 Acorn Woodpecker Melanerpes formicivorus RANGE: Resident west of the Cascade and Sierra Nevada Mountains from northwestern Oregon south through California to southern Baja California; and from northern Arizona, northern New Mexico, and western Texas south through the highlands of Central America to Colombia. STATUS: Common within its range. HABITAT: Inhabits mixed pine-oak woodlands and adjacent open grasslands along the Pacific Coast and the Southwest. SPECIAL HABITAT REQUIREMENTS: Oak trees to provide food and large snags that provide sites for nesting, roosting, and food storage. NEST: Excavates cavities in pine snags, living and dead oaks, sycamores, cottonwoods, and willows for nesting and roosting. May use the same cavity for several years. Nests communally and all birds of a community assist in feeding young. FOOD: Mostly consumes acorns, which are stored in holes drilled in communal trees. Also consumes sap from oaks from midwinter to summer, and hawks insects when available. REFERENCES: Beal 1911, MacRoberts 1970, MacRoberts and MacRoberts 1976, Overholser 1974a. 236 Gila Woodpecker Melanerpes uropygialis L 8!4" RANGE: Resident from southeastern California, extreme southern Nevada, central Arizona, and southwestern New Mexico south through Baja California and western Mexico to Jalisco. STATUS: Locally common. HABITAT: Inhabits desert mesas in association with creosotebush, mesquite, and saguaro and organ-pipe cactus. Also occurs in riparian areas and in foothill canyons among cottonwoods, willows, and sycamores. SPECIAL HABITAT REQUIREMENTS: Suitable nest sites in the Sonoran Desert such as cottonwoods, mesquites, and saguaros. NEST: Excavates holes mainly in saguaro but uses cottonwoods, willows, and mesquites at higher elevations and in riparian habitat. FOOD: Mainly eats insects; also eats fruits and nuts, and occasionally preys on nestling birds and eggs. REFERENCES: Bent 1939, Phillips et al. 1964, Terres 1980. 237 Golden-fronted Woodpecker Melanerpes aurifrons RANGE: Resident from southwestern Oklahoma and north-central Texas south through central Texas to Mexico and Central America. STATUS: Locally abundant; populations may be declining because of mesquite control programs. HABITAT: Prefers mesquite and riparian woodlands, but also inhabits semiarid brush country, pecan groves, and upland oak stands. SPECIAL HABITAT REQUIREMENTS: Trees large enough for nest cavity. NEST: Excavates nest hole in large living or dead trees, especially in mesquite, oak, and pecan. May also excavate holes in fence posts and telephone or electric poles; occasionally uses bird boxes. FOOD: Gleans much of its food, which consists of insects, acorns, pecans, wild fruits, corn, and occasionally citrus. REFERENCES: Bent 1939, Johnsgard 1979, Oberholser 1974a. 238 Red-bellied Woodpecker Melanerpes carolinus L 8V2" RANGE: Resident from southeastern Minnesota, south-central Wisconsin, southern Michigan, southern Ontario, central New York, and Massachusetts south to central Texas, the Gulf Coast, and southern Florida, and west to Iowa, eastern Nebraska, western Kansas, central Oklahoma, and north-central Texas. STATUS: Common. Range has been expanding westward along the river valleys of the Great Plains. HABITAT: Inhabits deciduous and coniferous forests of the southeastern United States, especially mature woodlands with dead and broken- topped trees. Also frequents farm woodlots, riparian forests, and orchards. SPECIAL HABITAT REQUIREMENTS: Mature woodlands with dead trees or trees with large dead limbs for nesting. NEST: Nests are excavated in a variety of sites, including trees, stumps, dead limbs, and poles. May nest in buildings or in nest boxes usually less than 40 feet above the ground. Prefers relatively soft deciduous tree species and dead trees or those with decayed stubs. FOOD: Eats mostly vegetable material including fruit, nuts, poison ivy seeds, pine seeds, and juniper berries, but also feeds on some wood¬ boring insect larvae found in dead wood. REFERENCES: Beal 1911, DeGraff et al. 1980, Johnsgard 1979, Reller 1972. 239 Yellow-bellied Sapsucker Sphyrapicus varius (includes Red-naped Sapsucker Sphyrapicus nuchalis) L 7%" RANGE: Breeds from eastern Alaska east to central Newfoundland, south to northeastern British Columbia, eastern North Dakota, New Hampshire, and locally in the Appalachians south to eastern Tennessee and western North Carolina; and in the Rocky Mountain region from south-central British Columbia to western Montana south, east of the Cascades, to east-central California and western Texas. Winters from Missouri, the Ohio Valley, and New Jersey south through Texas, southeastern United States to central Panama; in the West from southern California, central Arizona, and central New Mexico south to southern Baja California and Jalisco. STATUS: Common. HABITAT: Breeds in deciduous and mixed deciduous-coniferous forests in the eastern and northern parts of its range, especially in woodlands with Populus. In the Rocky Mountain region, occurs primarily in aspen forests or in coniferous forests where aspen is present. Uses a variety of forests and open woodlands, parks, and orchards in winter. SPECIAL HABITAT REQUIREMENTS: Dead or live trees with heartrot for cavity nests. NEST: Excavates cavities in snags or in live trees with rotten heartwood. Prefers aspens but will nest in ponderosa pine, birch, elm, butternut, cottonwood, alder, willow, beech, maple, and fir. May use the same nest tree for several years, but excavates a new cavity each year. FOOD: Drills rows of holes (sapwells) through the outer bark and consumes cambium and tree sap. Also eats a variety of insects attracted to the sapwells (ants are probably the dominant animal food). Fruits, mast, and Rhus seeds are included in the diet. REFERENCES: Beal 1911, DeGraff et al. 1980, Howell 1952, Johnsgard 1979, Lawrence 1967, Thomas et al. 1979. 240 Red-breasted Sapsucker Sphyrapicus ruber (split from Yellow-bellied Sapsucker) L 7 VS RANGE: Breeds from southeastern Alaska and coastal and central interior British Columbia south, west of the Cascades, to northwestern California, and in the Sierra Nevada to east-central California and extreme western Nevada; locally in the mountains of southern California and southern Nevada. Winters throughout the breeding range (except interior British Columbia) south through most of California (west of the deserts) to Baja California. STATUS: Locally common. HABITAT: Breeds in coniferous and conifer-aspen associations including the humid coastal lowlands. Also occurs in open woodlands and parks in winter. SPECIAL HABITAT REQUIREMENTS: Live or dead trees suitable for cavity nests. NEST: Apparently excavates a cavity in a variety of tree species, including aspen, alder, cottonwood, fir, willow, and birch. FOOD: Drills sapwells in a variety of tree species and consumes sap and cambium. Also feeds on a variety of insects, especially ants, and some fruits. REFERENCES: Beal 1911, Bent 1939. 241 Williamson’s Sapsucker Sphyrapicus thyroideus 9 RANGE: Breeds from extreme southern interior British Columbia, Idaho, western Montana, and Wyoming south in the mountains to northern and east-central California, central Arizona, and southern New Mexico. Winters generally from the breeding range south to Baja California, and east to western Texas and Mexico. STATUS: Common. HABITAT: Prefers mixed conifer-hardwood forests in the Rocky Mountain region but also inhabits the subalpine spruce-fir-lodgepole zone, and ponderosa pine, Douglas-fir, and aspen forests. SPECIAL HABITAT REQUIREMENTS: Dead or live trees infected with Fomes, a heartrot fungi, for cavity nest sites. NEST: Chooses different tree species for cavity nests in different regions. In some areas, nests primarily in conifers; in others, prefers aspen, especially those infected with Fomes. In Colorado and Arizona, mostly nests in aspen snags or live aspen infected with Fomes. FOOD: Drills rows of pits in the bark of lodgepole pine, hemlock, red and white firs, Jeffrey pine, and aspen and consumes sap and cambium. Eats ants for most of its animal food but also wood-boring larvae, moths of spruce budworm, and other insects. REFERENCES: Baily and Neidrach 1965, Beal 1911, Bent 1939, Burleigh 1972, Crockett and Hadow 1975, Hubbard 1965, Jackman 1975, Ligon 1961, Oliver 1970, Packard 1945, Rasmussen 1941, Tatschl 1967. 242 Picoides scalaris Ladder-backed Wood RANGE: Resident from southern interior California, irregularly to western Oklahoma south through Texas and Mexico to Central America. STATUS: Locally common. HABITAT: Occurs in wooded canyons, cottonwood groves, riparian woodlands in deserts, and dense growth of cholla cactus, creosotebush, catclaw, and other low-growing plants on borders of deserts. Also found in post oak and mesquite woodlands, and on lower slopes of mountains up to 5,500 feet. SPECIAL HABITAT REQUIREMENTS: Trees or other structures large enough for cavity nests. NEST: Excavates nest holes in a variety of trees (mesquite, screw bean, palo verde, hackberry, china tree, willow, cottonwood, walnut, and oak), usually 2 to 30 feet above ground. Sometimes uses saguaro, yucca stalks, telephone poles, and fence posts for nesting. FOOD: Eats mostly insects, especially larvae of wood-boring beetles, caterpillars, and ants, but also eats fruit of various cacti. REFERENCES: Bent 1939, Johnsgard 1979, Oberholser 1974a, Phillips et al. 1964. 243 Nuttall’s Woodpecker Picoides nuttallii RANGE: Resident from northern California south, west of the deserts and the Sierra divide, to Baja California. Casual or accidental in southern Oregon, southeastern California, and Arizona. STATUS: Common. HABITAT: Occurs in oak woodlands, live oak forests, and chaparral, and in canyons with sycamores, alders, cottonwoods, and bay trees growing along streams lined with live oaks. SPECIAL HABITAT REQUIREMENTS: Cavity nest sites. NEST: Excavates cavities in dead limbs and trunks of trees, 3 to 45 feet above ground in oak, willow, sycamore, cottonwood, elder, and alder trees. FOOD: Consumes a diet consisting of about 80 percent insects (beetles, bugs, caterpillars, and ants) which are gleaned from tree trunks and limb surfaces, or captured on the wing. Also eats wild fruits, poison oak seeds, and occasionally acorns. REFERENCES: Beal 1911, Miller and Bock 1972, Short 1971, Terres 1980. 244 Downy Woodpecker RANGE: Breeds from western and central Alaska, northern Alberta, northern Ontario, and Newfoundland south to southern California, central Arizona, the Gulf Coast, and southern Florida. Winters throughout the breeding range, but are mostly migratory in more northern populations and occurring irregularly southward. STATUS: Common throughout most of its range. HABITAT: Inhabits most of the wooded parts of North America, but absent or rare in arid deserts and less common in dense forests. Favors bottomlands but also inhabits open forests and woodlots, orchards, hummocks, farmyards, and urban areas. SPECIAL HABITAT REQUIREMENTS: Suitable cavity trees. NEST: Prefers to excavate its cavity-nests near the tops of dead trees or dead limbs of live trees in fairly open tree stands. Also nests in live trees, especially if heartrot is present. Generally, excavates new cavities each year; seldom reuses old cavities or cavities of other birds. In the fall, excavates fresh holes for winter roosts. FOOD: Consumes diet that is 75 percent animal and 25 percent vegetable material (beetles, mostly wood-boring larvae, make up a large portion of the diet). Also eats wild fruits, corn, poison sumac seeds, and mast. REFERENCES: Beal 1911, Bent 1939, DeGraff et al. 1980, Johnsgard 1979, Kilham 1970, Lawrence 1967, Thomas 1979. 245 Hairy Woodpecker Picoides villosus L 71/2' RANGE: Breeds from western and central Alaska, northern Saskatchewan, and Newfoundland south throughout most of North America to Central America and the Bahamas. Winters generally throughout the breeding range, with the more northern populations partially migratory southward. STATUS: Stable population throughout most of its range. HABITAT: Inhabits nearly all types of forest within its range, preferring bottomlands with large mature trees. Generally more abundant at the edge of woodlands. SPECIAL HABITAT REQUIREMENTS: Nest trees over 10 inches dbh. NEST: Excavates cavities in snags or in live trees with decaying heartwood. Usually chooses deciduous trees such as aspens, ashes, elms, or cottonwoods. FOOD: Consumes a diet that is about 80 percent animal food (wood¬ boring beetles removed from dead and diseased trees are an important source of food). Also eats other insects, fruits, corn, nuts, and cambium. REFERENCES: Beal 1911, Bent 1939, DeGraff et al. 1980, Johnsgard 1979, Kilham 1968, Lawrence 1967, Tate and Tate 1982, Thomas et al. 1979. 246 Strickland’s Woodpecker Picoides stricklandi (includes Arizona (Brown-backed) Woodpecker) RANGE: Resident from southeastern Arizona and extreme southwestern New Mexico south to Mexico. STATUS: Locally common within its limited range. HABITAT: In the northern part of range, occurs on mountain slopes and is primarily associated with oaks, but sometimes in riparian sycamores, cottonwoods, walnuts, and willows. In the southeastern part of range also associated with pines. SPECIAL HABITAT REQUIREMENTS: Cavity nest sites. NEST: Excavates holes in dead branches of live or dead trees, primarily walnuts, oaks, maples, and sycamores. FOOD: Feeds mainly on insects and their larvae, and some fruits and nuts. REFERENCES: Bent 1939, Davis 1965, Terres 1980. 247 Red-cockaded Woodpecker Picoides borealis RANGE: Resident locally from eastern Oklahoma, southern Missouri, northern Arkansas, northern Mississippi, northern Alabama, northern Georgia, southeastern Virginia, and southern Maryland south to eastern Texas, the Gulf Coast, and southern Florida, and north in the Cumberland plateau through eastern Tennessee to Kentucky. STATUS: Endangered. HABITAT: Endemic to the yellow pine forests of the southeastern United States, where hardwoods make up less than 35 percent of the tree stand. Generally, inhabits mature forests (at least 60 years old) or younger forests where groups of mature trees are present. It is found in forests dominated by several species of pine, but probably the largest populations are found where longleaf pine is prevalent. SPECIAL HABITAT REQUIREMENTS: Mature living pines with heartrot for nesting, and extensive pine stands for foraging. NEST: Excavates nest holes in mature living pines infected with red heartrot. Same pair may reuse a cavity for several years. Breeds cooperatively with auxiliary or helper birds (clan) aiding a mated pair in the rearing of young. Clan size is generally two to four birds at the beginning of the breeding season. FOOD: Prefers living pines for foraging substrate, especially larger pines. Consumes mostly insects (larvae of wood-boring insects, beetles, grubs, ants, crickets, caterpillars, scales, bark lice, and grasshoppers). Also consumes mast (primarily seeds of conifers), fruit pulp, and poison-ivy and bayberry seeds. REFERENCES: Baker 1971, Beal 1911, Crosby 1971, Hopkins and Lynn 1971, Jackson 1971, Johnsgard 1979, Lennartz 1984, Ligon 1970, 1971b, Oberholser 1974a, Steirly 1957. 248 White-headed Picoides albolarvatus Woodpecker RANGE: Resident from southern interior British Columbia, north-central Washington, and northern Idaho south through Oregon, east of the Cascades, to southern California and west-central Nevada. Casual in coastal and desert areas of southern California, but absent from the humid coastal coniferous forest. STATUS: Local. HABITAT: Primarily inhabits open ponderosa pine forests, but also occurs in sugar pine, Jeffrey pine, and red and white fir forests. Prefers forests with large trees and 40 to 70 percent canopy cover. SPECIAL HABITAT REQUIREMENTS: Dead trees for cavity nests. NEST: Usually excavates a new nest hole each year; seems to prefer dead pines. Nests in live and dead fir, oak, and aspen, with nest holes usually about 8 feet above the ground. FOOD: Consumes primarily pine seeds, during winter and early spring (60 percent of total diet) and insects and spiders during summer. REFERENCES: Bent 1939, Grinnel and Miller 1944, Ligon 1973, Tevis 1953, Verner and Boss 1980. 249 Three-toed Woodpe Picoides tridactylus (formerly Northern Three-toed Woe RANGE: Resident, often locally, from northwestern and central Alaska, northern Manitoba, northern Quebec, and Newfoundland south to western and southern Alaska, central Washington, and southern Oregon, in the Rocky Mountains to eastern Nevada, central Arizona, and south- central New Mexico, and to southwestern and central Alberta, southern Manitoba, northeastern Minnesota, central Ontario, northern New York, northern New England, and southern Quebec. STATUS: Locally common in western coniferous forests; rare in east. HABITAT: Primarily inhabits coniferous forests of the West, especially where fires have left large stands of dead trees. Also occasionally inhabits conifer stands in the Northeast. SPECIAL HABITAT REQUIREMENTS: Dead trees for cavity nests. NEST: Excavates nest cavities each year in dead trees or in dead limbs with decayed heartwood in live trees. Usually locates nest holes 5 to 12 feet above ground in pine, aspen, spruce, and cedar. FOOD: Feeds by probing and drilling for wood-boring larvae of moths and beetles (probably one of the most important birds in combating forest insect pests in the western United States). In Colorado, consumes spruce beetles for 65 percent of its annual diet and 99 percent of its winter diet. Also eats ants, wood-boring larvae, caterpillars, fruits, mast, and cambium. REFERENCES: Beal and McAtee 1912, Bent 1939, DeGraff et al. 1980, Jackman and Scott 1975, Johnsgard 1979, Koplin 1972, Massey and Wygant 1973, Thomas et al. 1979. 250 Black-backed Woodpe Picoides arcticus (formerly Black-backed Three-toed Woodpecker) L 8' RANGE: Resident, often locally, from western and central Alaska, southern Yukon, northern Manitoba, central Labrador, and Newfoundland south to southeastern British Columbia, through the Cascade, Siskiyou, and Warner Mountains and Sierra Nevada of Washington and Oregon to central California and west-central Nevada, through Montana to northwestern Wyoming and southeastern South Dakota, and to southwestern and central Alberta, southeastern Manitoba, northern Minnesota, north-central Michigan, northern New York, and northern New England. STATUS: Uncommon. HABITAT: Inhabits dense coniferous forests, especially in burned, swampy, cutover, or beetle-killed forests where dead trees are numerous. SPECIAL HABITAT REQUIREMENTS: Dead or live trees with dead heartwood for nesting and feeding sites. NEST: Usually excavates its cavities in snags or live trees with dead heartwood, especially in areas that have been burned or logged. Mostly nests in spruce, balsam fir, pines, or Douglas-fir, but also in maple, birch, cedar, and utility poles. Locates nest cavity usually less than 15 feet above the ground. FOOD: Flakes off bark of dead conifers to get at larvae of destructive wood-boring beetles, which make up about 75 percent of its food. Also eats weevils and other beetles, spiders, and ants, along with some wild fruit, mast, and cambium. REFERENCES: Beal and McAtee 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Thomas et al. 1979. 251 Northern Flicker Colaptes auratus (formerly Common Flicker) L 10 Vi" RANGE: Breeds from central Alaska, northern Manitoba, north-central Quebec, and Newfoundland south throughout most of North America and northwestern Mexico. Winters from southern Canada south through the breeding range. STATUS: Common. HABITAT: Commonly found near large trees in open woodlands, fields, meadows, and deserts throughout North America. In winter, occasionally seeks shelter in coniferous forests or in swamps. SPECIAL HABITAT REQUIREMENTS: Cavity nest sites. NEST: Prefers to nest near the top of broken-off stubs of dead trees in open country or in sparsely wooded parklike suburban areas. Also nests in and around openings in extensive forested areas. Excavates nests in dead or live trees of many species, including aspen, cottonwood, oak, willow, sycamore, pine, and juniper. FOOD: Searches for food, much of the time, on the ground. Consumes a diet that is about 60 percent animal food; of this, nearly 75 percent is ants. (Some flicker stomachs have contained over 2,000 ants.) The diet also includes other insects, weed seeds, cultivated grain, and the fruits of shrubs and trees. REFERENCES: Bailey and Niedrach 1965, Beal 1911, Conner et al. 1975 DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Lawrence 1967, Thomas et al. 1979. 252 Dryocopus pileatus Pileated Woodpecker L 15' RANGE: Resident from southern and eastern British Columbia, v * southwestern Mackenzie, central Manitoba, New Brunswick, and Nova Scotia south through Alberta, Washington, south-central Idaho, western Montana, and Oregon to northern California, west to eastern Dakotas, Missouri, and Oklahoma and south to east-central Texas, the Gulf Coast, and southern Florida. STATUS: Locally common but has become less common in areas where extensive agricultural or logging practices have eliminated large tracts of old-growth forests. HABITAT: Generally limited to mature coniferous, deciduous, and mixed forests with large, dead trees. Prefers woodlands near water. SPECIAL HABITAT REQUIREMENTS: Large dead trees for nesting and feeding. NEST: Usually excavates nest holes in dead tree trunks or dead limbs of living trees. Generally requires trees greater than 15 inches dbh for nest and roost cavities and generally uses ponderosa pine snags greater than 20 inches dbh. Nests in a variety of tree species, including beech, cottonwood, yellow-poplar, birch, oak, hickory, maple, hemlock, pine, ash, elm, basswood, and aspen. FOOD: Consumes a diet that is about 70 percent insects, with ants, especially carpenter ants and wood-boring beetles, predominating. Also consumes other insects, some wild fruits, mast, and seeds of sumac. REFERENCES: Beal 1911, Bull and Meslow 1977, Conner et al. 1975, DeGraff et al. 1980, Forbush and May 1955, Hoyt 1957, Johnsgard 1979. 253 Ivory-billed Woodpecker Campephilus principalis L 18 " RANGE: Formerly resident from eastern Texas, southeastern Oklahoma, northeastern Arkansas, southeastern Missouri, southern Illinois, southern Indiana, Kentucky, and southeastern North Carolina south to the Gulf Coast, southern Florida, and Cuba. STATUS: Endangered; may be extinct or nearly so. HABITAT: Inhabits old-age forests of bottomlands and swamps with dead and dying trees that provide a food source and nest sites. Most birds live in virgin or primitive tree stands, but virgin forests may not be essential as long as there are large numbers of recently dead trees to supply wood-boring grubs and large nesting trees. SPECIAL HABITAT REQUIREMENTS: Continual supply of dead and dying trees. NEST: Excavates a new hole each nesting season, seldom in the same tree, in almost all tree species occurring within its range, and in trunks of living and dead trees. FOOD: Consumes a diet that is about one-third wood-boring larvae. Most abundant in areas where recently dead and dying trees are numerous because of flooding, fire, insect attacks, or storms; stays as long as abundant wood-boring larvae are present. (Wood-boring larvae begin to decline after trees have been dead 2 or 3 years.) Also eats other insects and fruits. REFERENCES: Beal 1911, Cottam and Knappen 1939, Dennis 1967, Forbush and May 1955, Greenway 1958, Mackenzie 1977, Oberholser 1938, Pearson 1936, Tanner 1966, USDI Fish and Wildlife Service 1980. 254 Northern Beardless-Tyrannulet Camptostoma imberbe (formerly Beardless Flycatcher) L 3V2" RANGE: Breeds from southeastern Arizona, extreme southwestern New Mexico in Guadalupe Canyon, and Kenedy County in Texas south to Costa Rica. Winters in Mexico and Central America, casually to southern Arizona. STATUS: Fairly common to rare. HABITAT: In Arizona, occurs in cottonwoods, dense mesquite thickets, and in sycamore-live oak-mesquite associations. Along the lower Rio Grande Valley in Texas, inhabits mesquite woodlands, cottonwoods, willows, elms, and great luecaenas. NEST: Typically nests far out on a horizontal limb of a bush or tree up to 50 feet high, but usually near the ground. Usually nests along the edge of a grove, or among scattered trees in flat, sandy lowlands. Locates nest in a clump of mistletoe or sometimes between the bases of stems of palmetto fans. FOOD: Perches on the top branch of a tree or in lower branches to flycatch and glean scale insects, caterpillars, and ants. Occasionally eats small berries and seeds. REFERENCES: Bent 1942, Harrison 1979, Oberholser 1974a, Phillips et al. 1964, Terres 1980. 255 Olive-sided Flycatcher Contopus borealis L 6Vi" RANGE: Breeds from western and central Alaska and central Yukon to northern Ontario, south-central Quebec, and southern Labrador south to southern California across to western Texas, and east of the Rocky Mountains, to central Saskatchewan, northern Wisconsin, northeastern Ohio, and Massachusetts; also locally in the Appalachians to western North Carolina. Winters in South America and, casually, in southern California. STATUS: Local to fairly common. HABITAT: Inhabits montane and northern coniferous forests up to 10,000 feet in elevation, especially in burned-over areas with tall standing dead trees. Prefers forests of tall spruces, firs, balsams, and pines; groves of eucalyptus and Monterey cypress; taiga; subalpine coniferous forests; mixed woodlands near edges and clearings; and wooded streams and borders of northern bogs and muskegs. Prefers stands with a low percentage of canopy cover. SPECIAL HABITAT REQUIREMENTS: Tall, exposed perches such as snags or high, conspicuous dead branches. NEST: Usually hides nests in a cluster of needles and twigs on a horizontal branch of a conifer, well away from the trunk, usually between 15 and 50 feet above the ground. FOOD: Typically perches in tree tops and on high exposed limbs to hawk flying insects. REFERENCES: Bent 1942, Beal 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Terres 1980. 256 Greater Pewee Contopus pertinax (formerly Coues’ Flycatcher) RANGE: Breeds from central Arizona and southwestern New Mexico south to Central America. Winters in Mexico and Central America, casually north to southern Arizona. STATUS: Fairly common. HABITAT: Occurs in mountains up to 10,000 feet near the Mexico border, where it inhabits pine and pine-oak forests with an undergrowth of bushes. Also occurs in sycamore groves along mountain canyons. SPECIAL HABITAT REQUIREMENTS: Tall trees for feeding perches and for nesting. NEST: Locates nest on a horizontal fork 10 to 40 feet above the ground in a pine, sycamore, spruce, maple, oak, or other tall tree. Vigorously defends nest against jays, hawks, squirrels, and snakes. FOOD: Hawks insects from a perch high up in a tall tree. (There are no detailed reports of its food habits.) REFERENCES: Bent 1942, Harrison 1979, Phillips et al. 1964, Terres 1980. 257 Western Wood-Pewee Contopus sordidulus RANGE: Breeds from east-central Alaska, southern Yukon, and southern Mackenzie to northwestern Minnesota, south to Mexico, and east to western South Dakota, western Kansas, and western Texas. Winters in South America. STATUS: Common. HABITAT: Occurs in a variety of habitats including open deciduous and coniferous montane forests, pine-oak woodlands, floodplain forests, and wooded canyons. Found from sea level to the tops of coastal ranges, in cultivated stream valleys, in deciduous trees along borders of lakes and streams, in cities and towns, and in open, mature pine forests. It is generally adapted to drier environments than the eastern wood-pewee, and uses areas dominated by conifers. NEST: Locates nest on a horizontal limb or fork, dead or live, in a large variety of trees, usually deciduous, generally 15 to 40 feet above the ground. FOOD: Catches most of its food by hawking from a perch such as a dead branch. Eats insects, spiders, and a few wild berries. REFERENCES: Beal 1912, Harrison 1979, Johnsgard 1979, Terres 1980. 258 Eastern Wood-Pewee Contopus virens RANGE: Breeds from southeastern Saskatchewan to southern Quebec and New Brunswick, south to Texas, the Gulf Coast, and central Florida, and west to the eastern Dakotas, central Oklahoma, and south-central Texas. Winters in South America. STATUS: Common. HABITAT: Generally associated with deciduous forests; prefers woodlands with a relatively open understory but will use areas with a dense understory if the canopy above is incomplete or sparse. Also inhabits mixed forests, bottomlands, uplands, woodlots, orchards, parks, roadsides, and suburban areas planted to trees. Occurs in floodplain and river-bluff forests at the western edge of its range. Appears to be strongly associated with oaks, and throughout its range probably requires a predominance of hardwoods. NEST: Locates nest on a horizontal limb usually well out from the trunk, 9 to 65 feet above the ground, often on a dead limb in a living tree. Camouflages nest with spiderwebs and lichens. FOOD: Prefers to flycatch in a shady spot from mid to low level of the tree canopy. Eats insects, spiders, and millipedes, and also a few berries and seeds. REFERENCES: Beal 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979. 259 Yellow-bellied Flycatcher Empidonax flaviventris L 4V 2 " RANGE: Breeds from northern British Columbia and west-central and southern Mackenzie to southern Labrador and Newfoundland south to central Alberta, northern North Dakota, and northern Minnesota, across to southern Ontario, northeastern Pennsylvania, and Nova Scotia. Winters from Mexico to South America. STATUS: Common. HABITAT: Prefers predominantly coniferous forests of spruce and fir, frequenting low, swampy thickets bordering ponds and streams, spruce, cedar, tamarack and sphagnum bogs, spruce and alder swamps, wet mossy glades, and cool moist mountainsides. SPECIAL HABITAT REQUIREMENTS: Low, wet areas within coniferous forest. NEST: Nests on or near the ground, sometimes at the base of a tree or in a cavity formed by upturned roots, but more often beside a hummock or mound and well-hidden in sphagnum moss or other vegetation. May also nest in a damp, mossy crevice of rocks, but always in a quiet, concealed site. FOOD: Perches and feeds close to the ground, mainly on insects, which form 97 percent of the diet; occasionally eats a few berries. REFERENCES: Beal 1912, Bent 1942, DeGraff et al. 1980, Forbush and May 1955, Terres 1980. 260 Acadian Flycatcher Empidonax virescens RANGE: Breeds from southeastern South Dakota, northern Iowa, and extreme southeastern Minnesota to southern New York, Vermont, and Massachusetts, south to central and southern Texas, the Gulf Coast, and central Florida. Winters in Central and South America. STATUS: Common HABITAT: Inhabits the lowest tree canopy and understory layers of shady, humid riverbottom forests and wooded swamps. Prefers damp, lowland forests with an understory and uplands with wooded ravines near streams. Favors beech forests in the Northeast. SPECIAL HABITAT REQUIREMENTS: Mature, extensive deciduous forests with tall trees, a closed canopy, and open spaces in understory for feeding. NEST: Nests on a fork of a horizontal branch well away from the main trunk, usually 10 to 20 feet above the ground, often along a stream and t sometimes over water. Prefers open space below the nest to approach the nest easily. Favors lower branches of beech, dogwood, and witch- hazel, but also nests in oak, hickory, maple, basswood, and cherry. Occasionally is parasitized by brown-headed cowbirds. FOOD: Eats mostly insects; also eats some spiders and millipedes, and occasionally a few seeds and berries. REFERENCES: Beal 1912, Bent 1942, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Mumford 1964. 261 Alder Flycatcher Empidonax alnorum WILLOW AND ALDER FLYCATCHERS RANGE: Breeds from central Alaska and central Yukon to central and eastern Quebec, southern Labrador and southern Newfoundland, south to south-central British Columbia and southern Alberta, across to south- central Minnesota, eastern Pennsylvania, and Connecticut; also in the Appalachians south to western North Carolina. Winters in South America. STATUS: Common. HABITAT: Typically inhabits northern alder swamps, where it occupies a variety of habitats, including thickets of aspen parkland. Usually found near water in dense, low, damp thickets of alders, willows, sumacs, viburnum, elderberries, and red-osier dogwood bordering bogs, swamps, marshes, and along the banks of small streams and shores of ponds. SPECIAL HABITAT REQUIREMENTS: Forest openings and edges with dense, low shrubs. NEST: Nests in low trees or shrubs including dogwood, blackberry, hawthorn, viburnum, willow, spiraea, or alder, 1 to 6 feet above the ground, in an upright fork or saddled on a branch. FOOD: Catches at least 65 species of beetles as well as other flying insects, spiders, and millipedes; eats some fruits. REFERENCES: Bent 1942, DeGraff et al. 1980, Eckert in Farrand 1983b. Mousley 1931, Stein 1958, Terres 1980. Willow Flycatcher Empidonax traillii % RANGE: Breeds from central British Columbia and southern Alberta east to southern Wisconsin, southern Quebec, central Maine, and Nova Scotia south to southern California, western and central Texas, Arkansas, northern Georgia, and central and eastern Virginia. Winters in Mexico and Central America. STATUS: Common; population is generally stable or increasing throughout its range. HABITAT: Occurs in a variety of habitats ranging from brushy fields to willows, thickets along streams, prairie woodlots, shrubby swales, and open woodland edges. Prefers edge habitats that include thickets or groves of small trees and shrubs surrounded by grasslands, as well as the edges of gallery forests along rivers or streams. In areas where its range overlaps that of the alder flycatcher, prefers drier, smaller, more open shrubby habitat. SPECIAL HABITAT REQUIREMENTS: Fairly open and well-spaced shrubby habitats. NEST: Nests usually in horizontal forks or upright crotches of shrubs or small trees, usually between 3 to 25 feet above the ground, averaging about 4 to 6 feet. Commonly nests in dogwood, hawthorn, willow, buttonbush, elder, viburnum, and blackberry. Places nest at the outer edge of a shrub or thicket, so it can be easily approached. FOOD: Eats flying insects. REFERENCES: Bent 1942, DeGraff et al. 1980, Eckert in Farrand 1983b, Holcomb 1972, Johnsgard 1979, King 1955, Stein 1958, Tate and Tate 1982, Walkinshaw 1966. 263 Least Flycatcher Empidonax minimus RANGE: Breeds from southern Yukon and west-central and southern Mackenzie to southern Quebec and New Brunswick, south to southern British Columbia and central Montana, across to southwestern Missouri, northern Ohio, and central New Jersey; in the Appalachians to north¬ western Georgia. Winters in Mexico and Central America; also casually in southern California, southern Texas, and Florida. STATUS: Common; population is declining slightly in parts of its range. HABITAT: Associated with open deciduous forests, where it occurs along forest edges, burns, and clearings, floodplain forests, open shrublands, wooded margins of lakes and roads, orchards, shelterbelts, overgrown pastures, urban parks, and gardens. SPECIAL HABITAT REQUIREMENTS: Intermediate openness in the understory of open deciduous woodlands, and some edge habitat for nesting and feeding. NEST: Nests in upright crotch or on horizontal fork of deciduous or coniferous trees, usually saplings or small trees, including birch, red pine, cedar, apple, dogwood, oak, sugar maple, willow, and alder. Tends to nest at the edge of a clearing 10 to 20 feet above the ground, but will nest from 2 to 60 feet. FOOD: Feeds mainly on flying insects, most of which are caught on the wing, but some are gleaned from vegetation. Also takes a few fruits and seeds. REFERENCES: Beal 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, de Kirilene 1948, Tate and Tate 1982. 264 Hammond’s Flycatcher Empidonax hammondii RANGE: Breeds from east-central Alaska, southern Yukon, and south¬ western Alberta to northwestern Wyoming, south to east-central Cali¬ fornia, central Utah, northeastern Arizona, and north-central New Mexico. Winters in southeastern Arizona, Mexico, and Central America, casually in southern California. STATUS: Common. HABITAT: Inhabits tall, moist, closed-canopy montane conifer forests, sometimes with a deciduous understory. In Colorado, occurs from 7,500 to 11,000 feet in conifer-aspen woodlands; in California, in mature forests of medium to high canopy coverage from ponderosa pine up to lodge- pole pine forests. In the far North, prefers deciduous forests. SPECIAL HABITAT REQUIREMENTS: Nest sites that are cool and well shaded. NEST: Nests in a fork of a small tree or on a horizontal branch of a large conifer or deciduous tree, usually 25 to 40 feet above the ground. Uses birch, maple, ponderosa pine, western larch, and Douglas-fir for nesting. FOOD: Prefers to forage from the middle portions of tall conifers and aspens on flying insects. REFERENCES: Bailey in Farrand 1983b, Beaver and Baldwin 1975, Davis 1954, Terres 1980, Verner and Boss 1980. 9 265 Dusky Flycatcher Empidonax oberholseri l w RANGE: Breeds from southwestern Yukon, southern Alberta, southwest¬ ern Saskatchewan, and western South Dakota south to southern Cali¬ fornia, southern Nevada, central Arizona, and central and northeastern New Mexico. Winters in southern Arizona and Mexico: casual in southern California. STATUS: Common. HABITAT: Prefers shrubby sites or low- to intermediate-density forests with substantial shrub understory; generally avoids forests with a high percentage of canopy cover. Occurs in a variety of habitats, from mon¬ tane chaparral to moderately dense lodgepole pine forest, including many montane conifer types and aspen; especially favors mixed wood¬ lands or edge of small conifers and brush. In general, prefers drier, more open or patchier forests than Hammond’s flycatcher. NEST: Builds nests on upright or pendant twigs or in crotches of low shrubs or trees in relatively dry sites. Usually nests 3 to 8 feet, but up to 40 feet, above the ground in willow, alder, aspen, and other trees and shrubs. FOOD: Forages low over shrubby vegetation for flying insects. REFERENCES: Bailey in Farrand 1983b, Harrison 1979, Johnsgard 1979, Verner and Boss 1980. 266 Gray Flycatcher Empidonax wrightii RANGE: Breeds from south-central Washington and south-central Idaho to central Colorado, south to south-central California, central Arizona, and south-central New Mexico. Winters in central Arizona and Mexico, rarely in southern California. STATUS: Fairly common. HABITAT: Associated with arid woodland and brushy areas, where it inhabits tall sagebrush plains, pinyon-juniper woodlands, and arid, very open pine woods. During migration and in winter it occurs in arid scrub, riparian, and mesquite woodlands. NEST: Nests in a crotch of a thornbush, juniper, or sagebrush, 2 to 5 feet above the ground, sometimes in loose colonies. FOOD: Does most of its foraging in the spaces between bushes, and often flies to snatch insects from the ground. Catches insects from the size of tiny beetles to butterflies. REFERENCES: Bailey in Farrand 1983b, Phillips et al. 1964, Russell and Woodbury 1941, Terres 1980. * 9 267 Western Flycatcher Empidonax difficilis RANGE: Breeds from southeastern Alaska, northwestern and central British Columbia, and southwestern Alberta to western South Dakota, south along the coast and mountains to southwestern California, central Nevada, central and southeastern Arizona, and western Texas. Winters in Mexico. STATUS: Common. HABITAT: Found in a variety of wooded habitats; prefers moist, shaded forests, either coastal or lower montane, or higher in the Rockies and Great Basin ranges. Also inhabits hollows, canyon bottoms, riparian woodlands, and deciduous, coniferous, and mixed forests. SPECIAL HABITAT REQUIREMENTS: A sheltered nest site, possibly near a water source such as a stream, spring, or seep. NEST: May use a variety of sites for nesting; these include rock ledges or crevices of canyon walls, often concealed by ferns or clumps of mosses; crotch or tree limb projecting from the main trunk; behind flaps of loose bark; tree cavities; or old buildings. Nest height ranges from ground level up to 30 feet. FOOD: Commonly forages within shaded forests for insects and spiders. Also eats a few seeds. REFERENCES: Bailey in Farrand 1983b, Beal 1912, Beaver and Baldwin 1975 Davis et al. 1963, Johnsgard 1979, Verner and Boss 1980. 268 Buff-breasted Flycatcher Empidonax fulvifrons RANGE: Breeds very locally from the Huachuca and Chiricahua Mountains of east-central and southeastern Arizona through Mexico. Winters in Mexico. STATUS: Rare and local. HABITAT: Prefers open stands of pines and riparian trees, but also occurs in mixed pine and oak woods with shrubby undergrowth and on steep canyon slopes, from 5,000 to 8,500 feet. Favors open trees with bare, weedy, or grassy places. SPECIAL HABITAT REQUIREMENTS: Forest openings. NEST: Places nest on a branch 9 to 45 feet above the ground, often sheltered by an overhanging stub of a branch. Builds nests in pines, oaks, and sycamores. FOOD: Often forages for insects from the low scrubby understory or close to the trunk, low or at midlevel in pines. : REFERENCES: Bent 1942, Cottam and Knappen 1939, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b. 269 Black Phoebe Sayornis nigricans RANGE: Resident from northwestern California, southern Nevada, southwestern Utah, south-central Colorado, and western and west-central Texas south to South America. (Partially migratory, northern populations wander after the breeding season). STATUS: Common. HABITAT: Occurs in a variety of open and wooded habitats, usually near water, especially in drier regions. Found in the vicinity of streams, canals, lake margins, reservoirs, or other riparian sites flanked by grasslands, scattered trees, open woodlands or farmland. During migration and in winter, found at almost any source of water. SPECIAL HABITAT REQUIREMENTS: Vertical surface for nesting that is protected from rain, near a mud source, and preferably near water. NEST: Builds a mud nest commonly under bridges, in culverts and wells, under the eaves of buildings as well as on the vertical surfaces of cliffs, rimrocks, steep creek banks, and caverns. May use the same nest site year after year. FOOD: Catches insects over grassy fields and open water, or gleans them from plant foliage. Also eats some fruits of buckthorn. REFERENCES: Ohlendorf 1976, Phillips et al. 1964, Terres 1980. 270 Eastern Phoebe Sayornis phoebe L 5%" » RANGE: Breeds from northeastern British Columbia and west-central and southern Mackenzie to southwestern Quebec and central New Brunswick, south to southern Alberta, southwestern South Dakota, central New Mexico, and central and northeastern Texas acioss to northern Georgia and North Carolina. Winters from central Texas, the Gulf States, and Virginia south to Mexico and southern Florida; casually from Oklahoma, southern Ontario, and New England. STATUS: Common. HABITAT: Generally occurs near fresh running water in partially wooded habitats; frequents woodland edges, wooded ravines and cliffs, farms, and suburban areas where natural or artificial ledges are available for nesting. SPECIAL HABITAT REQUIREMENTS: Cliffs or ledges at stream-side clearings, or structures at forest openings for nesting. Perches 5 to 15 feet high for feeding. NEST: Nests on a ledge, usually sheltered above by an overhang, on natural or artificial structures, and generally near lakes or streams. May nest under bridges, culverts, or eaves of buildings, on cliffs, rock bluffs, or in ravines. Frequently uses nests from previous years, but is very adaptable in its nesting habits. Frequently the victim of cowbird parasitism. FOOD: In late winter and early spring, subsists mainly on seeds and wild berries; at other times, feeds on insects, usually hawking them from a perch. REFERENCES: Beal 1912, DeGraff et al. 1980, Forbush and May 1955, Hespenheide 1971, Johnsgard 1979, Tate and Tate 1982, Weeks 1979. 271 Say’s Phoebe Sayornis saya L 6>/4" RANGE: Breeds from western and northern Alaska, northern Yukon, northwestern and central Mackenzie, and central Alberta to southwestern Manitoba, south between coastal ranges and central prairie states to Mexico. Winters from northern California, Arizona, central New Mexico, and central Texas south to Mexico. STATUS: Common. HABITAT: Inhabits open arid regions, occurring in dry, barren foothills, sagebrush plains, scrubby pine-oak-juniper woodlands, bluffs and cliffs of the badlands, grasslands, canyons, and open areas near buildings. Unlike the eastern phoebe, it is independent of surface water. NEST: Prefers to nest in holes, crevices, on ledges, and on other protected horizontal surfaces of cliffs, rimrocks, steep creek banks, and caverns. Frequently nests in abandoned mine shafts, buildings, and under bridges. Also uses old nests of cliff swallows, barn swallows, and black phoebes; often uses the same nest in subsequent years or for successive clutches. FOOD: Eats mostly insects (primarily grasshoppers) and some berries. REFERENCES: Beal 1912, Bent 1942, Johnsgard 1979, Ohlendorf 1976, Terres 1980. 272 t Vermilion Flycatcher Pyrocephalus rubinus L 5" RANGE: Breeds from southern California, southern Nevada, central Arizona, central New Mexico, and western Oklahoma south to South America. Winters from southern California and southern Nevada to the Gulf Coast, east to south-central Florida, and south to Central America. STATUS: Common. * HABITAT: Found in the arid Southwest, occuring almost exclusively near water. Favor wooded groves of cottonwood, willow, oak, mesquite, and sycamore bordering rivers, especially near open, brushy, grassy, or * agricultural fields. Also occurs in widely spaced junipers and oaks, and in dry washes on the plains. NEST: Builds nest on a small, horizontal forked branch usually 8 to 20 , feet, but sometimes 40 to 50 feet, above the ground, and usually near a stream or other source of water. Nests in willow, sycamore, mesquite, cottonwood, oak, paloverde, hackberry, and other trees and bushes. • FOOD: Forages from a conspicuous perch, often only a few feet above the ground or water, for insects such as bees, grasshoppers, and small beetles. REFERENCES: Bent 1942, Harrison 1979, Johnsgard 1979, Taylor and Hanson 1970, Terres 1980, Terrill in Farrand 1983b. 273 Dusky-capped Flycatcher Myiarchus tuberculifer (formerly Olivaceous Flycatcher) L 5 3 A‘ RANGE: Breeds in southeastern Arizona and southwestern New Mexico; also in Central and South America. Winters from Mexico to South America. STATUS: Fairly common. HABITAT: Generally found below 6,000 feet but does occur up to 7,500 feet in montane pine-oak woodlands. Prefers dense scrub oak thickets on hillsides but also occurs along canyon streams where trees grow thick enough to provide deep shade. SPECIAL HABITAT REQUIREMENTS: Natural cavities or old woodpecker holes in trees for nesting. NEST: Builds nests in natural cavities in trees and stumps or in old woodpecker holes, 4 to 50 feet above the ground, in oaks, sycamores, or ashes. FOOD: Eats small insects. REFERENCES: Bent 1942, Cottam and Knappen 1939, Harrison 1979, Phillips et al. 1964, Terres 1980. 274 Ash-throated Flycatcher Myiarchus cinerascens L 6 V 2 " » * RANGE: Breeds from northwestern Oregon and eastern Washington to Colorado and western Kansas, south to Mexico. Winters from southern California and central Arizona south to Central America. STATUS: Common. HABITAT: Inhabits mesquite and cactus deserts, rocky mesas, shrubby canyons, oak groves on hillsides, mesquite thickets along creek bottoms, open pinyon-juniper woodlands, and open groves of sycamore, oak, willow, or cottonwood along stream courses. Stands with a low percent¬ age of canopy cover are preferred. Occurs from sea level to 9,000 feet in California, but is most frequently found at lower elevations. SPECIAL HABITAT REQUIREMENTS: Natural tree cavities or old wood¬ pecker holes for nesting. NEST: Uses a variety of cavities for nesting; natural cavities or knot holes in trees and stumps of mesquite, ash, oak, sycamore, juniper or cottonwood, or old woodpecker holes. May also nest behind loose pieces of bark, in abandoned nests of cactus wrens, in cavities in saguaro, in artificial structures, or in stalks of yucca or agave. Usually nests less than 20 feet above the ground. FOOD: Forages over low shrubs, hawking insects and spiders. Also eats a few fruits and seeds. REFERENCES: Beal 1912, Bent 1942, Johnsgard 1979, Terres 1980. 275 Great Crested Flycatcher Myiarchus crinitus RANGE: Breeds from east-central Alberta and central and southeastern Saskatchewan to south-western Quebec and central New Brunswick, south to central and southeastern Texas, the Gulf Coast, and Florida, and west to the eastern Dakotas, western Kansas, and west-central Oklahoma. Winters in central and southern Florida and from Mexico to South America. STATUS: Common. HABITAT: Prefers fairly extensive hardwood forests but is commonly found in old orchards and woodlots in farming country, clearings in mixed and deciduous forests, and wooded residential areas. Prefers forests with mature trees and fairly open canopies but will also use second-growth woodlands. SPECIAL HABITAT REQUIREMENTS: Cavities in middle-aged to mature trees, preferably in deciduous forests. NEST: Nests in woodpecker holes or in natural cavities in live or dead trees, usually 10 to 20 feet, but sometimes from 3 to 75 feet, above the ground. May also use artificial structures such as bird houses and other hollows, with little preference shown for the shape of the opening or the cavity expanse. FOOD: Forages in the forest canopy and gleans a variety of insects and spiders from crevices in the bark of trees; also eats some fruits. REFERENCES: Beal 1912, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Mousley 1934a. 276 L 7V4" Brown-crested Flycatcher Myiarchus tyrannulus (formerly Wied’s Crested Flycatcher) % RANGE: Breeds from southeastern California, extreme southern Nevada, southwestern Utah, Arizona, and southwestern New Mexico south to Central America. Winters in Mexico and Central America; rarely in south¬ ern Florida. STATUS: Fairly common. HABITAT: Inhabits saguaro deserts, riparian deciduous woodlands, and shade trees in urban areas. In Texas, occurs in open woodlands of mes- quite, hackberry, and ash; in Arizona, frequents cottonwood, willow, and sycamore woodlands. SPECIAL HABITAT REQUIREMENTS: Natural tree cavities or aban¬ doned woodpecker holes for nesting. NEST: Builds nest in abandoned woodpecker holes in saguaro or in cavities in cottonwoods, sycamores, mesquite, or old fence posts 5 to 30 feet above the ground. Sometimes nests in bird houses. FOOD: Probably eats beetles, other flying insects, and some wild berries and fruits; has been observed eating hummingbirds. REFERENCES: Bent 1942, Gambona 1977, Oberholser 1974a, Phillips et al. 1964. 277 Great Kiskadee Pitangus sulphuratus (formerly Kiskadee Flycatcher) RANGE: Resident in southern Texas north to Webb County and the Corpus Christi area and south to South America. STATUS: Locally common. HABITAT: Found in the lower Rio Grande Valley, along river beds, streams, ponds, and stagnant watercourses among large trees of mes- quite, huisache, palm, baldcypress, and willow, with thick undergrowth of shrubs and small trees. Also inhabits groves and orchards and is found in trees around ranches and urban areas. NEST: Builds nest in trees or tall shrubs, usually 10 to 20 feet, but up to 30 feet, above ground in brushy sites, low woods, along streams, marshes, or lagoons, or in cultivated areas. May nest in mesquite, palm, pine, acacia, or in a thorny bush. FOOD: Fishes for minnows, tiny fishes, and tadpoles by perching on a low branch overhanging water and diving, barely breaking the water’s surface. Also eats water insects, flying insects, and some fruits and berries, especially in winter. REFERENCES: Bent 1942, Oberholser 1974a, Terres 1980. 278 Sulphur-bellied Flycatcher Jiyiodynastes luteiventris L 6%" RANGE: Breeds from southeastern Arizona to Central America. Winters in South America. STATUS: Fairly common. HABITAT: Inhabits riparian mountain canyons, from 5,000 to 7,500 feet in elevation, where sycamore, oak, walnut, Arizona cypress, and pine are common. SPECIAL HABITAT REQUIREMENTS: Natural cavities in trees or aban¬ doned northern flicker holes. NEST: Usually builds nest in a natural cavity, typically a knothole where a large branch has broken off and a cavity has rotted out, 20 to 50 feet above the ground in living sycamores. Occasionally nests in an old flicker hole or a nest box placed high in a tree. Constructs nest on top of a loose platform built inside the cavity. FOOD: Eats a variety of insects and spiders; also a few small fruits and berries. REFERENCES: Bent 1942, Cottam and Knappen 1939, Ligon 1971a, Phillips et al. 1964. 279 Tropical Kingbird Tyrannus melancholicus L 7" RANGE: Breeds from southeastern Arizona to South America. Winters in Mexico south to South America. STATUS: Uncommon and local. HABITAT: Inhabits groves of tall trees, especially cottonwoods, next to ponds or flowing streams at low elevations. Frequently occurs along with both the western and Cassin’s kingbirds. NEST AND FOOD: Has nesting and food habits similar to those of west¬ ern and Cassin’s kingbirds, with which it closely associates. REFERENCES: Kaufman in Farrand 1983b, Terres 1980. 280 Couch’s Kingbird Tyrannus couchii [split from Tropical Kingbird) RANGE: Resident from southern Texas north to Webb and Kenedy Counties, south to Central America. STATUS: Fairly common. HABITAT: Frequents the borders of woods, chaparral, and trees along lakes, ponds, rivers, and stagnant watercourses, where it inhabits groves of mesquite, ebony blackbead, retaima, granjena, persimmon, and thorny bushes. It also frequents urban areas. NEST: Nests on a branch or in a fork of a tree 8 to 20 feet above the ground in woodlands and brush, along marshy or brushy margins of lakes or rivers, and in cultivated areas. FOOD: Frequently perches on tall trees to hawk insects. REFERENCES: Bent 1942, Harrison 1979, Kaufman in Farrand 1983b, Oberholser 1974a. 281 Cassin’s Kingbird Tyrannus vociferans RANGE: Breeds from central California, southern Utah, Colorado, and southeastern Montana south to Mexico, and east to western Texas. Winters in Mexico and Central America; irregularly from central California. STATUS: Fairly common. HABITAT: Occurs in open country such as plains and semideserts, in a variety of habitats from desert riparian areas up to 7,500 feet and in open woodlands in southwestern mountains. Inhabits pinyon-yucca, pinyon-juniper, pine-oak, and ponderosa pine woodlands, canyons of sycamores, and in California, open valley woodlands and grasslands of the foothills among scattered oaks, cottonwoods, and sycamores. SPECIAL HABITAT REQUIREMENTS: Tall trees for nesting. NEST: Usually nests in fairly tall trees such as pine, oak, cottonwood, walnut, hackberry, or sycamore. Places nest near the end of a horizonta limb 8 to 40 feet, but up to 100 feet, above the ground. Also places nests in bushes and on posts. FOOD: Primarily eats insects, but also spiders and fleshy fruits. REFERENCES: Beal 1912, Bent 1942, Hespenheide 1964, Johnsgard 1979, Ohlendorf 1974, Terres 1980, Terrill in Farrand 1983b. 282 Thick-billed Kingbird Tyrannus crassirostris L 7/a RANGE: Breeds from the Patagonia and Guadalupe mountains in southeastern Arizona, and Guadalupe Canyon in extreme southwestern New Mexico south to Mexico. Winters in Mexico. STATUS: Rare; first discovered in the United States in 1958, the range of this Mexican species has expanded northward since the middle of the 20th century. HABITAT: Occura near sycamore trees in streamside habitats dominated by cottonwood, willow, and mesquite. NEST: In the United States, nests in streamside sycamores 50 to 60 feet above the ground. FOOD: Presumably eats insects; commonly flies great distances between perches. REFERENCES: Levy 1959 , Oberholser 1974a, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b. 283 Western Kingbird Tyrannus verticalis RANGE: Breeds from southern interior British Columbia to southern Manitoba and western Minnesota south to Baja California, Mexico, and southern and south-central Texas; rarely or sporadically eastward to southern Ontario, Missouri, Arkansas, and Louisiana. Winters in small numbers along the Atlantic and Gulf Coasts from South Carolina to southern Florida and west to southern Louisiana; and from Mexico to Central America. STATUS: Common. HABITAT: Occurs in almost any open habitat with scattered trees at low to moderate elevations, especially in agricultural regions. Commonly occurs near edge habitats such as shelterbelts, woodland borders, orchards, and hedgerows. NEST: Builds nests in a variety of sites but prefers trees, when available May nest against the trunk, in a crotch, or on a horizontal branch 8 to 40 feet above the ground in cottonwoods, oaks, sycamores, willows and other trees; if no trees are available, nests in bushes, on utility poles, or a variety of structures. FOOD: Flycatches from a perch on poles, fence posts, or tree tops in open areas for a variety of insects; also eats millipedes, spiders, and some fruits. REFERENCES: Beal 1912, Bent 1942, Hespenheide 1964, Johnsgard 1979, Ohlendorf 1974, Terres 1980, Terrill in Farrand 1983b. 284 Eastern Kingbird Tyrannus tyrannus RANGE: Breeds from southwestern and north-central British Columbia, southern Mackenzie, and central Manitoba to southern Quebec and New Brunswick, south to northeastern California, northern Utah, northwestern and central New Mexico, the Gulf Coast, and Florida. Winters in South America. STATUS: Common. HABITAT: Frequents open areas with scattered trees or tall shrubs; forest edges or hedgerows along pastures, swamps, marshes, fields, or highways; open country around orchards; brushy streamsides; and sometimes open woodlands. SPECIAL HABITAT REQUIREMENTS: Open habitats with perches for flycatching. NEST: Often builds nest over water on a tree limb well away from the main trunk, or occasionally in shrubs or on an artificial structure, locating nest 10 to 20 feet, but sometimes 2 to 60 feet, above the ground. Builds nest in the crotch of a tree, on top of a dead stub, or on a fence post if no trees are available. In New England, frequently nests in the upper horizontal limbs of apple trees. FOOD: Consumes over 200 kinds of insects and more than 40 kinds of fruits, catching most insects by hawking from a perch. REFERENCES: Beal 1912, Bent 1942, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Terres 1980. 285 Gray Kingbird Tyrannus dominicensis RANGE: Breeds along the Atlantic and Gulf Coasts from southeastern South Carolina south to the Florida Keys, and west to southern Alabama and islands off the coast of Mississippi, south in coastal regions to South America. Winters from Hispaniola south to South America, casually in southern Florida. STATUS: Locally common. HABITAT: Found within the coastal zone, where it occurs in mangrove swamps, marsh edges, along roadsides, and in woodlands, groves, and yards in urban or rural areas. SPECIAL HABITAT REQUIREMENTS: Habitats within the coastal zone. NEST: Nests in a fork or saddle on a horizontal limb of a tree or shrub 3 to 17 feet above the ground, often hanging over water. Prefers mangroves for nesting, but also nests in oaks, acacia, sea grape, casuarina, and cabbage palm. Shows a strong attachment to the nesting site, returning yearly to the same tree or clump of trees. FOOD: Flycatches from exposed perches for insects; also eats lizards and fruits of tropical trees. REFERENCES: Bent 1942, Harrison 1975, Sykes in Farrand 1983b, Terres 1980. 286 Scissor-tailed Flycatcher RANGE: Breeds from southeastern Colorado, southern Nebraska, and north-central Missouri south to western and southern Texas and western Louisiana; isolated breeding in northeastern Mississippi, central Tennessee, and central Iowa. Winters in southern Florida and from Mexico to Central America; casually in southern Louisiana. Tyrannus forficatus STATUS: Common. HABITAT: Occurs on plains, prairies, mesas, and flats, and around pastures, woodland clearings, ranches, and farms. Perches for long periods on tall prairie plants, limbs of dead trees, utility wires, or fences. SPECIAL HABITAT REQUIREMENTS: Open habitats with elevated perches. NEST: Typically nests in cottonwoods, elms, or other hardwood species, in exposed sites 6 to 30 feet above the ground. Prefers isolated trees to those growing in clumps or heavier cover; occasionally uses fence posts, telephone poles, windmill towers, or buildings for nest sites. FOOD: Flies from a perch to catch a variety of insects, but also picks up insects from the ground. Also eats few fruits, berries, and seeds. REFERENCES: Beal 1912, Fitch 1950, Johnsgard 1979, Oberholser 1974a. 287 Rose-throated Becard RANGE: Breeds in southeastern Arizona and in southern Texas (Cameron and Hidalgo Counties) and in Mexico and Central America. Winters in Mexico and Central America. STATUS: Rare and local. HABITAT: Inhabits mature groves of trees situated near flowing water, preferably stands of sycamore, cottonwood, and willow. NEST: Builds an immense bushel-basket nest of strips of fibrous plant stems, suspending it from twigs at the end of a drooping branch 30 to 60 feet above the ground. Often places nests in sycamores but also uses cottonwoods, baldcypress, and willows. Will often build in the same site as the previous year’s nest, or very close to the site. FOOD: Perches on interior branches rather than exposed perches while feeding. Eats insects and some wild fruit. REFERENCES: Oberholser 1974b, Phillips 1949, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b. 288 Horned Lark Eremophila alpestris L RANGE: Breeds in North America from western and northern Alaska, the Arctic Coast of northern Canada, Prince Patrick, Devon, and Baffin Islands, and northern Labrador south to Mexico, southwestern Louisiana, central Missouri, northern Alabama, and North Carolina. Winters from southern Canada south throughout the breeding range, and locally or irregularly to the Gulf Coast and southern Florida. STATUS: Locally common. HABITAT: Inhabits a wide variety of open habitats, from coastal dunes and alpine tundra to prairies and deserts. Prefers areas with a minimum of vegetation, such as natural or planted low-stature grasslands, cultivated and plowed fields, golf courses, airports, and other relatively barren areas. In winter, groups in small-to-enormous flocks on open, barren sites similar to its breeding habitat. SPECIAL HABITAT REQUIREMENTS: Bare ground for nesting. NEST: Nests in a depression on the ground, placed so that the upper edge of the nest is level with the ground surface. Often paves the nest with small pebbles along a portion of the rim. Places nest where there is little or no vegetational cover around the nest, or next to a clump of grass or a rock. FOOD: In summer, feeds primarily on insects; in winter, it consumes seeds of grasses, weeds, and waste grains. REFERENCES: Beal and McAtee 1912, Beason and Franks 1974, Cottam and Hanson 1938, DeGraff et al. 1980, Johnsgard 1979, Pickwell 1931, Terres 1980, Verbeek 1967. 289 Purple Martin Progne subis RANGE: Breeds from southwestern British Columbia south to Baja California; and from northeastern and east-central British Columbia and central Alberta to southern Ontario and New Brunswick south to Mexico, the Gulf Coast, and southern Florida, and west to eastern Idaho and central Utah. Local in the Rocky Mountains but avoids most other mountainous areas. Winters in South America, casually in Florida. STATUS: Locally common; of special concern on blue list. HABITAT: Inhabits open and cut over woodlands, open grassy river valleys, meadows around pools, shores of lakes, marsh edges, agricul¬ tural lands, saguaro deserts, parks and towns. Prefers habitats near open water. In the East, breeds almost exclusively in artificial colonial martin houses; in the West, still uses woodpecker-made cavities to a large extent. SPECIAL HABITAT REQUIREMENTS: Large, multiroomed martin houses, tree cavities, or abandoned woodpecker holes for nesting, and open spaces for foraging. NEST: Originally nested in cavities in large snags but is now largely dependent upon man-made martin houses. Nests colonially in houses preferably set 15 to 20 feet above the ground in open settings near suitable perches such as wires. Also uses cavities in cliffs or among loose rocks, and crevices in old buildings. In the west, still depends on old woodpecker holes for nesting; in the Arizona deserts, nests in old woodpecker holes in saguaro cacti. FOOD: Catches flying insects on the wing for most of the diet. Also picks up a few insects and spiders from the ground. REFERENCES: Allen and Nice 1952, Beal 1918, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Tate and Tate 1982, Terres 1980. 290 Tree Swallow Tachycineta bicolor L 5" RANGE: Breeds from western and central Alaska and central Yukon to northern Quebec and central Labrador south along the Pacific Coast to southern California and south-central New Mexico, generally sporadic or irregular as a breeder east of the Rocky Mountain States and south of the upper Mississippi and Ohio Valleys, or along the Atlantic Coast south of Massachusetts. Winters from southern California, southwestern Arizona, Texas, the Gulf Coast, and the Atlantic Coast from New York south to Central America. STATUS: Common. HABITAT: Prefers open woodlands near ponds, small lakes, or marshes. Occurs around farmlands, river bottomlands, beaver ponds, wooded swamps, and marshes where dead standing trees are in or near water. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting; suitable cavity trees must have a minimum dbh of 10 inches, and open feeding areas such as meadows, marshes, or open water. NEST: Prefers to nest in natural cavities and abandoned woodpecker holes, but if nesting holes are scarce, will accept nest boxes placed in open fields or use crevices in buildings. Uses cavities in the trunk or limb of live or dead trees, especially if the cavity is 3 to 15 feet above water. Usually nests singly but is loosely colonial if there are abundant suitable cavities and a good food supply. FOOD: Feeds mostly on flying insects over open areas. In early spring and in cold weather when insects are scarce, subsists on wild berries and seeds, especially bayberries. REFERENCES: Beal 1918, Chapman 1955, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Scott et al. 1977, Terres 1980, Thomas et al. 1979. 291 Violet-green Swallow Tachycineta thalassina l w RANGE: Breeds from central Alaska, central Yukon, and extreme southwestern Mackenzie south to Mexico, and east to southwestern Saskatchewan, western South Dakota, and western Nebraska. Winters from central coastal and southern California south to Central America. STATUS: Common. HABITAT: Inhabits coniferous, deciduous, and mixed forests, preferring open or broken woods, or the edges of dense woodlands. Occurs around towns, in woodland clearings, especially near lakes and streams, and if snags are present, in canyons, and in mountains from the foothills to near timberline. SPECIAL HABITAT REQUIREMENTS: Cavities or crevices for nesting and open terrain or forest openings for feeding. NEST: Builds nests in old woodpecker holes, natural tree cavities, crevices in rocky cliffs, nesting boxes, niches of old buildings, and when cavities are scarce, in old nests of cliff swallows and burrows of bank swallows. In Colorado, prefers cavities in ponderosa pine but also nests in aspen and other trees. FOOD: Consumes only insects, which catches and eats while on the wing. REFERENCES: Bailey and Niedrach 1965, Beal 1918, Bent 1942, Combellack 1954, Johnsgard 1979, Scott et al. 1977, Terres 1980. 292 Northern Rough-winged Swallow S telgidopteryx serripennis L W RANGE: Breeds from southeastern Alaska, central British Columbia, and southern Alberta to southwestern Quebec and central Maine south to Central America and south-central and southwestern Florida. Winters from southern Texas, southern Louisiana, and southern Florida south to Mexico and Central America. STATUS: Fairly common. HABITAT: Inhabits open country, including open woodlands, wherever a suitable nest site near water can be found. In the East, frequents rocky gorges, shale banks, stony road cuts, railroad embankments, river valleys, and stream banks. In the Midwest and West, often found around gravel pits, stream banks, and other exposed banks of sand, dirt, or gravel. SPECIAL HABITAT REQUIREMENTS: Suitable nest sites preferably near, but up to 1/2 mile from water. NEST: Excavates nests in banks of clay, sand, or gravel or uses abandoned bank swallow or kingfisher burrows and sometimes natural rock crevices, drainpipes, culverts, cracks in bridges, and crevices in buildings. May nest singly, in scattered groups, or in small colonies; tends to be more colonial in the western part of its range. FOOD: Feeds on the wing, catching primarily flies and other flying insects. REFERENCES: Beal 1918, DeGraff et al. 1980, Johnsgard 1979, Lunk 1962. 293 Bank Swallow Riparia riparia L4H" RANGE: Breeds from western and central Alaska and central Yukon to central Quebec and southern Labrador, south to southern California, western Nevada, southern New Mexico, southern Texas, northern Alabama, eastern Virginia, and casually, northwestern North Carolina and south-central South Carolina. Winters in South America. STATUS: Locally common; population is declining over parts of its range. HABITAT: Prefers grasslands and cultivated fields but uses a variety of open habitats, usually near water and suitable nest sites. Nests in riverbanks, borrow pits, gravel pits, road cuts, sand banks and other exposed banks of sand, gravel or clay. SPECIAL HABITAT REQUIREMENTS: Vertical banks of sand, gravel, or clay in an open habitat, preferably near lakes, ponds, or marshes. NEST: Excavates a burrow near the top of a vertical bank (or repairs an existing burrow) ranging from 9 inches to 6 feet, but generally about 2 feet, in length. Forms dense colonies, with up to several hundred nests in a bank. FOOD: Catches primarily flies while flying over water or grasslands, especially pastures. REFERENCES: Allen 1933, Beal 1918, Beyer 1938, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Peterson 1955, Tate and Tate 1982. I 294 Cliff Swallow Hirundo pyrrhonota 15" RANGE: Breeds from western and central Alaska and central Yukon to northern Ontario, southern Quebec and New Brunswick south to Mexico, southwestern Louisiana, northern portion of the Gulf States and southern North Carolina: also in the Lake Okeechobee region of southern Florida. Winters in South America. STATUS: Common in the West, locally fairly common in the East; overall populations are stable or increasing, except in some northeastern States where it is of special concern on the blue list for declining species. HABITAT: Originally restricted to the vicinity of cliffs and banks; now occurs over open country around farmlands, towns, bridges, dams, freeway overpasses, and other areas near mud supplies and potential nest sites. SPECIAL HABITAT REQUIREMENTS: A vertical substrate with an overhang for nest attachment, a supply of mud suitable for nest construction, fresh water with a smooth surface for drinking, and an open foraging area near the nest site. NEST: Originally nested on bluffs, cliffs, deep gorges in mountains, and sometimes on the side of large pine trees and in caves; has adapted to building its gourdlike mud nests under the eaves of, or in, buildings, under bridges, in culverts, on the face of dams, and under freeway overpasses. Forms colonies of up to several hundred nests in favorable locations. FOOD: Consumes insects caught while flying high, often above 100 feet, as nearly 100 percent of the diet. REFERENCES: Beal 1918, Bent 1942, DeGraff et al. 1980, Emlen 1954, Forbush and May 1955, Johnsgard 1979, Mayhew 1958, Samuel 1971, Tate and Tate 1982. 295 Cave Swallow Hirundo fulva L 4Vt" RANGE: Breeds from Carlsbad Caverns in southeastern New Mexico and from western and south-central Texas south through Mexico. Winter range is unknown. STATUS: Locally fairly common; range is expanding as it adapts to human-altered environment. HABITAT: Originally restricted to open country in the vicinity of limestom caves and sinkholes; has adapted its nesting habits to artificial struc¬ tures. Now also nests around culverts and bridges in the northern part c its range where water and mud are available. SPECIAL HABITAT REQUIREMENTS: Extensive roughened or pitted surfaces for nesting in caves, water for drinking, and mud suitable for nest construction. NEST: Forms colonies and in caves, tends to build its mud nests in isolated crevices and pockets, or under overhanging ledges. Also nests in sinkholes, in highway culverts, and under bridges. May reuse nest year after year and will sometimes share the same nest site with barn swallows. FOOD: Needs a source of water, such as a seep, spring, open water tank, or pond for drinking. (The diet of this swallow in the United States is currently unknown.) REFERENCES: Martin 1974, 1981, Selander and Baker 1957, Wauer and Davis 1972. 296 Barn Swallow Hirundo rustica L 6" RANGE: Breeds from south-coastal and southeastern Alaska and southern Yukon across to central Manitoba, northern Ontario, and southern Quebec south to Mexico, the Gulf Coast, north-central Florida, and southern North Carolina. Winters in Central and South America, casually north to the southwestern United States and southern Florida. STATUS: Common. HABITAT: Occurs virtually throughout the whole United States wherever suitable nest sites are found, but favors farmlands, open forests, rural, and suburban areas. SPECIAL HABITAT REQUIREMENTS: Overhead protection, especially buildings, for nesting. NEST: Originally nested on cliffs and in caves and rock crevices in mountains, along rocky coasts, and on high shores of lakes and rivers. Still uses such sites in the north and on the Pacific Coast, but in other areas nests on horizontal beams or ledges inside barns or other buildings, or under bridges, culverts, or wharves. Usually nests colonially. FOOD: Prefers to feed over water or fields, catching flying insects (especially flies) on the wing. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Samuel 1971. 297 Gray Jay Perisoreus canadensis RANGE: Breeds from western and central Alaska, northern Mackenzie, and southwestern Keewatin across to northern Quebec and northern Labrador, south to northern California, central Idaho, east-central Arizona, Black Hills of South Dakota, central Saskatchewan, northern Minnesota, southern Ontario, and northern New England. Winters generally throughout the breeding range. STATUS: Locally common. HABITAT: Inhabits northern coniferous forests, especially dense spruce and pine. Occasionally occurs in mixed forests and deciduous woodlands near coniferous forests. SPECIAL HABITAT REQUIREMENTS: Conifer forests. NEST: Builds nest in late winter while there is still deep snow in the woods, typically in a crotch or on a horizontal branch near the trunk of a conifer, often less than 10 feet but up to 30 feet above the ground. Usually hides well. FOOD: Regularly catchesfood, producing a special saliva that helps bind the food together, so it can be firmly held in conifer foliage. Is omnivorous and typically eats insects, conifer seeds, berries, young birds, small mammals, lichens, fungi, and carrion. Commonly steals food from campers. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Goodwin 1976, Johnsgard 1979, Ouellet 1970, Rutter 1969. 298 Steller’s Jay Cyanocitta stelleri Lir RANGE: Resident from south-coastal and southeastern Alaska to southwestern Alberta south to southern California, Arizona, central New Mexico, Central America, and western Texas, and east to western Montana and western Nebraska. STATUS: Common. HABITAT: Generally inhabits coniferous forests throughout its range, but occasionally occurs in mixed and deciduous woodlands, and ventures into orchards and gardens. Is especially frequent in the ponderosa pine zone but limited numbers occur in the pinyon-juniper and spruce-fir zones. Favors edges of forest openings over extensive, unbroken woodlands. SPECIAL HABITAT REQUIREMENTS: Predominately coniferous woodlands. NEST: Usually builds nest on a horizontal branch of a conifer from 8 to 40 feet, but up to 100 feet, above the ground. Occasionally locates nest in a shrub or tree cavity. FOOD: Usually forages on the ground and in trees for seeds, nuts, acorns, fruits, insects, spiders, and bird eggs and nestlings. REFERENCES: Gibson in Farrand 1983b, Goodwin 1976, Johnsgard 1979, Wilmore 1977. 299 Blue Jay Cyanocitta cristata L 10' RANGE: Resident from extreme east-central British Columbia and central and southeastern Alberta to southern Quebec and Newfoundland south to central and southeastern Texas, the Gulf Coast, and southern Florida, and west to eastern Montana and east-central New Mexico. Northern populations are partly migratory to the southern parts of the breeding range. STATUS: Common. HABITAT: Inhabits deciduous and mixed woodlands, especially prefering those with oak, beech, and hickory, but also occurs in coniferous forests, preferably where pines predominate. Also frequents wooded islands, farms, gardens, parks, cities — almost anywhere trees are found in grassland areas. NEST: Hides nest well in the fork, crotch, or outer branches of trees, occasionally in shrubs, typically from 10 to 25 feet, but ranging from 5 to 50 feet, above the ground. Prefers conifer thickets in mixed woodlands for nesting. FOOD: Forages from the tree tops to the ground for its food, which consists of 76 percent vegetable and 24 percent animal matter. Feeds omnivorously, primarily on mast but also takes a variety of other foods. Also eats grains, weed seeds, wild fruits, insects, a few mice, young birds and eggs, fish, salamanders, and crustaceans. Is easily drawn to bird feeders. REFERENCES: Beal 1904, DeGraff et al. 1980, Goodwin 1976, Johnsgard 1979, Wilmore 1977. 300 Green Jay Cyanocorax yncas RANGE: Resident in southern Texas; also from Mexico to South America. STATUS: Fairly common. HABITAT: Found in dense thickets and woodlands along the lower Rio Grande, in mesguite woodlands, and thickets of mesquite, retaima, and hackberry. In winter, wanders in small flocks in more open country, frequenting woods of huisache, ebony blackbean, and anacua. NEST: The nest is built in a fork or outer branches of small trees or bushes from 5 to 15 feet above the ground, in dense woods or thickets along streams. FOOD: Feeds omnivorously, including insects, spiders, seeds, acorns, palmetto fruit, and eggs and young of birds in the diet. REFERENCES: Cottam and Knappen 1939, Oberholser 1974b, Terres 1980, Wilmore 1977. 301 Scrub Jay Aphelocoma coerulescens RANGE: Resident from southwestern Washington to southern Wyoming and Colorado south to Baja California, western and west-central Texas, and Mexico: also on Santa Cruz in the Channel Islands in California, and in central Florida. STATUS: Locally common; population is declining in Florida because of clearing and development of orange groves. HABITAT: Inhabits a variety of brushy areas from dense chaparral, open woodlands and residential areas in the Pacific States to pinyon-juniper, scrub oak, and less frequently mixed oak and ponderosa pine in the interior West, and humid scrub-oak communities in Florida. Prefers borders of brushy ravines and wooded creek bottoms. In Florida, usually found near small openings or at the edge of the scrub rather than in dense, unbroken scrub. SPECIAL HABITAT REQUIREMENTS: Scrub habitats. NEST: Builds nest in pinyons, oaks, tall shrubs, or vine tangles, gener¬ ally less than 10 feet, but up to 30 feet, above the ground. Usually nests singly, but in Florida may nest in scattered colonies with up to 6 nests in small tracts of scrub. FOOD: Consumes many kinds of insects and other invertebrates. Also eats acorns, seeds of palmetto, grains, fruits, mice, eggs and nestlings of birds, small reptiles, and frogs. REFERENCES: Goodwin 1976, Flarrison 1975, Johnsgard 1979, Tate and Tate 1982, Terres 1980, Terrill in Farrand 1983b, Wilmore 1977. 302 Gray-breasted Jay Aphelocoma ultramarina (formerly Mexican Jay) RANGE: Resident from central Arizona, southwestern New Mexico, and western Texas south to Mexico. STATUS: Common. HABITAT: Inhabits canyons and hillsides of oak and pine-oak woodlands, adjacent riparian forests, and occasionally ventures into adjacent pure pine woodlands or more open scrub, from 2,000 to 9,000 feet in elevation. SPECIAL HABITAT REQUIREMENTS: Live oaks within its home range. NEST: Builds nest in a fork, crotch, or on a horizontal branch of oaks or pines, typically 10 to 25 feet, but ranging from 6 to 54 feet above the ground. May nest singly or in loose colonies, and participates in com¬ munal nest building. FOOD: Usually forages on the ground and in trees for acorns, insects, lizards, eggs and young birds, fruits, and seeds. REFERENCES: Goodwin 1976, Gross 1949, Terres 1980, Wilmore 1977. 303 Pinyon Jay Gymnorhinus cyanocephalus L 9' RANGE: Breeds from central Oregon to western South Dakota, south to Baja California, northwestern and east-central Arizona, central New Mexico, and western Oklahoma. Winters throughout the breeding range and irregularly from southern Washington to northwestern Montana south to Mexico and central Texas. STATUS: Common. HABITAT: Typically inhabits pinyon-juniper woodlands of the foothilfs and lower mountain ranges in the West, but also occurs in open ponderosa pine forests where the soil is dry and trees are small and scattered. Locally nomadic outside the nesting season in response to fluctuating food supplies; during years of a poor seed crop, may move in flocks hundreds of miles to find food. SPECIAL HABITAT REQUIREMENTS: Open woodlands for nesting and an adequate supply of seeds, especially pinyon nuts. NEST: Usually nests in scattered colonies of up to 100 birds, usually with just one nest per tree. Builds nest away from the center of a tree on a low, southerly facing horizontal limb, and generally 6 to 20 feet, but up to 85 feet, above the ground. Pinyons, junipers, ponderosa pines, and scrub oaks are common nest trees. FOOD: Forages in trees and on the ground, principally for seeds, espe¬ cially pinyon nuts, but also seeds of ponderosa pines and other conifers, and caches them for use during the next breeding season. It also eats fruits, berries, insects, and eggs and nestlings of small birds. REFERENCES: Baida and Bateman 1972, Goodwin 1976, Johnsgard 1979, Terres 1980, Wilmore 1977. 304 Clark’s Nutcracker Nucifraga columbiana RANGE: Resident from central British Columbia and southwestern Alberta to western and southeastern Wyoming, south through the mountains of Washington, Oregon, California, and Nevada to Baja California, and in the Rockies to east-central Arizona and southern New Mexico. Occasionally wanders during nonbreeding season to lower mountains and lowlands beyond breeding range. STATUS: Common. HABITAT: Prefers high-altitude rocky sites with open or broken coniferous forest and clearings, but occurs in the mountains from 3,000 to 13,000 feet. Inhabits a variety of coniferous forest types including ponderosa pine, pinyon-juniper, and spruce-fir. SPECIAL HABITAT REQUIREMENTS: Seeds of pines for food. NEST: Nests mostly between 6,000 and 8,000 feet in elevation in a wind- sheltered site. Builds nest in a conifer, well out on a branch or in a bushy top from 7 to 150 feet above the ground. FOOD: Forages on the ground and in trees, primarily on the seeds of conifers and on insects. Also eats seeds of lupine, oats, grains, berries, small mammals, eggs and nestlings of birds, and carrion. REFERENCES: Bevier in Farrand 1983b, Cottam 1945, Goodwin 1976, Mewaldt 1956, Terres 1980, Verner and Boss 1980, Wilmoer 1977. 305 Black-billed Magpie Pica pica RANGE: Resident from south-coastal and southern Alaska to northern Alberta, central Manitoba, and western Ontario south to northeastern and east-central California and south-central Nevada, across to western and northeastern Oklahoma. Casual north and east of range in fall and winter. STATUS: Common. HABITAT: Typically inhabits open country with short, scattered, clumped or grazed vegetation, or exposed ground, and with patches of scrub, large bushes, or trees. Avoids dense forests and strictly desert regions. Found nearly up to timberline in the mountains; frequents a variety of open habitats including sagebrush, agricultural lands, pastures, grasslands, forest edges, streamsides with tall thickets and scattered trees, open woodlands, and urban areas. SPECIAL HABITAT REQUIREMENTS: Open country for foraging, trees or large bushes for nesting and cover. NEST: Builds a bulky stick nest from a few feet to 25 feet above the ground in a variety of trees or tall bushes, especially thorny ones. Nests in small, scattered colonies along streams, in woods, or in thickets, less often on buildings, cliff ledges, high banks, or on the ground among bushy cover. Sometimes reuses the old nest but usually builds a new nest each year. FOOD: Forages mainly on the ground, sometimes in trees or shrubs, for insects, especially grasshoppers. Also eats snails, slugs, millipedes, spiders, fishes, reptiles, amphibians, young birds and eggs, small mammals, carrion, and wild and cultivated fruits. REFERENCES: Ballard in Farrand 1983b, Goodwin 1976, Johnsgard 1979, Kalmbach 1927, Linsdale 1937, Terres 1980, Wilmore 1977. 306 Yellow-billed Magpie Pica nuttalli Yellow-billed L 16" Black-billed L18" RANGE: Resident in California in the Sacramento and San Joaquin Valleys, and in valleys of the coast ranges from San Francisco Bay south to Santa Barbara County. STATUS: Common. HABITAT: In farming country, foothills, and valleys, inhabits broken oak woodland interspersed with grasslands or cultivated lands, open riparian woodland, oak savannah, and vacant city lots. Avoids areas with high strong winds, cold snowy winters, and very dry hot summers. Prefers tall trees in a linear arrangement, such as trees bordering streams, parklike groves, or orchards. SPECIAL HABITAT REQUIREMENTS: Tall trees near open water for nesting. NEST: Believed to mate for life; breeds in small loose colonies, with each pair in a different tree. Prefers tall trees usually about 50 feet. Builds nest near the tree top on a small limb far out from the trunk, sometimes in a mistletoe clump. Occasionally reuses nest, but usually builds a new one each year. Locates nest in sycamore, oak, cottonwood, and digger pine. FOOD: Consumes a diet of about half insects, especially grasshoppers, but also eats grains, acorns, cultivated and wild fruits, and carrion. REFERENCES: Goodwin 1976, Kalmbach 1927, Linsdale 1937, Terres 1980, Verbeek 1973, Wilmore 1977. 307 American Crow Corvus brachyrhynchos RANGE: Breeds from north-central British Columbia and southwestern Mackenzie to central Quebec and southern Newfoundland south to Baja California, central Arizona, southern New Mexico, central and southeast¬ ern Texas, the Gulf Coast, and southern Florida. Winters from southern Canada south throughout the breeding range. STATUS: Common. HABITAT: Most often inhabits open and semiopen habitats, favoring open deciduous, coniferous, and mixed forests, wooded river bottoms, groves, orchards, woodlands adjacent to agricultural land, suburban areas, parks, and woodlots. NEST: Builds a nest, a large platform of sticks, usually on a horizontal branch or in a crotch of a tree near the trunk, 10 to 75 feet above the ground. Prefers conifers and oaks as nest trees, but where trees are lacking, will build nests on the ground, on shrubs, or on telephone pole crossbars. FOOD: Prefers to forage in cultivated fields. Has an omnivorous diet that is three-fourths vegetable foods, including cultivated grains, seeds, wild and cultivated fruits, and nuts. Also eats insects, millipedes, spiders, small crustaceans, small reptiles, frogs, small mammals, eggs and young of birds, and carrion. REFERENCES: DeGraff et al. 1980, Goodwin 1976, Johnsgard 1979, Lehman in Farrand 1983b, Wilmore 1977. 308 Northwestern Crow 'orvus caurinus L RANGE: Resident along the Pacific Coast from south-coastal and southeastern Alaska south to the Puget Sound area in northwestern Washington. STATUS: Common. HABITAT: Occurs along the coast, rarely straying farther than a mile from tidal waters, where it inhabits saltwater beaches, towns, and the wooded shores of bays, especially where there are small coniferous trees. NEST: Nests in scattered pairs or in loose colonies, usually in a crotch of a low tree or bush 10 to 20 feet, but up to 70 feet, above the ground. Occasionally builds nest on the ground under overhanging boulders, under bushes or windfalls, on the side of a sandy bank, or in a hole in a cliff. Will locate nest in apple, hemlock, Douglas-fir, and spruce trees. FOOD: Scavenges refuse from beaches, along with mollusks and other shellfishes. Forages for insects, especially grasshoppers, in nearby cultivated fields. Also eats crabs, mussels, dead fish and other carrion, eggs of other birds, and wild and cultivated fruits. REFERENCES: Bent 1946, Goodwin 1976, Terres 1980, Wilmore 1977. 309 Fish Crow Corvus ossifragus RANGE: Resident from New York and Massachusetts south along the Atlantic Coast to southern Florida and west to southern Texas; inland along major river systems to southern Illinois and east-central Oklahoma. STATUS: Locally common; range is extending northward and into drier regions. HABITAT: Inhabits low coastal areas, especially wooded marine shore¬ lines, coastal marshes and beaches, brackish bays, fertile farmlands up to 100 miles from the coast, inland wetlands, and forests near rivers and lakes. Occasionally occurs in pine forests, orchards, old dry fields, and abandoned farmlands overgrown with natural grasses and pines. NEST: Nests singly or in loose colonies of 2 to 4 pairs, each nesting in a separate tree. Builds nest near the tops of trees, especially pines, usually 10 to 90 feet, but up to 150 feet, above the ground; rarely in tall shrubs. Usually locates nest near water in a large fork or on a horizontal limb close to the trunk. FOOD: Gathers food from the ground or trees, most commonly from tidal flats, beaches, rookeries, and riverbanks. Eats a diet that includes crabs, shrimps, and other crustaceans; stranded and dead fish; insects; eggs and young of birds; small reptiles; wild fruits; cultivated grains; seeds; and carrion. REFERENCES: DeGraff et al. 1980, Goodwin 1976, Johnsgard 1979, Wilmore 1977. 310 Chihuahuan Raven Corvus cryptoleucus L \7Vi RANGE: Resident from south-central and southeastern Arizona, central and northeastern New Mexico, northeastern Colorado, and south-central Nebraska south to Mexico, and east to western Kansas and central Texas. (Northeastern populations, especially those in Nebraska and Kansas, migrate southward in winter.) STATUS: Locally common. HABITAT: Favors open, arid grasslands interspersed with yucca, mesquite, and cactus. Also occurs in deserts, on the open plains, and in arid farmlands, extending into the foothills. Outside the breeding season, often forms large flocks and roosts communally in canyons and gulches. NEST: Usually builds nest in isolated trees or bushes, 4 to 40 feet above the ground; often uses same nest year after year. Uses sycamore, mesquite, willow, oak, yucca, cottonwood and other species for nest sites, but also nests on utility poles and windmill towers. FOOD: Typically eats insects, especially grasshoppers and beetles, cultivated grains, small reptiles, fruits of cacti, carrion, eggs and young of birds, and scraps of human food. REFERENCES: Bent 1946, Goodwin 1976, Johnsgard 1979, Terrill in Farrand 1983b, Wilmore 1977. 311 Common Raven L 21 Corvus corax RANGE: Resident from Alaska and northern Canada south through the western United States to Baja California and Mexico, and east to the eastern edge of the Rockies, western Oklahoma, and central Texas; east of the Rockies, south to central Saskatchewan, northern Wisconsin, southern Ontario, Vermont, and southeastern Maine; also locally in the Appalachians to northwestern Georgia. STATUS: Common to locally common; reinvading its historic range and colonizing new areas. HABITAT: Occurs in a wide variety of habitats but is most often found in open woodlands and mountainous and coastal regions. Inhabits rocky seacoasts, steep canyons, boreal forests, deserts, foothills, mountains, arctic tundra, and wooded marine islands. Tends to avoid extensive, dense forests. SPECIAL HABITAT REQUIREMENTS: Cliff ledges or tall trees for nesting. NEST: Usually builds nest high up in a tall coniferous tree or on a cliff ledge that is sheltered overhead and undercut or nearly vertical below. Generally selects locations inaccessible to humans and will sometimes use the same site in successive years. FOOD: Scavenge for road kills along highways, and eats small mammals, reptiles, frogs, eggs, young and wounded birds, insects, mollusks, cultivated grains, mast, fruits, and other plant material. Eats all types of carrion, from small to large mammals to fishes. REFERENCES: DeGraff et al. 1980, Goodwin 1976, Harlow et al. 1975, Hooper 1977, Johnsgard 1979, Knight and Call 1980, Terrill in Farrand 1983b, White and Cade 1971, Wilmore 1977. 312 Slack-capped Chickadee 'arus atricapillus L 4 Vi' RANGE: Resident from western and central Alaska, Saskatchewan, southern Quebec, and Newfoundland south to northwestern California, northeastern Nevada, central New Mexico, northeastern Oklahoma, central Indiana, and northern New Jersey, and in the Appalachians at higher elevations. Wanders irregularly south in winter. STATUS: Common. HABITAT: Prefers mixed woodlands but also inhabits deciduous and coniferous forests. Will inhabit dense woodlands to thickets, orchards, and urban areas, wherever suitable nesting cavities exist or can be excavated. SPECIAL HABITAT REQUIREMENTS: Comparatively open sites near deep woods, and dead standing trees larger than 4 inches dbh for nesting and feeding. NEST: Usually excavates own nest holes in soft decayed wood of a dead tree or branch stub; will use existing cavities of other birds or bird houses. Prefers to nest in tree species that occur in early serai stages such as aspen, paper birch, yellow birch, willow, basswood, maple, and white ash. Favors trees adjacent to open areas in forest or edge situations for nest sites. Generally roosts in dense foliage rather than cavities. FOOD: Forages from the ground to the tree tops for a variety of insects, conifer seeds, and fruits. REFERENCES: Bailey and Niedrach 1965, Bent 1946, Brewer 1961, Forbush and May 1955, Johnsgard 1979, Martin et al. 1951, Odum 1941a, 1941b, 1942, Thomas et al. 1979. 313 Carolina Chickadee Parus carolinensis RANGE: Resident from southern Kansas, central Illinois, central Ohio, and central New Jersey south to central and southeastern Texas, the Gulf Coast and northern peninsular Florida. Wanders casually to the north and southward. STATUS: Common. HABITAT: Inhabits coniferous and deciduous woodlands; prefers forest and forest edge habitats similar to, but more moist and warm than, those preferred by the black-capped chickadee. Also frequents swamps, thickets, second-growth woodlands, parks, and brushy areas. SPECIAL HABITAT REQUIREMENTS: Standing dead trees for excavating cavities. NEST: Usually excavates nest holes in dead, decayed tree trunks or in dead limbs of living trees. Occasionally nests in old woodpecker holes or natural cavities. (The nest and eggs of this species cannot be distin¬ guished from those of the black-capped chickadee.) Chooses willow, pine, cottonwood, poplar, pear, and cherry for nest trees. FOOD: Has very similar food habits to those of the black-capped chickadee. REFERENCES: Bent 1946, Brewer 1961, 1963, Johnsgard 1979, Pitts 1976. 314 Mexican Chickadee Parus sclateri L 4'A" RANGE: Resident in Mexico, the Chiricahua Mountains of southeastern Arizona, and the Animas Mountains of southwestern New Mexico. STATUS: Locally common. HABITAT: Occurs in almost any habitat with conifers up to 7,000 to 8,000 feet in elevation, even where trees are sparse, but prefers pine-oak woodlands and montane pine and spruce-fir forests, primarily in mesic habitats. Outside the breeding season, may also be found in groves of Arizona cypress at lower elevations. SPECIAL HABITAT REQUIREMENTS: Dead trees for cavity nest. NEST: Excavates its own cavity in a dead stub, tree trunk, or branch. FOOD: Probably eats insects and seeds similar to other chickadees. (No specific information is available on food habits). REFERENCES: Bent 1946, Harrison 1979, Phillips et al. 1964, Terres 1980. 315 Mountain Chickadee Parus gambeli RANGE: Resident from northwestern and central British Columbia, southwestern Alberta, western and south-central Montana, and Colorado south to Baja California, southern Nevada, central and southeastern Arizona, southern New Mexico, and extreme western Texas. STATUS: Common. HABITAT: Inhabits open coniferous forests from 6,000 to 11,000 feet in elevation. In winter, often ranges downslope to the foothills, frequenting oaks, and cottonwoods and willows along streams. SPECIAL HABITAT REQUIREMENTS: Decayed trees or stubs for excavating nests, or old cavity nests. NEST: Usually nests in natural cavities or abandoned woodpecker holes, but will excavate its own holes in rotted wood. Many nest in aspen trees associated with a conifer forest. FOOD: Gleans much of its food from foliage, especially large volumes of adult and larval insects. When densities of lodgepole needle miners are high, consumes these insects for much of its diet. Also eats seeds, spruce buds, and fruits. REFERENCES: Bent 1946, DeWeese et al. 1979, Phillips et al. 1964, Scott et al. 1977, Telford and Herman 1963, Winternitz 1973. 316 Siberian Tit D arus cinctus formerly Gray-headed Chickadee) L 4W RANGE: Resident from northern Alaska east across northern Yukon to northwestern Mackenzie, and south locally to western and central Alaska. STATUS: Uncommon. HABITAT: Inhabits spruce, willow, aspen, and birch stands along rivers at the northern limit of the boreal forest. During winter, wanders widely through river valleys. SPECIAL HABITAT REQUIREMENTS: Natural cavities, abandoned woodpecker holes, or trees with soft, dead wood for nesting. NEST: Builds nest inside an abandoned woodpecker hole or a natural cavity in a tree or stump; may excavate a cavity in trees with soft, dead wood. FOOD: Eats adult, larvae, and eggs of insects in summer, and seeds of conifers and berries in winter. REFERENCES: Bent 1946, Gibson in Farrand 1983b, Terres 1980. 317 Boreal Chickadee Parus hudsonicus RANGE: Resident from western and central Alaska and central Yukon to northern Ontario and Labrador, south to extreme north-central Washington, northwestern Montana, northern Minnesota, northern Michigan, northern New York, Maine, and Nova Scotia. After the breeding season, wanders irregularly to the south. STATUS: Fairly common. HABITAT: Associated with northern coniferous forests, where it inhabits spruce, balsam, and dense pine woodlands, white cedar and hemlock swamps, bogs, and occasionally birch and streamside willows. SPECIAL HABITAT REQUIREMENTS: Available cavities, or decaying trees with soft heartwood and hard exterior layers and bark for exca¬ vating nest cavities. NEST: Excavates nest cavity in trees or stubs, preferably with soft and decayed heartwood but hard outer layers; it may use natural cavities or old woodpecker holes. Uses cavity opening from 1 to 10 feet above the ground, facing upward rather than laterally like the nests of other chick¬ adees. Selects nest sites more for the softness of the heartwood than for the species of tree. FOOD: Consumes adults, larvae, pupae, and eggs of insects found by gleaning and probing tree trunks, bark crevices, and foliage. Also extracts seeds from cones, and eats fruits. REFERENCES: Bent 1946, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, McLaren 1975, Terres 1980. 318 Chestnut-backed Chickadee Parus rufescens RANGE: Resident from south-central and southeastern Alaska, western British Columbia, northern Idaho, western Alberta, and northwestern Montana south through the coast ranges to southern California and through the Cascades and Sierra Nevadas to central California. STATUS: Common; has extended its range in California in the past 20 years. HABITAT: Prefers low-elevation, coastal, mesic coniferous forests of pines, cedar, tamarack, and hemlock. Also inhabits along streams and in adjacent deciduous woodlands. SPECIAL HABITAT REQUIREMENTS: Available tree cavities or rotted snags suitable for nest excavation. NEST: Builds nest in natural cavities, or abandoned woodpecker holes, or excavates cavities in soft, rotted tree stubs. Prefers pine, oak, and Douglas-fir snags. FOOD: Gleans much of its food from tree trunks, but also from rotting logs on the ground. Primarily eats insects but also takes spiders, some fruit pulp, and conifer seeds. REFERENCES: Beal 1907, Bent 1946, Grinnell and Miller 1944, Root 1964, Terres 1980. 319 Bridled Titmouse Parus wollweberi L RANGE: Resident from central and southeastern Arizona and southwest¬ ern New Mexico, south locally to Mexico. STATUS: Common. HABITAT: Generally found in oak woodlands and pine-oak associations from 5,000 to 7,000 feet in elevation. In winter, may move down slope along streams where cottonwoods are present. SPECIAL HABITAT REQUIREMENTS: Natural cavities in living or dead oaks. NEST: Usually builds nest in natural cavities of dead and living oak, but will also use cavities in cottonwood, willow, and mesquite. FOOD: Spends much of its time foraging in crevices in bark, on tree trunks, and on branches, presumably for adults, larvae, and eggs of insects. REFERENCES: Bent 1946, Phillips et al. 1964, Terres 1980. 320 Plain Titmouse Parus inornatus RANGE: Resident from southern Oregon, northeastern Nevada, southern Wyoming, and western Oklahoma, south to Baja California, central and southeastern Arizona, southern New Mexico, and extreme western Texas. STATUS: Common. HABITAT: In California, prefers oak woodlands; in the Great Basin and desert mountain ranges, occurs in pinyon-juniper woodlands. Greatest numbers inhabit evergreen trees in dry woodlands of the Southwest. SPECIAL HABITAT REQUIREMENTS: Nest and roost cavities. NEST: Usually builds nest in natural cavities or old woodpecker holes, primarily in oak trees; will also use nest boxes if available. Is capable of excavating its own cavity in rotted wood. FOOD: Gleans much of its food, which is predominantly insects, from limbs, twigs, and from the ground. Also eats leaf galls, weed seeds, pinyon nuts, acorns, oats, and cherries. REFERENCES: Bent 1946, Dixon 1949, Johnsgard 1979, Root 1964, Terres 1980, Wetmore 1964. 321 Tufted Titmouse Parus bicolor L RANGE: Resident from northeastern Nebraska, central and eastern Iowa, southern Wisconsin, northern Ohio, southern Ontario, central New York, western Massachusetts, and southwestern Connecticut south to western Texas, the Gulf Coast, and southern Florida, and west to central Kansas, eastern Oklahoma and eastern Mexico. STATUS: Common. HABITAT: Associated with eastern coniferous and deciduous forests, where it prefers woodland swamps and river bottoms. Also occurs in orchards, low, rich woodlands, woodlots, city parks, and suburban areas. SPECIAL HABITAT REQUIREMENTS: Natural cavities or woodpecker holes for nesting. NEST: Usually nests in natural tree cavities or old woodpecker holes, generally 10 to 20 feet, but ranging from 3 to 90 feet, above the ground. Occasionally nests in bird boxes. FOOD: Gleans food from branch and leaf surfaces during spring and summer and from branch surfaces and the ground in winter. Feeds primarily on insects, especially caterpillars; also eats snails; spiders; berries; seeds of sumac, yellow-poplar, alder, poison ivy, and bayberry; and some mast. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Gillespie 1930, Johnsgard 1979, Laskey 1957, Martin et al. 1951, Terres 1980. 322 Verdin Auriparus flaviceps L 3 Vi" CT RANGE: Resident from northeastern Baja California, southern California, southern Nevada, northern Arizona, southwestern Utah, central New Mexico, and central Texas south into Mexico. Casually in southwestern California and southwestern Oklahoma. STATUS: Common. HABITAT: Inhabits brushy valleys, oak slopes, and other semiarid habi¬ tats where there are stiff-twigged and thorny bushes, or trees such as mesquite, hackberry, hawthorn, catclaw, screw bean, paloverde, and cholla. NEST: Builds a nest that is an oval or ball-shaped mass, up to 8 inches in diameter, of thorny twigs anchored to a limb of almost any tree or shrub species found within its range. Also builds roosting or winter nests. FOOD: Eats mostly insects, searching among terminal twigs, buds, and under leaves for insects, their eggs and larvae. Also eats spiders, pulp from seed pods of paloverde, mesquite, and ironwood; and fruits of wolf berry and date palm. REFERENCES: Johnsgard 1979, Taylor 1971, Terres 1980, Whitaker 1943. 323 Bushtit Psaltriparus minimus RANGE: Resident from extreme southwestern British Columbia, western Washington, western and southern Oregon, southwestern Idaho, northern Nevada, north-central Utah, southwestern Wyoming, north-central Colo¬ rado, western Oklahoma, and central Texas south to Baja California, central and southeastern Arizona, and Mexico. STATUS: Locally common. HABITAT: Found most frequently in pinyon-juniper habitats, but also occurs in tall sagebrush, mountain-mahogany, chaparral, brushy or tree- lined river banks, and in hillside aspen groves. NEST: Builds a gourd-shaped nest of twigs, mosses, roots, lichens, oak leaves, and flowers, that is hung from a branch in a clump of leaves. FOOD: Gleans insects and spiders from foliage of trees and shrubs; also eats some fruit. REFERENCES: Addicott 1938, Johnsgard 1979, Terres 1980. 324 ^ed-breasted Nuthatch Yitta canadensis RANGE: Breeds from south-coastal and southeastern Alaska, southern Yukon, central Manitoba, and Newfoundland south to southern California, central and southeastern Arizona, central Colorado, Wyoming, southwest¬ ern North Dakota, southern Manitoba, southern Michigan, and north- central Ohio; in the Appalachian Mountains to eastern Tennessee and western North Carolina; and south to southeastern Pennsylvania, southern New Jersey, and southern New York. Winters throughout most of the breeding range except at the higher latitudes and elevations, irregularly south to Baja California, southern Arizona, southern Texas, and central Florida. STATUS: Common. HABITAT: Prefers coniferous forests, but sometimes occurs in mixed and deciduous woodlands. SPECIAL HABITAT REQUIREMENTS: Cavities in trees with a minimum dbh of 12 inches for nest sites, or soft dead wood for cavity excavation. NEST: Generally uses natural cavities or woodpecker holes for nesting, but can excavate its own cavity in rotted stubs or dead branches. Typ¬ ically locates nest 15 feet above the ground, but sometimes from 5 to 40 feet. Smears pitch below or around the entrance hole, even when the nest is in a deciduous tree or nest box. FOOD: Pries open conifer cone scales and removes seeds for much of its food. Also feeds on spiders and some insects. REFERENCES: Bent 1948, deKiriline 1952, Forbush and May 1955, Terres 1980, Thomas et al. 1979. 325 White-breasted Nuthatch Sitta carolinensis L5" RANGE: Resident from northwestern Washington, southern British Columbia, central Montana, southern Manitoba, northern Minnesota, northern Michigan, New Brunswick, and Nova Scotia south to Baja California, southern Nevada, central and southeastern Arizona, the highlands of Mexico, western and east-central Texas, and northern Florida. Absent from most of the Great Plains. STATUS: Common. HABITAT: Occurs primarily in deciduous and mixed forests, and locally in coniferous forests. Prefers open woodlands, pinyon-juniper, forest edges, parks, and partly open situations with scattered trees. SPECIAL HABITAT REQUIREMENTS: Natural tree cavities for nesting, preferably in trees with a minimum of 12 inches dbh. NEST: Prefers natural cavities in living trees at almost any height for nesting, but will use cavities in dead or dying trees or old woodpecker holes. Rarely, if ever, excavates its own cavity. Chooses apple, elm, maple, aspen, and ponderosa pine for nest trees. FOOD: Gleans insects from the bark of tree trunks and limbs, but also searches for seeds on the ground. In fall and winter, primarily eats mast, sunflower seeds, and corn. During spring and summer, eats a myriad of arthropods. REFERENCES: Bailey and Niedrach 1965, Bent 1948, DeGraff et al. 1980, Forbush and May 1955, Kilham 1968, Scott and Patton 1975, Thomas et al. 1979. 326 Pygmy Nuthatch Sitta pygmaea RANGE: Resident from southern interior British Columbia, northern Idaho, western Montana, central Wyoming, and southwestern South Dakota south to Baja California, Mexico, southern Nevada, central and southeastern Arizona, central New Mexico, western Texas, and western Oklahoma. STATUS: Common. HABITAT: Generally associated with pine forests; prefers open, parklike forests, especially among ponderosa pines in the lower coniferous forest zone. Occurs less frequently in pinyon-juniper and pines of the Pacific Coast. SPECIAL HABITAT REQUIREMENTS: Pine forests with dead trees for cavity nest sites. NEST: Usually excavates nest cavity near the top of a dead pine where the wood is well rotted, or in the underside of a dead branch about 5 to 60 feet above the ground, often at least 25 feet up. Occasionally nests in aspen snags. FOOD: Searches for food in the tops of pine trees, consuming many insects and conifer seeds. Consumes a diet of about 80 percent insects and spiders. REFERENCES: Bent 1948, Grinnell and Miller 1944, Johnsgard 1979, Norris 1958, Phillips et al. 1964, Scott et al. 1977. 327 Brown-headed Nuthatch Sitta pusilla RANGE: Resident from southeastern Oklahoma, central Arkansas, the northern portions of the Gulf States, northern Georgia, eastern Tennessee, western North Carolina, south-central and eastern Virginia, southern Maryland, and southern Delaware south to eastern Texas, the Gulf Coast, and southern Florida. STATUS: Locally common to rare in parts of its range. HABITAT: Prefers open pine or pine-hardwood woodlands, particularly burned-over areas or clearings where there are dead trees or old stumps. Virtually never found outside the coastal plain of the Southeast, or pine habitats. SPECIAL HABITAT REQUIREMENTS: Dead trees or stumps for nest excavation. NEST: Excavates nest cavities in dead trees and stumps (often fire- blackened) or in posts or poles. Cavities are seldom over 13 feet high and usually less than 5 feet above the ground. Occasionally uses old woodpecker holes or natural cavities. FOOD: Eats mainly insects in summer, foraging for food on tree branches and trunks. Mostly eats pine seeds in winter. REFERENCES: Bent 1948, Norris 1958, Pearson 1936, Terres 1980. 328 Brown Creeper Certhia americana L4%" RANGE: Breeds from southwestern, central, and southeastern Alaska, central Alberta, central Manitoba, and Newfoundland south to southern California, across to extreme western Texas, southeastern Nebraska, southeastern Missouri, southern Ontario, eastern Ohio, and West Virginia; in the Applachians to eastern Tennessee and western North Carolina; and to the lowlands of Virginia, Maryland and Delaware. Breeds also through Mexico into Central America. Winters generally through the breeding range, withdrawing from the higher latitudes and elevations and south throughout the eastern United States and southern Texas, the Gulf Coast, and central Florida. STATUS: Inconspicuous, but locally common. HABITAT: Inhabits dense coniferous, deciduous, and mixed woodlands, montane forests, and wooded swamps with standing dead trees with loose bark. During migration and in winter, occurs in open woodlands, scrub forests, parks, and suburban trees. SPECIAL HABITAT REQUIREMENTS: Dead trees with loose bark, preferably with a minimum dbh of 10 inches. NEST: Constructs nest between loose bark and the trunk of a live, dead, or dying tree, generally 5 to 15 feet above the ground. Occasionally nests in natural cavities or old woodpecker holes. FOOD: Explores tree trunks and branches for insects and larvae. Also eats a small amount of mast. REFERENCES: Bent 1948, Davis 1978, DeGraff et al. 1980, Johnsgard 1979, Terres 1980, Thomas et al. 1979. 329 STATUS: Locally common; there are no apparent ecological factors to hinder the spread of this species along coastal, tropical, southeastern Florida. HABITAT: Has adapted well to exotic trees and shrubs in large suburban yards around Kendall, Florida. Generally stays under cover of vegetation, but occasionally perches in the open. Flocks together in the nonbreeding season and begins roosting assemblages in July and August. SPECIAL HABITAT REQUIREMENTS: Ornamental trees and shrubs old enough to bear berries and fruits for food. NEST: Currently nests only within suburbs, using virtually any shrub, hedge, or small tree. Builds nest in the crotch of a low shrub or small tree 2 to 8 feet above the ground. FOOD: Eats fruits, berries, flowers, nectar, and some insects. The fruit of the Brazil peppertree is a very important food item during winter months. REFERENCES: Carleton and Owre 1975, Fisk 1966, Sykes in Farrand 1983b. 330 Cactus Wren Oampylorhynchus brunneicapillus RANGE: Resident from southern California, southern Nevada, south¬ western Utah, and central Arizona to central and southern Texas south to Mexico. STATUS: Common. HABITAT: Inhabits southwestern deserts, primarily where there are abundant cacti and thorny trees, especially large cholla, mesquite, and paloverde. Also frequents riparian brush and trees in towns of arid regions. SPECIAL HABITAT REQUIREMENTS: Thorny shrubs or trees for nesting sites. NEST: Usually constructs a conspicuous nest in cholla cactus, catclaw, or other thorny shrubs or trees, from 3 to 14 feet, but typically 4 to 9 feet, above the ground. Occasionally may nest in orange trees, old woodpecker holes, or in a hollow cornice of a building. Covered roosting nests are built for use throughout the year. FOOD: Feeds mostly on the ground, but also gleans insects from branches of trees and shrubs. Consumes insects, some spiders, occasionally lizards and tree frogs, cactus fruit, berries, and some seeds. REFERENCES: Anderson and Anderson 1959, 1973, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983b. 331 Rock Wren Salpinctes obsoletus RANGE: Breeds from south-central British Columbia and southern Alberta to the western Dakotas south (east of the coast ranges in Washington, Oregon and northern California) to Baja California and Central America, and east to western Nebraska and central and southern Texas. Winters from northern California, southern Nevada, and southern Utah to north- central Texas south through the southern portions of the breeding range, wandering to lower elevations. STATUS: Fairly common. HABITAT: Primarily inhabits arid and semiarid environments, preferring open, rocky areas such as rock outcrops, canyons, fractured cliff faces, talus slopes, and dry earth banks. May be found up to 10,000 feet in the Rocky Mountains and shows no preference for areas with water through¬ out its range. SPECIAL HABITAT REQUIREMENTS: Rough, rocky surfaces with crevices for foraging and cover. NEST: Typically locates nest on slopes of loose rocks and boulders, in crevices of canyon walls, or sometimes in rodent cavities in banks or in tree holes. Builds a well hidden nest, often with a small runway of stones, sometimes 8 to 10 inches long, leading to the nest. FOOD: Forages almost exclusively in open or relatively unvegetated sites. Gleans insects and spiders from boulders, rocks, barren ground, and crevices. REFERENCES: Bent 1948, Johnsgard 1979, Verner and Boss 1980, Webster in Farrand 1983b. 332 Canyon Wren Catherpes mexicanus RANGE: Resident from southern interior British Columbia and eastern Washington to Wyoming, southeastern Montana, and southwestern South Dakota south to Baja California and Mexico. STATUS: Fairly common. HABITAT: Found in two primary habitats; areas with water, such as boulder-strewn streams, rocky canyons, and river gorges, and major rock formations, such as tall cliffs, large caves, mesas, and buttes. Prefers cool, shaded canyons with rock outcrops. SPECIAL HABITAT REQUIREMENTS: Small cliffs, talus, or rock outcrops for nesting and foraging. NEST: Favors ledges in caverns or rocky crevices for nest sites but sometimes attaches nest to a rock face in a cave or wide crevice. May also nest in buildings. FOOD: Forages mainly in secluded or covered habitats, gleaning insects and spiders from rock surfaces or the ground. REFERENCES: Bent 1948, Johnsgard 1979, Verner and Boss 1980, Webster in Farrand 1983b. 333 Carolina Wren Thryothorus ludovicianus RANGE: Resident from eastern Nebraska, Iowa, and southeastern Minnesota across to southern Ontario, extreme southwestern Quebec, and southern New England, south to Mexico, the Gulf Coast, and Florida. STATUS: Common; populations are beginning to recover in the northern portion of its range where they had previously declined. HABITAT: Found in a variety of habitats from lowland streambank tangles to upland brushy slopes and woodland edges, especially in moist areas with thickets and undergrowth such as honeysuckle, greenbrier, and brush piles. Also frequents cutover forests, cultivated areas with brush heaps or old buildings, and suburban parks and gardens. In winter, moves to low, flat ground near tidewater creeks in the Northeast, and to narrow valleys and deep ravines in other areas. SPECIAL HABITAT REQUIREMENTS: Low, brushy vegetation. NEST: Prefers nest sites that are fairly well enclosed. Typically nests in natural tree cavities, woodpecker holes, overturned root cavities, bird- houses, under rocks, and in building crevices. Locates nest usually less than 10 feet above the ground, sometimes in low shrubs or in grasses. FOOD: Consumes a diet that is about 94 percent animal food, nearly all insects gleaned from trees, shrubs, and the ground. Also eats some snails, lizards, tree frogs, berries, and seeds. Will come to bird feeders for food. REFERENCES: Armistead in Farrand 1983b, Bent 1948, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Nice and Thomas 1948, Scott et al. 1977, Tate and Tate 1982. 334 Bewick’s Wren 'hryomanes bewickii RANGE: Breeds from southwestern British Columbia, southern Wyoming, eastern Nebraska, southeastern Minnesota, southern Ontario, and south¬ eastern New York south to Mexico, central Texas, the northern portions of the Gulf States, central Georgia and central South Carolina. Resident in the West; winters from the northern limits of the breeding range, southern Kansas, the lower Ohio Valley, and North Carolina to Mexico, the Gulf Coast, and central Florida. STATUS: Scarce and local throughout the eastern portion of the breeding range, locally common in the West. HABITAT: Generally associated with dense, brushy habitats such as thickets of mesquite, oaks, and cacti; chaparral; mixtures of pine, junipers, and oaks; dense growths of alder, cottonwood, and willow. In the Southwest, occurs in mountain canyons up to 6,000 feet in elevation. SPECIAL HABITAT REQUIREMENTS: A brushy understory and cavities for nesting. NEST: Nests near the ground in secluded natural tree cavities, old wood¬ pecker holes, rock crevices, deserted buildings, birdhouses, or in almost any cavity where a nest could be built. (Nest and nesting sites are like those of the house wren, and the two species usually compete when in the same area.) FOOD: Gleans small insects and spiders (about 97 percent of diet) from low trunks and branches of trees and brush, usually under dense cover. REFERENCES: Beal 1907, Bent 1948, Johnsgard 1979, Miller 1941, Tate and Tate 1982, Verner and Boss 1980. 335 House Wren Troglodytes aedon RANGE: Breeds from southern and east-central British Columbia and northern Alberta east to southwestern Quebec and New Brunswick, and south to Baja California, Mexico, western and northern Texas, central Arkansas, southern Tennessee, and North Carolina. Winters from south¬ ern California to northern Texas, the northern portion of the Gulf States, and coastal Maryland south to Mexico, the Gulf Coast, and Florida. STATUS: Common. HABITAT: Originally associated with deciduous forests and open woods, it has adapted to woody vegetation in cities, towns, and around farms. Frequents edges of woodlands, open forests, clearings, swampy wood¬ lands, orchards, farmlands, and suburban gardens. Ranges from the plains up to near timberline in the West but avoids high elevations in the East. SPECIAL HABITAT REQUIREMENTS: Woody vegetation and cavities for nesting. NEST: Uses almost any type of cavity as a nest site, including natural cavities in trees, fenceposts, or stumps, woodpecker holes, and bird- houses or other artificial cavities with openings preferably about 1 inch in diameter. Typically chooses a nest site less than 10 feet above the ground. FOOD: Gleans and hawks insects, which form 98 percent of its diet. REFERENCES: Bent 1948, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Kendeigh 1941. 336 \N inter Wren troglodytes troglodytes L 3V4" RANGE: Breeds from coastal southern and southeastern Alaska and northern British Columbia to central Quebec and southern Labrador, south to central California, central Idaho, southeastern Manitoba, south¬ ern Wisconsin, and southeastern New York, and in the Appalachians to northeastern Georgia. Winters from southern Alaska and British Columbia east to northeast Colorado, central Iowa, southern Michigan, and Massachusetts south to southern California, southern Texas, the Gulf Coast, and central Florida. STATUS: Generally uncommon. HABITAT: Primarily inhabits dense undergrowth of coniferous forests, generally near water. Favors spruce and fir forests, but sometimes inhabits dense mixed and hardwood forests. Frequents thickets near woodland streams, boreal swamps and bogs, banks of marshy ditches, and slash piles. In winter, prefers coniferous and deciduous woodlands with a dense understory, especially in moist areas. SPECIAL HABITAT REQUIREMENTS: Moist coniferous woodlands with low woody vegetation, or low-lying cold bogs or swamps. NEST: Usually nests under an upturned root of a tree or under a stump, in a hollow log, brush heap or rocky crevice, or rarely in an old woodpecker hole. Does not typically build its nest in an enclosed cavity as other wrens do. FOOD: Insects and spiders gleaned from the ground form almost the entire diet. REFERENCES: Bent 1948, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979. 337 Sedge Wren Cistothorus platensis (formerly Short-billed Marsh Wren) L 3 3 A RANGE: Breeds from extreme east-central Alberta and central Saskatchewan east to northern Michigan and southern New Brunswick, south to east-central Arkansas, central Kentucky, and southeastern Virginia, and west to central North Dakota and eastern Kansas. Winters from western Tennessee and Maryland to northeastern Mexico, Texas, the Gulf Coast, and Florida. STATUS: Scarce and local; populations are declining in the Northeast and Midwest. HABITAT: Inhabits wet meadows and the damp upper margins of marshes and sphagnum bogs. In the Northeast, commonly inhabits sedge meadows, shallow sedge marshes with scattered shrubs and little or no standing water, and coastal brackish marshes of marsh hay cord- grass with scattered low shrubs and herbs. In the Midwest, prefers wet meadows dominated by sedges, cottongrass, mannagrass, and reed grass, but also frequents emergent vegetation associated with marshes, and retired croplands and fields. SPECIAL HABITAT REQUIREMENTS: Wet meadows or drier edges of marshes for nesting. NEST: May be loosely colonial in good habitat, otherwise nests singly. Builds nest over land or water in dense vegetation such as canarygrass. sedges, or bulrushes, but shuns cattails; usually places nest 1 to 3 feet above the substrate. Males build many unlined dummy nests, but few are used by females. FOOD: Gleans insects and spiders from the ground and surrounding marsh vegetation. REFERENCES: Crawford 1977, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Mousley 1934b, Tate and Tate 1982, Vickery in Farrand 1983b, Walkinshaw 1935. 338 Vlarsh Wren 'istothorus palustris formerly Long-billed Marsh RANGE: Breeds from southwestern and east-central British Columbia and northern Alberta east to northern Michigan and eastern New Brunswick, south to Baja California, southwestern Arizona, extreme western and southern Texas, the Gulf Coast, and east-central Florida. 3enerally very local in interior North America. Winters in coastal areas throughout the breeding range, and in the interior from the southern United States to Mexico. STATUS: Locally common. HABITAT: Prefers large fresh or brackish marshes with an abundance of tall emergent vegetation such as cattails, loosestrife, sedges, or rushes. Also frequents prairie sloughs, pond and sluggish river shores, marsh- fringed lakes, and the banks of tidal rivers bordered with tall emergent vegetation. Prefers large marshes grown with narrow-leaved cattails to those with broad-leaved cattails. SPECIAL HABITAT REQUIREMENTS: Marshy habitats with tall emergent vegetation. NEST: Builds domed elliptical nest preferably in cattail stands of moderate density, 3 to 5 feet above the marsh substrate, which is generally shallow water. Usually attaches to cattails or other tall emergent vegetation, but may place it in small bushes or trees. Constructs many dummy nests and uses some for roosting. FOOD: Gleans insects and spiders from surrounding marsh vegetation and the surface of the water; also hawks for insects and eats a few snails. REFERENCES: Bent 1948, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Low and Mansell 1983, Verner 1965, Verner and Engelsen 1970, Vickery in Farrand 1983b, Welter 1935. 339 American Dipper Cinclus mexicanus (formerly Dipper) RANGE: Resident from western and northeastern Alaska and north- central Yukon to southwestern Alberta, north-central Montana, and southwestern South Dakota south to southern California, north-central and southeastern Arizona, southern New Mexico, Mexico, and Central Mexico. STATUS: Fairly common. HABITAT: Found along rapidly flowing mountain streams in the West, with numerous falls and cascades, and beds filled with large rocks and boulders. Primarily in the vicinity of coniferous forests from 2,000 feet to timberline; less frequently found in the vicinity of mountain ponds and lakes. SPECIAL HABITAT REQUIREMENTS: Clear, permanent streams or rivers. NEST: Usually locates nest over water, either under overhanging rock ledges or under bridges, from just above water level to 15 feet high. Places nest on a rock in midstream, behind a waterfall, or more commonly, in a niche in a rock wall, or sometimes among the roots of a fallen tree. FOOD: Eats mainly insect larvae and adults, snails, and fish fry. Searches for food while completely submerged under fast-flowing water. Catches some insects in the air by hawking. REFERENCES: Bakus 1959, Hann 1950, Johnsgard 1979, Thut 1970. 340 Golden-crowned Kinglet Regulus satrapa RANGE: Breeds from southern Alaska to northern Alberta, southern Quebec, and Newfoundland south in the coastal and interior mountains to southern and eastern California, southern Utah, south-central New Mexico, Mexico, Guatemala, and east of the Rockies to southern Manitoba, north-central Michigan, New York, eastern Tennessee, western North Carolina, northern New Jersey, and southern Maine. Winters from south-coastal Alaska and southern Canada south to northern Baja California, through the breeding range to Guatemala, the Gulf Coast, and central Florida. STATUS: Common in parts of its range; has declined in western regions. HABITAT: Breeds primarily in dense coniferous forests, especially where spruce is present. Winters in coniferous forests and occasionally in deciduous woodland scrub and brush. NEST: Builds a globular nest with entrance at the top, woven into the twigs of a horizontal limb of a conifer. FOOD: Forages over leaves, branches, and trunks, feeding almost entirely on insects and their eggs (bark beetles, scale insects) and especially plant lice. In summer, feeds mainly on flying insects. REFERENCES: Bent 1949, DeGraff et al. 1980, Forbush and May 1955, Tate and Tate 1982, Terres 1980. 341 Ruby-crowned Kinglet Regulus calendula RANGE: Breeds from northwestern and north-central Alaska, northern Saskatchewan, northern Ontario, and Newfoundland south to southern Alaska, in the mountains to southern California, southern Arizona, south- central New Mexico, and east-central Colorado, and east of the Rockies to central Alberta, southern Manitoba, northeastern Minnesota, northern Michigan, northern New York, northern Maine, and Nova Scotia. Winters from southern British Columbia, Idaho, northern Arizona, Nebraska, southern Ontario, and New Jersey south to Baja California, southern Texas, southern Florida, and through Mexico to Guatemala. STATUS: Locally common. HABITAT: Generally inhabits coniferous forests or coniferous-deciduous woodlands during the summer breeding season. In migration and during winter also found in deciduous forests, open woodlands, brush, and scrub. NEST: Usually attaches nest to pendent twigs beneath a horizontal spruce branch (occasionally fir or pine), generally from 15 to 60 feet above the ground. FOOD: Gleans or hawks its food, which consists mainly of insects and spiders. Also eats some elderberries and weed seeds. REFERENCES: Beal and McAtee 1912, DeGraff et al. 1980, Forbush and May 1955, Terres 1980. 342 Blue-gray Gnatcatcher Polioptila caerulea RANGE: Breeds from southern Oregon, northern California, southern Idaho, central Utah, Colorado, Nebraska, western Iowa, southeastern Minnesota, Michigan, southwestern Quebec, central New York, central Vermont and southern Maine south to Baja California, to southeastern Texas, the Gulf Coast, and southern Florida, throughout Mexico to Central America. Winters from southern California, southern Nevada, western and central Arizona, central Texas, the southern portions of the Gulf States, and on the Atlantic Coast from Virginia south through Mexico to Central America. STATUS: Common in parts of its range, but numbers fluctuate. HABITAT: In the Southeast, it inhabits forested river bottoms and upland pine woods with an understory of oaks. In other areas, it may inhabit open scrub and woodlands, or tall trees of closed canopy along river flood plains. Throughout the West it breeds in oaks, pinyon-juniper, and less frequently in chaparral. SPECIAL HABITAT REQUIREMENTS: An abundant supply of arthropods. NEST: Places nest saddled on a horizontal limb 4 to 70 feet high (average 25 feet), in a conifer or deciduous tree, but usually in deciduous oaks. FOOD: Gleans food from the tips of branches, leaf surfaces, and bark; also hawkes flying insects from perches. Mostly eats arthropods, principally insects, and some spiders. REFERENCES: Forbush and May 1955, Root 1967, 1969, Terres 1980. 343 Black-tailed Gnatcatcher Polioptila melanura RANGE: Resident from southwestern California and northwestern Baja California south locally to southern Baja California and from northwestern Baja California, southeastern California, southern Nevada, western and central Arizona, southern New Mexico, and western and southern Texas south into Mexico. STATUS: Common, but may be declining. Listed on the blue list for declining species in 1982. HABITAT: Found in the lower elevations of the Southwest, where it prefers desert brush and scrub, especially mesquite and creosote bush, and in coastal sagebrush and thorn forests. NEST: Places its small, deep cup, invariably low (2-4 feet above ground) in buckthorn, laurel, sumac, sagebrush, cactus, or other desert plant. FOOD: Gleans insects and some spiders from branches and twigs of shrubs. Also eats small amounts of seeds. REFERENCES: Harrison 1979, Tate and Tate 1982, Terres 1980, Terrill in Farrand 1983c. 344 Eastern Bluebird ialia sialis L 5W RANGE: Breeds from southern Saskatchewan, southern Quebec, and western Nova Scotia south to southern Texas and southern Florida, and west to the Dakotas, western Kansas, Texas, and southeastern New Mexico; also in southeastern Arizona and through the highlands of Mexico to Central America. Winters from the middle portions of the eastern United States south throughout the breeding range. STATUS: Population low but stable, many dependent on nest boxes. HABITAT: Inhabits fields, forest edges, open woodlands, and open country with scattered trees, and in coniferous, deciduous, and riparian woodlands. SPECIAL HABITAT REQUIREMENTS: Low cavities for nesting and perches for foraging. NEST: Nests in old woodpecker holes, hollows of decayed trees, crevices of rocks, and hollows in wooden fence posts when available. Many now nest in artificial nest boxes placed in open areas or at the edge of a forest. FOOD: Catches insects and spiders by flying from a perch to the ground or hawking. Eats fruits and a few seeds during winter. REFERENCES: Beal 1915a, Forbush and May 1955, Hartshorne 1962, Pearson 1936, Rustad 1972, Tate and Tate 1982, Thomas 1946. 345 Western Bluebird Sialia mexicana RANGE: Resident from southern British Columbia, western and south- central Montana, and north-central Colorado south through the mountains to Baja California, western and southern Nevada, southern Utah, western and southeastern Arizona, central New Mexico, western Texas, and in the highlands of Mexico. Wanders in winter to lowland areas throughout the breeding range, and to islands off California and Baja California. STATUS: Overall population is low but stable. HABITAT: Mostly inhabits open ponderosa pine forests of the transition zone but is also found in other open coniferous, deciduous, and mixed forests, partly open country with scattered trees, savannah, and riparian woodlands. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting and perches for feeding. NEST: Usually nests in old woodpecker holes, but also uses natural cavities and nest boxes. Prefers to locate nests in rather open forests or at forest edges. FOOD: Sometimes hawks insects from high perches; otherwise flies to the ground from low perches to catch prey. Mostly eats insects and spiders but also some fruits such as elderberries and mistletoe berries. REFERENCES: Beal 1915a, Herlugson 1982, Tate and Tate 1982. 346 Mountain Bluebird Sialia currucoides L 6" RANGE: Breeds from east-central Alaska, southern Yukon, and western Manitoba south in the mountains to southern California, central and southeastern Nevada, northern and east-central Arizona, and southern New Mexico, and east to northeastern North Dakota, western South Dakota, and central Oklahoma. Winters from southern British Columbia and western Montana south to Baja California, Mexico, and southern Texas, and east to eastern Kansas, western Oklahoma, and central Texas. STATUS: Population is low but stable. HABITAT: Nests in nearly all forest types of the Rocky Mountain region, usually from 7,000 to 11,000 feet in open forests or near forest edges. During migration and in winter, also frequents grasslands, open brushy country, and agricultural lands. SPECIAL HABITAT REQUIREMENTS: Cavity nests and feeding perches. NEST: Usually nests in old woodpecker holes or natural cavities in dead trees in open areas or near forest edges. Will also use nest boxes. FOOD: Hawks from high perches or flies to the ground to catch its prey. Probably more insectivorous than the other bluebirds; nearly 92 percent of the diet is animal material; the small amount of vegetable food includes fruits, hackberry seeds, and cedar berries. REFERENCES: Beal 1915a, Burleigh 1972, Herlugson 1982, Scott et al. 1977, Tate and Tate 1982. 347 Townsend’s Solitaire Myadestes townsendi RANGE: Breeds from east-central and southeastern Alaska, to west- central Mackenzie south in the mountains to southern California, northern and east-central Arizona, central New Mexico, and northern Mexico, and east to southwestern Alberta, western and southern Montana, southwestern South Dakota and northwestern Nebraska. Winters from southern British Columbia, southern Alberta, Montana, and South Dakota south to Baja California, the southern limit of the breeding range in Mexico, and east to western Missouri, western Oklahoma, and central Texas. STATUS: Common within parts of its range. HABITAT: During summer, it is found in montane and subalpine coniferous forests and in thickets and brushy areas adjacent to rocky cliffs up to 12,000 feet in elevation. Winters in open woodland, pinyon- juniper associations, chaparral, desert, and riparian woodlands. SPECIAL HABITAT REQUIREMENTS: Juniper berries for winter food. NEST: Builds nest on the ground, partly concealed at the base of a pine or fir, under overhanging banks, or among the roots of a fallen tree. FOOD: Gleans food from the ground, foliage, and fruiting stems, and hawks for flying insects. Eats insects and spiders during the summer, and mostly juniper berries, along with a few other fruits and seeds, during winter. REFERENCES: Beal 1915b, Poddar and Lederer 1982, Terres 1980, Verner and Boss 1980. 348 rusty race t/eery Zatharus fuscescens L 6" RANGE: Breeds from south-central and southeastern British Columbia to New Brunswick and southwestern Newfoundland, south to central Oregon, southern Idaho, northeastern South Dakota, northern Illinois, and northern Ohio, in the mountains through West Virginia, western and central Maryland, eastern Kentucky, western and central Virginia, eastern Tennessee, and western North Carolina to northwestern Georgia, and in the Atlantic region to eastern Pennsylvania, central New Jersey, and the District of Columbia. Also in east-central Arizona. Winters in South America. STATUS: Common. HABITAT: Inhabits low, moist, deciduous woods, bottomland forests, wooded swamps, and damp ravines; prefers sapling stands of deciduous second-growth or open woods with fairly dense undergrowth of ferns, shrubs, and trees. SPECIAL HABITAT REQUIREMENTS: Moist woodlands with understory of low trees or shrubs. NEST: Builds a bulky nest on or near the ground at the base of a shrub, on a mossy stump, in a clump of weeds, or occasionally in a low shrub or tree. FOOD: Often forages on the forest floor, turning leaves with bill in search of food; occasionally searches for food in trees. Consumes a diet that is about 57 percent animal and 43 percent vegetable. REFERENCES: Beal 1915b, DeGraff et al. 1980, Forbush and May 1955, Terres 1980. 349 Gray-cheeked Thrush RANGE: Breeds from northern Alaska, southern Keewatin, and New¬ foundland south to southern Alaska, northwestern British Columbia, northeastern Saskatchewan, eastern New York, Massachusetts, central Vermont, northern New Hampshire, central Maine, and northern Nova Scotia. Winters in South America. STATUS: Common in portions of its range. HABITAT: During summer, inhabits coniferous forests (primarily spruce) and tall shrubby areas in taiga. During migration and in winter, also found in deciduous forests and open woodlands. Generally inhabits mountain tops in the Northeast. SPECIAL HABITAT REQUIREMENTS: Coniferous forests. NEST: Usually builds its nest in willows, alders, or spruces, from ground level to 20 feet, but usually about 6 feet above ground on divergent branches close to the trunk. FOOD: Generally gleans food from the ground; eats insects, spiders, earthworms, crayfish, and berries. REFERENCES: Beal 1915b, DeGraff et al. 1980, Terres 1980. 350 Bwainson’s Thrush 'atharus ustulatus RANGE: Breeds from western and central Alaska, northern Saskatchewan, central Quebec, and Newfoundland south to southern Alaska, southern and east-central California, central Utah, north-central New Mexico, extreme northern Nebraska, eastern Montana, southern Manitoba, northern Minnesota, southern Ontario, northern Pennsylvania, and southern Maine. Also in eastern West Virginia, western Virginia, and western Maryland. Winters from Mexico south. STATUS: Rare to locally common. HABITAT: In summer, inhabits dense coniferous forests (especially spruce) and dense tall shrubbery, or recent clearcuts in low damp areas or near water. In parts of range, prefers aspen-poplar forests and willow or alder thickets, and occasionally breeds in coniferous-deciduous forests. In winter, frequents deciduous forests. SPECIAL HABITAT REQUIREMENTS: Damp forests or adjacent water. NEST: Builds a bulky cup nest, usually near the trunk, on a horizontal branch of a conifer 2 to 20 feet above the ground. FOOD: Gleans food from the forest floor, foliage, and branch surfaces. Also eats insects, spiders, millipedes, and small fruits and berries. REFERENCES: Beal 1915b, DeGraff et al. 1980, Forbush and May 1955, Terres 1980, Verner and Boss 1980. 351 Hermit Thrush Catharus guttatus RANGE: Breeds from western and central Alaska, northern Saskatchewan, and Newfoundland south to southern Alaska, in the mountains to south¬ ern California, southern Nevada, southern New Mexico, and western Texas, and east of the Rockies to central Alberta, central Wisconsin, southern Ontario, central Pennsylvania, western Virginia, western Maryland, southern New York, and in the Black Hills in southwestern South Dakota. Winters from southern British Columbia and the northern United States south to Baja California, Mexico, southern Texas, and southern Florida. STATUS: Common. HABITAT: In summer, inhabits coniferous, mixed, and deciduous forests with intermediate to high canopy coverage; responds negatively to intensive tree harvests. During migration and in winter, also inhabits chaparral, riparian woodlands, arid pine-oak associations, and desert scrub. SPECIAL HABITAT REQUIREMENTS: Relatively undisturbed, rather dense forests. NEST: Usually builds nest in a depression on the ground, under rock ledges, or under low overhanging limbs. Sometimes locates nests in shrubs or small trees near the ground, especially in the West. FOOD: Gleans most of its food from the ground. Eats insects, spiders, snails, and earthworms, plus considerable amounts of wild fruits in fall and winter. REFERENCES: Beal 1915b, DeGraff et al. 1980, Forbush and May 1955, Szaro and Baida 1982, Terres 1980, Verner and Boss 1980. 352 Wood Thrush Hylocichla mustelina RANGE: Breeds from southeastern North Dakota, northern Michigan, northern Vermont, southwestern Maine, and Nova Scotia south to east- central Texas, the Gulf Coast and northern Florida, and west to eastern South Dakota, central Kansas, and eastern Oklahoma. Winters from southern Texas south to Central America. STATUS: Common. HABITAT: Inhabits cool, mature, lowland deciduous or mixed forests, particularly damp situations and near swamps or water. In New England, also found on wooded slopes; has adapted to gardens and city parks. SPECIAL HABITAT REQUIREMENTS: Deciduous or mixed forests with tall trees. NEST: Builds a compact cup nest on a horizontal limb, in a fork of a sapling or tree, or well hidden in dense shrubbery, generally 6 to 50 feet (average 10 feet) above ground. FOOD: Gleans most of its food from the ground, but occasionally gleans insects from tree foliage. Eats insects, spiders, snails, earthworms, and berries. REFERENCES: Beal 1915b, Brackbill 1943, DeGraff et al. 1980, Forbush and May 1955, Terres 1980. 353 American Robin Turdus migratorius RANGE: Breeds from western and northern Alaska, southern Keewatin, northern Quebec, Labrador, and Newfoundland south to southern California, central and southeastern Arizona, Mexico, southern Texas, and central Florida. Winters from southern Alaska, the northern United States and Newfoundland south to Baja California, southern Texas, and southern Florida, throughout Mexico to Central America. STATUS: Abundant. HABITAT: Found in nearly all habitats from tree limit in sparsely wooded barrens up to 12,000 feet in the mountains of the West, along forest borders, hedges, orchards, gardens, city parks, and in suburban yards. SPECIAL HABITAT REQUIREMENTS: Mud for nest building. NEST: Places nest, constructed of mud and vegetation, on almost any substantial support, usually in a fork or on a horizontal branch of a shrub or tree; rarely on the ground. FOOD: Feeds mostly on the ground, searching for almost any edible substance; but also picks fruits from trees and shrubs. Consumes a diet that is about 42 percent animal and 58 percent vegetable. REFERENCES: Beal 1915b, Forbush and May 1955, Knupp et al. 1977, Terres 1980, Young 1955. 354 Varied Thrush Ixoreus naevius RANGE: Breeds from western and northern Alaska, and northwestern and western Mackenzie south through central and southern Alaska, southwestern Alberta, northwestern Montana, northern Idaho, Washington, and Oregon to extreme northwestern California. Winters from southern Alaska, southern British Columbia, and northern Idaho south through Washington, Oregon, and California to Baja California. STATUS: Uncommon. HABITAT: Favors moist, dense stands of fir near mountain lakes, but also inhabits humid coastal and interior montane coniferous forests. Occasionally found in deciduous forests with a dense understory and in tall shrub stands. NEST: Builds a bulky nest on a horizontal branch or in a crotch of a tree or shrub, usually 10 to 15 feet above ground. FOOD: Generally forages for food on the ground, but also gleans food from foliage and fruiting stems. Consumes a variety of vegetable and animal foods including insects, snails, earthworms, acorns, weed seeds, berries, and fruits. REFERENCES: Beal 1915b, Terres 1980, Verner and Boss 1980. 355 Wrentit Chamaea fasciata RANGE: Resident in coastal areas from northwestern Oregon south to northwestern Baja California, and in interior areas of northern and central California. STATUS: Common resident in suitable habitat. HABITAT: Primarily inhabits chaparral but also inhabits other habitats from coastal brushlands up to ponderosa and black oak vegetation types where dense stands of shrubs are present. SPECIAL HABITAT REQUIREMENTS: Dense shrubs. NEST: Builds a compact cup nest of coarse bark, plant fibers, and grasses, which it conceals 1 to 4 feet above the ground in shrubs such as Baccharis. FOOD: Gleans food from bark surfaces of shrubs; eats primarily insects, spiders, and small fruits. REFERENCES: Erickson 1938, Terres 1980, Verner and Boss 1980. 356 Gray Catbird Dumetella carolinensis RANGE: Breeds from southern British Columbia, southern Ontario, and Nova Scotia, south to central New Mexico and northern Florida, and west to northern and south-central Washington, south-central and eastern Oregon, north-central Utah, and central and northeastern Arizona. Winters from north-central and eastern Texas, the central portions of the Gulf States and in the Atlantic Coastal lowlands from Long Island south along the Gulf-Caribbean slope of Central America. STATUS: Common in breeding season. HABITAT: Prefers dense thickets of shrubby edge habitat, but also inhabits shrubs, briars, vines along woodland borders, dry marsh edges, roadside shrubs, old house sites, abandoned fields, and fence rows. SPECIAL HABITAT REQUIREMENTS: Low, dense, shrubby vegetation. NEST: Hides nest about 3 to 10 feet above the ground in almost any dense woody vegetation such as multiflora rose, barberry, lilac, mock- orange, osage orange, a hedge, or conifer tree. FOOD: Gleans food from the ground; about half of diet is insects. In the fall, eats a variety of fruits. REFERENCES: Forbush and May 1955, DeGraff et al. 1980, Nickell 1965, Terres 1980. 357 Northern Mockingbird Mimus polyglottos (formerly Mockingbird) RANGE: Resident in scattered localities across southernmost Canada from Alberta to Newfoundland: generally resident from northern California, northern Utah, and southern North Dakota, east to southern Maine and south into Mexico. Some northern birds move southward in winter. STATUS: Common. HABITAT: Inhabits a variety of open to partly open landscapes such as farm hedges, isolated shrub patches and trees of the prairie, orchards, woodland edges, pastures with scattered fruit-bearing shrubs, and suburbs and cities; absent from forest interiors. SPECIAL HABITAT REQUIREMENTS: Low, dense, woody vegetation, elevated perches, and a variety of persistent edible fruits. NEST: Builds nest in a fork or on a limb of a small tree, shrub, or vine (preferably evergreen), typically 3 to 10 feet above the ground. Usually produces two broods each year; builds a new nest for each brood. FOOD: Gleans insects from the ground and foliage, or hawks from the air. Eats fruits in fall and winter. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Laskey 1962, Terres 1980, Verner and Boss 1980. 358 Sage Thrasher Oreoscoptes montanus RANGE: Breeds largely between the Sierra Nevada-Cascade Mountain axis and the Rocky Mountains from southern British Columbia, Montana, and Wyoming south to east-central California, southern Nevada, northern Arizona, New Mexico, and northwestern Texas. Winters from northern Arizona’and central Texas south to northern Mexico. STATUS: Common throughout most its range. HABITAT: Mainly limited to semiarid sagebrush plains, but may extend into junipers and mountain-mahogany habitats near sagebrush. SPECIAL HABITAT REQUIREMENTS: Sagebrush. NEST: Sometimes nests on the ground under sagebrush, but usually in branches near the main stem of sagebrush plants, 1 to 3 feet above ground. May also nest in other low-growing shrubs such as greasewood, horsebrush, rabbitbrush, and saltbush. FOOD: Gleans food from the ground, including great numbers of grasshoppers, Mormon crickets, and other insects. Also eats fruits and berries in the fall. REFERENCES: Reynolds 1981, Reynolds and Rich 1978, Rich 1978, Terres 1980, Webster in Farrand 1983c. 359 Brown Thrasher L 10' Toxostoma rufum RANGE: Breeds from southeastern Alberta east to New England and south to Colorado, northern and eastern Texas, the Gulf Coast, and southern Florida. Winters in the south from Texas eastward, ranging north in the Mississippi Valley to Illinois and along the Atlantic Coast to Massachusetts. Rare visitor as far west as the Pacific Coast in migration and winter. STATUS: Common in parts of its range. HABITAT: During summer, inhabits dry thickets in wooded and farming country, brushy pastures, second-growth woods, fencerows, brier patches, roadsides, and sometimes shrubbery of gardens. SPECIAL HABITAT REQUIREMENTS: Low, dense, woody vegetation for nesting and cover. NEST: Builds a bulky nest in any of a variety of shrubs (usually thorny) or low trees, up to 14 feet from the ground, but sometimes on the ground under a small shrub. FOOD: Gleans food from the ground or in shrubs. In spring, eats almost entirely insects, spiders and worms; in summer and fall, eats mostly fruits, mast (mainly acorns), and waste corn. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Kaufman in Farrand 1983c, Terres 1980. 360 _ong-billed Thrasher r oxostoma longirostre L 10' RANGE: Resident in southern Texas and eastern Mexico. Accidental in western Texas. STATUS: Abundant within its range. HABITAT: Inhabits the brush country of southern Texas, especially dense mesquite. Also found among bottomland willow, huisache, condalia, and various other shrubs. SPECIAL HABITAT REQUIREMENTS: Dense shrubs. NEST: Nests in pricklypear, yucca, mesquite, or other shrubby plants, usually 4 to 8 feet above the ground, and near the center of the plant. FOOD: Uncovers food in debris on the ground and in loose soil; mostly eats antiions, ants, beetles, bugs, termites, spiders, and hackberries. REFERENCES: Cottam and Knappen 1939, Kaufman in Farrand 1983c, Oberholser 1974b, Terres 1980. 361 L 8 V t " Bendire’s Thrasher Toxostoma bendirei RANGE: Breeds in southeastern California, southern Nevada, southern Utah, western New Mexico, and Sonora. Winters from central-southern Arizona to Sinaloa. Rare in fall to the southern coast of California. STATUS: Locally common. HABITAT: Inhabits open desert habitats, especially areas with tall vegetation, cholla cactus, creosote bush, and yucca. May also inhabit pinyon-juniper-sage communities, but tends to avoid large areas of continuous, dense brushy cover and grasslands. SPECIAL HABITAT REQUIREMENTS: Desert communities. NEST: Builds a compact nest in almost any of the shrubby vegetation found within its habitat, usually 3 to 10 feet above the ground. FOOD: Spends much of its time on the ground searching and digging for food. Probably has food habits similar to those of other thrashers. REFERENCES: Bent 1948, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983c. 362 Curve-billed Thrasher RANGE: Resident from central and southeastern Arizona, central and northeastern New Mexico, southeastern Colorado, northwestern Okla¬ homa, southwestern Kansas, and western and central Texas south to southern Mexico. STATUS: Common to abundant. HABITAT: Inhabits deserts with extensive thickets of thorny shrubs (paloverde and mesquite) and dense large cactus (saguaro and cholla), and in brushy riparian and residental areas. Prefers areas with thorny shrubs and thickets at the edge of woodlands. SPECIAL HABITAT REQUIREMENTS: Desert shrubs. NEST: Usually builds nest in the fork of cholla cactus but may be in yucca, clumps of mistletoe in mesquite and cottonwood, or in mesquite or similar thickets. Favors nest sites at the edge of woodlands or in cultivated areas. FOOD: Digs in the soil with its curved bill searching for food, and eats beetles, other insects, seeds of cacti, berries, and other fruit. REFERENCES: Oberholser 1974b, Terres 1980, Terrill in Farrand 1983c. 363 California Thrasher Toxostoma redivivum RANGE: Resident in California north to Humboldt and Shasta Counties (west of the Cascade Mountains-Sierra Nevada and the deserts), and in northwestern Baja California. STATUS: Fairly common in suitable habitat. HABITAT: Found most abundantly along mountain bases and up to 5,000 feet elevation. Prefers slopes covered with chaparral or early tree stages of blue oak savannah, digger pine-oak, and riparian deciduous types; also found in foothill towns with mixed brush and short trees, but avoids areas with dense tree canopies. SPECIAL HABITAT REQUIREMENTS: Dense shrubs. NEST: Builds a large, well-concealed loose cup nest near the ground in a large bush or scrubby tree. FOOD: Digs in the soil and turns over leaves with its bill in search of food. Eats a diet consisting of insects, spiders, seeds of berries, hazel¬ nuts, weed seeds, and small fruits. REFERENCES: Beal 1907, Terres 1980, Verner and Boss 1980, Webster in Farrand 1983c. 364 Crissal Thrasher Toxostoma dorsale L 10 Vi" RANGE: Resident from southeastern California, southern Nevada, southwestern Utah, northwestern and central Arizona, central New Mexico, and western Texas south to northeastern Baja California, central Sonora and central Chihuahua. STATUS: Common. HABITAT: Inhabits the more heavily vegetated areas of the southern deserts, such as tall brush along rivers and large washes, and dense mesquite or chaparral from sea level to 6,000 feet elevation. SPECIAL HABITAT REQUIREMENTS: Dense desert shrubs. NEST: Most commonly builds a nest saddled on a branch or in a fork of mesquite trees, but also in willow, sagebrush, greasewood, or other desert shrubs, usually 2 to 8 feet above the ground. FOOD: Eats berries, wild grapes, and insects. (Little is known about its food habits.) REFERENCES: Bent 1948, Phillips et al. 1964, Terres 1980, Webster in Farrand 1983c. 365 L 9 Vi' Le Conte’s Thrasher Toxostoma lecontei RANGE: Resident in the southern San Joaquin Valley of California and across the deserts east of the coast ranges and the Sierra Nevadas to southern Nevada and southwestern Utah, south to Baja California, northern Sonora, and western and south-central Arizona. STATUS: Uncommon and local. HABITAT: Lives in the hottest, driest, most barren deserts of the Southwest, and is most abundant in saltbush and open creosotebush deserts. SPECIAL HABITAT REQUIREMENTS: Deserts with scattered shrubs. NEST: Builds its nest, a bulky mass of thorny twigs and sticks, usually in cholla cactus but also in other desert shrubs or trees such as mesquite, paloverde, saltbush, sage, ocotillo, and ironwood, usually 4 to 8 feet above the ground. FOOD: Has food habits that are probably similar to those of other thrashers. REFERENCES: Bent 1948, Phillips et al. 1964, Terres 1980, Webster in Farrand 1983c. 366 Water Pipit \nthus spinoletta /inter RANGE: Breeds on the Arctic tundra and in the mountains of the West and in Maine. Winters on the Pacific Coast from British Columbia south, and from the southern United States south through Mexico into Central America. STATUS: Common. HABITAT: Breeds in alpine and arctic habitats with rough features such as tussocks, tilted rocks, or eroded spots for nest sites. Requires nesting habitat that is free of snow early in the breeding season; prefers moss- grown slopes with southern exposures. SPECIAL HABITAT REQUIREMENTS: Alpine or arctic tundra with some rough features. NEST: Builds nest on the ground in the shelter of a rock or bank, beside a mossy hummock, or at the base of a tussock. FOOD: Forages while walking, gleaning food from the ground and low vegetation. Eats seeds of grasses and weeds, insects, spiders, mites, small mollusks, and crustaceans. REFERENCES: Forbush and May 1955, Gibb 1956, Sutton and Parmalee 1954a, Terres 1980, Verbeek 1970. summer 367 Sprague’s Pipit Anthus spragueii RANGE: Breeds from north-central Alberta, central Saskatchewan, and west-central and southern Manitoba south to Montana, western South Dakota, North Dakota, and northwestern Minnesota. Winters from the southern tier of the Great Plains States south through Mexico. STATUS: Common. HABITAT: Primarily inhabits extensive areas of grasslands dominated by grasses of medium height. Also inhabits large alkaline meadows and meadow zones of large alkali lakes. SPECIAL HABITAT REQUIREMENTS: Extensive prairie. NEST: Constructs nest of grasses in hollows of the ground, and in clumps of grasses or grasslike plants. FOOD: In summer, mostly eats insects and weed seeds. In winter, flocks frequently in weed-grown fields, eating weed seeds. REFERENCES: Bent 1950, Forbush and May 1955, Johnsgard 1979, Terres 1980. 368 Sohemian Waxwing ombycilla garrulus RANGE: Breeds from central Alaska, Yukon, southwestern Mackenzie, and northern Manitoba south to northern parts of Washington, Idaho, and Montana, central Saskatchewan, and central Manitoba. Winters south to Washington, Colorado, the Great Lakes, and Maine, east to Ontario, southern Quebec, Nova Scotia, and the northern tier of states. Winters irregularly to California, Arizona, northern New Mexico, and northern Texas. STATUS: Common. HABITAT: During summer, inhabits open coniferous forests, muskegs, and less frequently, mixed coniferous-deciduous woodlands. Wanders in large flocks in winter and may be abundant wherever food is available. NEST: Constructs nest of twigs, grasses, and lichens, usually on a horizontal branch of an isolated spruce, tamarack, or pine in open muskeg, 4 to 50 feet above the ground. FOOD: In summer, mostly eats insects, catching many of them by hawking from high perches. In fall and winter, eats mostly fruits, with fruits of mountain-ash and berries of cedar and juniper the most important winter foods. In spring, also eats sap from maple trees. REFERENCES: Bent 1950, Eckert in Farrand 1983c, Forbush and May 1955, Terres 1980. 369 Cedar Waxwing Bombycilla cedrorum RANGE: Breeds from southeastern Alaska, central British Columbia, Alberta, Saskatchewan, northern Manitoba, Ontario, central Quebec, and Newfoundland south to northern California, Nevada, Utah, Colorado, South Dakota, central Missouri, Illinois, Indiana, northern Georgia, western North Carolina, and Virginia. Winters from southern British Columbia, Montana, Saskatchewan, Manitoba, Ontario, New York, and New England south to Central America. STATUS: Locally common to rare. HABITAT: Inhabits a wide variety of open coniferous and deciduous forests, forest edges, farmsteads, parks, and residential areas, but absent from dense forests. During winter, found almost anywhere that trees and shrubs with persistent fruits are present. SPECIAL HABITAT REQUIREMENTS: Fruit- and berry-producing trees and shrubs. NEST: Builds its nest semicolonially in dense coniferous thickets (often cedar) but will use a variety of deciduous trees and shrubs. Places nest on a horizontal limb, often in a crotch next to the main trunk, 4 to 50 feet above the ground. FOOD: Gleans insects from leaf surfaces or hawks from perches. In summer, consumes a diet of about 20 percent insects. In fall and winter, eats nearly all fruits and berries. REFERENCES: DeGraff et al. 1980, Eckert in Farrand 1983c, Forbush and May 1955, Lea 1942, Putnam 1949, Terres 1980. 370 L 6'A" 'hainopepla lainopepla nitens 3ANGE: Breeds from central California, southern Nevada, southern Jtah, southern New Mexico, and western Texas south to Baja California ind into Mexico. Winters from southern California, southern Nevada, central Arizona, southern New Mexico and western and southern Texas south into Mexico. STATUS: Locally common to uncommon or rare. HABITAT: In deserts, primarily inhabits washes, riparian areas, and other habitats that support a brushy growth of mesquite and paloverde. In more northern and coastal areas, inhabits oak chaparral and riparian oak woodlands. SPECIAL HABITAT REQUIREMENTS: Trees or shrubs and berries (especially mistletoe). NEST: Builds nest (almost exclusively by the male) in a forked limb of a mesquite, cottonwood, hackberry, willow, sycamore, oak, or citrus tree, often in a clump of mistletoe, 4 to 50 feet above the ground. FOOD: During the breeding season, captures many insects by hawking from high perches. During other periods, primarily eats fruits and berries (especially mistletoe berries). REFERENCES: Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983c, Verner and Boss 1980. 371 Northern Shrike Lanius excubitor L8" RANGE: Breeds in Alaska, the Yukon, southwestern Mackenzie, and northern parts of Manitoba, Quebec, and Labrador. Winters from southern Alaska and the southern half of Canada south to northern California, central Nevada, northern Arizona, northern New Mexico, Kansas, northern Missouri, central Illinois, Indiana, Ohio, Pennsylvania, and New Jersey. STATUS: Locally common in summer range; uncommon in winter. HABITAT: Inhabits a broad belt of coniferous forest or taiga across Canada and Alaska during summer; strongly prefers forest edges, open willow brush, and brush-bordered swamps and bogs. Prefers semiopen country with short grasses and scattered trees or shrubs during winter. SPECIAL HABITAT REQUIREMENTS: Elevated perches, short vegetation. NEST: Builds a bulky, loose nest of twigs, in spruces, willows, or bushes. 5 to 20 feet above the ground. FOOD: Attacks prey from an elevated perch by hawking or hovering, then diving and pouncing. Mostly eats small birds and mammals; also eats insects (especially grasshoppers and crickets), and some snakes, lizards, and frogs. REFERENCES: Bent 1950, DeGraff et al. 1980, Eckert in Farrand 1983c, Miller 1931, Terres 1980. 372 Loggerhead Shrike .anius ludovicianus RANGE: Breeds from central Alberta, central Saskatchewan, southern Manitoba, Minnesota, central Wisconsin, central Michigan, and south¬ eastern Ontario, south to Mexico and the Gulf Coast. Very rare or absent from most of the Appalachians, Pennsylvania, New York, and New England. Winters in the southern half of the United States and in Mexico. STATUS: On the 1982 Blue List for declining species as a species of concern (mostly in the East); common in parts of the West. HABITAT: Inhabits open country with scattered shrubs or small trees such as shelterbelts, cemeteries, farmsteads, or hedgerows in the plains country and Midwest. In the West, breeds in savannah, pine-oak wood¬ lands, and chapparal types, and prefers very open stands. NEST: Builds a bulky, cup-shaped nest in a variety of shrubs and low, dense trees, rarely less than 3 feet or more than 25 feet above the ground. Hides the nest well below the crown of the bush or tree. FOOD: Sometimes hawks for aerial insects, but takes most of its prey as it dives to the ground from an elevated perch. In the West, eats about 83 percent insects; in the East, 68 percent. Eats mostly grasshoppers and crickets, but also a variety of other insects, small mammals, birds, and reptiles. REFERENCES: Beal and McAtee 1912, Eckert in Farrand 1983c, Johnsgard 1979, Kridelbaugh 1983, Miller 1931, Morrison 1981, Porter et al. 1975, Tate and Tate 1982. 373 European Starling Sturnus vulgaris (formerly Starling) winter summer RANGE: Introduced to North America from Europe. Breeds from south¬ eastern Alaska and the southern half of all Canadian provinces south to Mexico, the Gulf of Mexico, and southern Florida. Still uncommon in some recently occupied parts of the Southwest and the Rocky Mountains, but continues to expand range. Migrates sometimes in the northern part of the range. STATUS: Abundant in most of its range. HABITAT: Occupies a great variety of habitats from suburban to rural and woodlands wherever suitable nesting sites occur. Appears to favor thickly settled agricultural areas and tends to avoid dense forests away from human habitation. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting. NEST: Normally selects an old woodpecker or natural cavity in trees, utility poles, or fence posts but is extremely adaptable in its choice of nest sites. Usually nests earlier than many other cavity nesters and may be a serious competitor for available nest cavities. FOOD: Generally forages on the ground. Is sometimes considered a serious agricultural pest in some areas because it eats practically all grains, digs up sprouting seeds, and eats livestock feed. However, is considered beneficial in some areas because more than half of the diet is animal material, including clover weevils, cutworms, and japanese beetles. Also eats a variety of domestic and wild fruits. REFERENCES: Forbush and May 1955, Kalmbach 1928, Kessel 1953, 1957, Planck 1967, Royall 1966, Small 1974. 374 White-eyed Vireo Vireo griseus L 4 Vi" RANGE: Breeds from southeastern Nebraska and central Iowa to southern Michigan, southern Ontario, and southern Massachusetts, south through eastern Texas and Florida into eastern Mexico. Winters from southern Texas south to Central America and east across the Gulf Coast to Florida, north to central coastal North Carolina. STATUS: Common. HABITAT: Inhabits deciduous thickets, woodland edges, brambles, undergrowth, hedgerows, and the dense understory of bottomland woodlands, generally favoring thickets near water. May inhabit highlands in the South. SPECIAL HABITAT REQUIREMENTS: Low shrubby vegetation. NEST: Builds a cone-shaped cup nest that is suspended from forked twigs of a low shrub or tree, usually well concealed, 1 to 8 feet above the ground. FOOD: Gleans its food, primarily insects, from branches and leaves. Eats mostly animal matter (90 percent of diet), but eats some berries and small fruits in winter. REFERENCES: Chapin 1925, Forbush and May 1955, Johnsgard 1979, Nolan and Wooldridge 1962, Sykes in Farrand 1983c. 375 Bell’s Vireo Vireo bellii RANGE: Breeds from southern California (local and rare), southern Nevada, Arizona, southern New Mexico, north into the Midwest (east of the Rocky Mountains) to North Dakota and east to Illinois and south to southwestern Tennessee, Arkansas, northwestern Louisiana, Texas, and Mexico. Winters from Mexico south to Central America. STATUS: Rare to absent from some former ranges in California, declining in Kansas, Oklahoma, and Texas. (Some decline may result from cowbird parasitism.) HABITAT: Throughout most its range, inhabits streamside willows. In the arid Southwest, lives along water courses and marshes where mesquite is mixed with cottonwood, saltcedar, elderberry, and desert hackberry. In the Great Plains, generally associated with thickets near streams and rivers, or with second-growth scrub, forest edges, and brush patches. SPECIAL HABITAT REQUIREMENTS: Dense riparian shrubs. NEST: Builds a small, basketlike cup nest attached to a forked branch of mesquite, hackberry, catclaw, oak, willow, ash, cottonwood, or low shrub, usually near water and seldom more than 5 feet above the ground. FOOD: Mostly eats animal matter (insects and spiders), gleaned from leaves and branches; also eats a few berries. REFERENCES: Chapin 1925, Forbush and May 1955, Johnsgard 1979, Tate and Tate 1982, Verner and Boss 1980, Terrill in Farrand 1983c. 376 Black-capped Vireo RANGE: Breeds in central Oklahoma locally through central and western Texas to north-central Mexico. Winters mainly in western Mexico. STATUS: Fairly common to uncommon. Populations decreasing. HABITAT: Generally inhabits the dense, low, ragged-topped thickets growing in hot, rocky hillsides, including stands of oaks, mescalbean, sumac, cedar, or other chaparral brush; prefers scrub oaks. May also be found in prairie ravines and early successional stages of brushlands. Prefers habitat arranged in rectangular or oval formation instead of linear. SPECIAL HABITAT REQUIREMENTS: Low, dense shrubs or trees. NEST: Builds a deep, cuplike nest suspended from a fork of slender twigs in trees or shrubs, usually 2 to 6 feet above the ground. Prefers oaks in parts of its range, but uses other tree and shrub species. FOOD: Gleans insects from leaves of shrubs (especially oaks). Also eats a few spiders and small fleshy fruits. REFERENCES: Graber 1961, Kaufman in Farrand 1983c, Johnsgard 1979, Oberholser 1974b. 377 Gray Vireo Vireo vicinior RANGE: Breeds locally from southern California, southern Nevada, southern Utah, and northwestern and central New Mexico south to Baja California, central and southeastern Arizona, southern New Mexico, western Texas, and central Mexico. Also in western Oklahoma. Winters ir southern Arizona, western Texas, and western Mexico. STATUS: Rare to locally common. HABITAT: Inhabits thorn scrub, oak-juniper woodland, pinyon-juniper, dry chaparral, mesquite, and riparian willow habitats. Favors dry chaparral that forms a continuous zone of twigs 1 to 5 feet above ground. SPECIAL HABITAT REQUIREMENTS: Low, dense shrubs. NEST: Builds a basketlike cup nest that is suspended 3 to 8 feet above the ground from the forks of twigs in a variety of low, thorny shrubs and small trees. FOOD: Gleans food (mostly insects) from leaves and branches of scrub oaks and other thickets. REFERENCES: Chapin 1925, Johnsgard 1979, Terres 1980, Terrill in Farrand 1983c. 378 Solitary Vireo 'ireo solitarius L4%" tocky Mt. ace RANGE: Breeds from central British Columbia east through central Canada to northern Ontario and Newfoundland, southwest of and through the Rockies to southern California and west Texas, south through Mexico to Honduras, and east of the Rockies to North Dakota, Illinois, and Massachusetts; in the Appalachian and Piedmont regions to eastern Tennessee, Alabama, Georgia, South Carolina, North Carolina, Virginia, and Maryland. Winters from southern California, central Texas, the northern portions of the Gulf States and North Carolina south to Costa Rica. STATUS: Common. HABITAT: Usually inhabits coniferous or coniferous-deciduous forests, especially spruce and tamarack swamps in parts of its range. Seems to prefer open mixed forests with considerable undergrowth. NEST: Builds a deep cup nest that is suspended from the fork of a horizontal branch, generally 3 to 20 feet above the ground, often about midway in a small conifer, but occasionally in a small deciduous tree or shrub. FOOD: Gleans most food from twigs and foliage but occasionally hawks for flying insects. Mostly eats insects, plus a few spiders and small fruits. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Harrison 1975, Johnsgard 1979, Petersen in Farrand 1983c, Terres 1980. 379 Yellow-throated Vireo Vireo flavifrons RANGE: Breeds from southern Manitoba, Minnesota, southern Ontario, New Hampshire and southwestern Maine south to eastern Texas, the Gulf Coast and central Florida and west to the Dakotas, Nebraska, Kansas, Oklahoma, and west-central Texas. Winters mainly in Mexico and Central and South America, but a few winter in Florida. STATUS: Rather uncommon to common. HABITAT: Generally associated with mature, moist deciduous forests, especially river bottom forests or north-facing slopes (in southern parts of range), and prefers open woodlands with partially open canopies. Found less frequently in wooded residential areas, seldom in dense deciduous forests, and rarely in coniferous forests. SPECIAL HABITAT REQUIREMENTS: Open, mature deciduous woodlands. NEST: Suspends nest between the forks of a slender branch, usually near the trunk, of a deciduous tree, preferably a large oak, normally over 30 feet above the ground (range 3 to 60 feet.) FOOD: Gleans most of its food from branches and foliage; occasionally hawks for flying insects. Mostly eats insects, spiders, and a few snails, but also a few small fruits and berries. REFERENCES: Chapin 1925, Forbush and May 1955, James 1976, Johnsgard 1979. 380 Hutton’s Vireo 'ireo huttoni RANGE: Resident from southwestern British Columbia south through western Washington, Oregon, and California to Baja California, and from central Arizona, southwestern New Mexico, and western Texas south to Guatemala. STATUS: Fairly common. HABITAT: Primarily inhabits live oak forests but also pine-oak communities; mountain canyons with sycamores, maples, and tall chaparral; or streamside willows. Prefers tree stands with open canopies. SPECIAL HABITAT REQUIREMENTS: Live oaks. NEST: Builds a hanging cup nest that is anchored to horizontal twigs and usually near the tips of the branches, mostly in oaks. Locates nest 7 to 25 feet above the ground. FOOD: Gleans food from foliage and small twigs, and occasionally hawks flying insects. Mostly eats insects, plus some spiders and small berries. REFERENCES: Chapin 1925, Harrison 1979, Verner and Boss 1980, Terrill in Farrand 1983c. 381 Warbling Vireo Vireo gilvus RANGE: Breeds from southeastern Alaska, northern British Columbia, and southern Mackenzie southeast to southern Ontario and southern New Brunswick, south to northern Mexico, Alabama, and Virginia. Winters mostly in Mexico and Central America. STATUS: Common and widespread. HABITAT: Inhabits open deciduous and mixed deciduous-coniferous forests, especially streamside vegetation, but also in groves, scrubby hillside trees, and residential areas. In mixed forests, generally associated with the deciduous trees, and prefers forests with a substantial forb or shrub layer and low to intermediate canopy cover. SPECIAL HABITAT REQUIREMENTS: Scattered deciduous trees or wooded streamsides. NEST: Builds a cup nest that is usually suspended from a horizontal branch of a deciduous tree, often poplar or aspen, generally in branches well away from the tree trunk and higher than those of other vireos (20 to 90 feet above the ground). FOOD: Gleans much of its food from the mid to upper canopy of deciduous trees. Eats mostly animal matter but includes some small fruits. REFERENCES: Chapin 1925, DeGraff et al. 1980, Forbush and May 1955, Harrison 1975, James 1976, Johnsgard 1979. 382 Philadelphia Vireo Vireo philadelphicus extremes RANGE: Breeds from east-central British Columbia to central Manitoba and southwestern Newfoundland, south to south-central Alberta, north- central North Dakota, northeastern Minnesota, southern Ontario, northern New Hampshire, northern Vermont, and Maine. Winters in Central America. STATUS: Uncommon to rare. HABITAT: Inhabits open deciduous, coniferous, or mixed forests, wood¬ land edges, burned or cutover areas with young deciduous regeneration, and willow and alder thickets along streams. Is restricted to aspen groves in parts of its range. NEST: Builds a deep cup nest that is suspended from a horizontal, forked branch of a deciduous tree or shrub, usually 10 to 40 feet above the ground. FOOD: Mainly (93 percent) eats animal material (insects and some spiders), most of which is gleaned from foliage and branches; catches some flying insects by hawking. Also eats some wild fruits. REFERENCES: Chapin 1925, DeGraff et al. 1980, Johnsgard 1979, Petersen in Farrand 1983c. 383 Red-eyed Vireo Vireo olivaceus (includes Yellow-green Vireo (Vireo flavoviridis)) RANGE: Breeds from southwestern British Columbia and southern Mackenzie southeast to central Ontario and the Maritime Provinces, south to northern Oregon, eastern Colorado, western Oklahoma to central Texas, the Gulf Coast, and central Florida. Very rare in California, Arizona, and southern Texas. Winters in South America. STATUS: Abundant; rare in the Southwest. HABITAT: Inhabits open deciduous and mixed forests with dense understory of saplings, in wooded clearings, or borders of burns. Found in both upland and river-bottom forests, and sometimes in residential areas where abundant shade trees provide a continuous canopy. Seldom found where conifers make up 75 percent or more of the basal area. SPECIAL HABITAT REQUIREMENTS: Deciduous trees with dense understory. NEST: Builds nest in deciduous or coniferous trees or shrubs. Suspends deep cup nest from a horizontal fork of a slender branch, usually in dense foliage 5 to 10 feet above the ground, but sometimes as high as 60 feet. FOOD: Consumes insects, gleaned from leaf surfaces in mid to upper tree canopies, for about 85 percent of the diet. Also eats spiders, a few snails, wild fruits, and berries. REFERENCES: Chapin 1925, Forbush and May 1955, James 1976, Johnsgard 1979, Laurence 1953a. 384 Black-whiskered Vireo /ireo altiloquus RANGE: Breeds in central and southern Florida and on islands in the Caribbean; winters in South America. STATUS: Common within much of its limited United States range. HABITAT: Lives mostly in mangroves along the coasts of Florida and the Florida Keys. SPECIAL HABITAT REQUIREMENTS: Dense mangroves. NEST: Suspends deep cup nest between twigs (usually in red mangroves but occasionally in other trees or shrubs) as high as 15 feet above the ground or water. FOOD: Gleans food, primarily spiders, from foliage. Also eats insects, fruits of barberries, and ragweed seeds. REFERENCES: Bent 1950, Chapin 1925, Sykes in Farrand 1983c, Terres 1980. 385 Bachman’s Warbler Vermivora bachmanii RANGE: Formerly bred in northeastern Arkansas, southeastern Missouri, south-central Kentucky, central Alabama, and southeastern South Caro¬ lina. May still breed in South Carolina. Winters in Cuba and the Isle of Pines. STATUS: Rarest songbird in North America; endangered, possibly extinct. Reasons for its decline are not clear. HABITAT: Inhabits low, moist, deciduous woodlands and swamps of the southern coastal plain, where it probably occurred primarily in sweet bay-swamp tupelo-red maple associations of headwater swamps, sweet gum-willow oak associations of wet flats, and in bottomland hardwoods. Appeared to use forest bottomlands and headwater swamps that were inundated with water for relatively short periods of time. SPECIAL HABITAT REQUIREMENTS: Mature deciduous swamp forests NEST: Locates nest in canebreaks and thickets 2 to 5 feet above the ground in and along the margins of low, wet forested habitats. (Little is known about its nesting habits.) FOOD: Forages in dense foliage high up in trees for spiders and insects. (Little is known about feeding habits.) REFERENCES: Griscom and Sprunt 1979, Hooper and Hamel 1977, Mackenzie 1977, Meanley and Mitchell 1958, Sykes in Farrand 1983c. 386 Blue-winged Warbler L4V," Vermivora pinus RANGE: Breeds from eastern Nebraska and southeastern Minnesota east to southern Vermont and southern Maine, and south to northwestern Arkansas, northern Alabama, northern Georgia, western South Carolina, and Delaware. Winters in Mexico south to Central America. STATUS: Uncommon. HABITAT: Inhabits rank growth near the borders of swamps or streams, woodland edges, brushy overgrown fields and pastures, thickets, and second-growth woods. Prefers old fields with saplings greater than 10 feet tall. Prefers more moist habitats than the golden-winged warbler, a closely related species with which it competes and, in some areas, hybridizes. SPECIAL HABITAT REQUIREMENTS: Brushy habitats near water. NEST: Builds nests on the ground, attached to upright stems of weeds or grass clumps. Usually nests in loose aggregations or colonies among bushes, ferns, tangles of vines, or grasses. FOOD: Gleans insects and spiders from leaves, twigs, and buds among branches near the tops of trees. REFERENCES: Confer and Knapp 1981, DeGraff et al. 1980, Griscom and Sprunt 1979, Harrison 1975, Johnsgard 1979, Robbins et al. 1983. 387 Golden-winged Warbler Vermivora chrysoptera L 4 Vi" 9 RANGE: Breeds from southern Manitoba, central Minnesota, and north¬ ern Wisconsin east to southern Vermont and eastern Massachusetts, and south to southeastern Iowa, southern Ohio, and southern Connec¬ ticut, and in the Appalachian Mountains south to northern Georgia. Breeding range in the Northeast and Appalachians has been decreasing in recent years, partly as a result of displacement by blue-winged warblers. Winters in Central and South America. STATUS: Locally common. HABITAT: Inhabits openings in deciduous forests or forest edges where there is a dense understory of forbs, grasses, or ferns. Also commonly inhabits damp fields heavily vegetated with thick grass, overgrown pastures, dense scrubby thickets, second-growth woods, and brush- bordered lowland areas. Generally occupies higher and drier areas than the blue-winged warbler, although there is a broad overlap in habitats. SPECIAL HABITAT REQUIREMENTS: Brushy edge habitats or openings in cover with saplings, forbs, and grasses. NEST: Nests in loose aggregations or colonies on or close to the ground. Usually locates nest within the shade of a forest edge, supported by weed stalks such as goldenrod, or by tufts of grass, or on a substrate of dead leaves. FOOD: Gleans insects and spiders chickadeelike, from the ground to tree tops. REFERENCES: Confer and Knapp 1981, DeGraff et al. 1980, Eyer 1963, Griscom and Sprunt 1979, Johnsgard 1979, Parkes in Farrand 1983c, Tate and Tate 1982. 388 Tennessee Warbler Vermivora peregrina L 4 Va" RANGE: Breeds from southeastern Alaska and southern Yukon across Canada to north-central Quebec and southern Labrador, and south to south-central British Columbia, northwestern Montana, northern Minnesota, northeastern New York, and southern Maine. Winters from Mexico to South America. STATUS: Common. HABITAT: In northern coniferous and deciduous woodlands, inhabits forest openings with grasses, dense shrubs, and scattered clumps of young trees, open spruce and tamarack or white cedar bogs where sphagnum moss is abundant, brushy hillsides, and occasionally dry pine lands. (Little is known about its breeding biology.) SPECIAL HABITAT REQUIREMENTS: Brushy, semi-open country. NEST: Nests singly or sometimes in loose colonies. Conceals nest on moist ground, typically in sphagnum-covered hummocks or among grasses, protected by bog plants overhead, or less frequently, on dry hillsides under the cover of shrubs or saplings. FOOD: Forages on terminal twigs, gleaning while hopping from branch to branch, eating almost entirely insects. Also eats some spiders and fruits. REFERENCES: Bent 1953a, DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979. 389 Orange-crowned Warbler Vermivora celata RANGE: Breeds from western and central Alaska and central Yukon across Canada to northern Ontario, central Quebec, and southern Labrador south to southwestern and central California, central Utah, southern New Mexico, and extreme western Texas, and east of the Rockies, to southern Saskatchewan, central Ontario, and south-central Quebec. Winters from coastal and southern California, central Arizona, Texas, the southern portion of the Gulf States, and South Carolina south to Central America. STATUS: Common in the West; rare in the East. HABITAT: Occurs in a variety of woodland and brushy habitats, especially sites with considerable shrub cover. Prefers chaparral, brushy open woods, woodland edges of low deciduous growth, burns, overgrown pastures, riparian thickets, and the edges of clearings. In Oregon, found in the mountains up to 6,000 feet, inhabiting dense mixed groves of aspen, alder, willow, and pine in meadows of subalpine parks. SPECIAL HABITAT REQUIREMENTS: Dense shrubs for nesting. NEST: Conceals nest on the ground in a bramble tangle, hummock, at the base of a bush or stump, or occasionally up to 3 feet high in low, dense bushes. FOOD: Gleans insects from foliage in shrubs and small trees at heights ranging from 5 to 30 feet. Also eats some spiders, leaf galls, seeds, and fruits. REFERENCES: Bent 1953a, Griscom and Sprunt 1979, Harrison 1979, Verner and Boss 1980. 390 slashville Warbler 1 ermivora ruficapilla RANGE: Breeds from southern interior British Columbia and northwestern Montana south to northwestern and south-central California and extreme west-central Nevada; and from central Saskatchewan to southern Quebec, Nova Scotia, and New Brunswick, south to southern Manitoba, southern Wisconsin, southern Michigan, northern New Jersey, and Rhode Island. Winters from southern Texas south to Mexico and Central America, rarely in California and southern Florida. STATUS: Common. HABITAT: Prefers brushy sphagnum bogs and open second-growth woodlands. Also occurs in regenerating areas that have been burned or cut, overgrown pastures and fields, woodland edges, and in swales, slashings, and undergrowth of mixed forests, especially those with aspen or birch. Also found in woodlands, but generally on poor sites. SPECIAL HABITAT REQUIREMENTS: Scattered trees interspersed with brush. NEST: Conceals nest on the ground in a small depression, sometimes in a sphagnum hummock, often with an overhead cover of ferns or other overhanging vegetation. FOOD: Forages from the ground to the treetops, gleaning food (chiefly insects) from the trunk, leaves, and branches, and hawking flying insects. REFERENCES: DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979, Lawrence 1948, Petersen in Farrand 1983c. 391 Virginia’s Warbler Vermivora virginiae RANGE: Breeds from east-central California, central Nevada, southeast¬ ern Idaho, and southern Wyoming south to south-central California, central and southeastern Arizona, southern New Mexico, and extreme western Texas. Winters in Mexico. STATUS: Common. HABITAT: Inhabits arid montane woodlands from 6,000 to 9,000 feet, preferring scrubby brush interspersed with pinyon-juniper and yellow pine. Frequents dense growths of low scrub oaks, mountain-mahogany, and chokecherry, rocky steep slopes and ravines, chaparral, riparian willow and alder thickets, and open spruce and fir forests near scrubby thickets. SPECIAL HABITAT REQUIREMENTS: Scrubby vegetation for nesting. NEST: Builds nest on the ground, embedded among dead leaves or in loose soil, sometimes at the base of a bush, or hidden under a tussock of grass, but usually concealed by overhanging vegetation. FOOD: Forages on the ground, as well as in foliage, and hawks insects on the wing. REFERENCES: Bent 1953a, Griscom and Sprunt 1979, Terrill in Farrand 1983c, Van Tyne 1936. 392 Colima Warbler Vermivora crissalis L 43/4" RANGE: Breeds in the Chisos Mountains in extreme western Texas and northern Mexico. Winters in Mexico. STATUS: Rare and local. HABITAT: Inhabits forested canyons and slopes between 6,000 and 7,000 feet, where it frequents thickets of young maples and oaks along dry stream beds, clumps of small oaks along mountain slopes, and mixed woods of maple, oak, Arizona cypress, and yellow pine. SPECIAL HABITAT REQUIREMENTS: Oak thickets above 6,000 feet. NEST: Nests on the ground among fallen leaves and vines, which may partly or completely conceal the nest. Locates nest on rocky slopes, or adjacent to dry stream beds among small rocks and leaves where there are clumps of small oaks. FOOD: Occupies the lower tree branches and forages for caterpillars and other insects in Arizona cypress and neighboring vegetation. (No food studies on this species in the United States.) REFERENCES: Blake 1949, Griscom and Sprunt 1979, Oberholser 1974b, Van Tyne 1936. 393 Lucy’s Warbler Vermivora luciae RANGE: Breeds from southeastern California, southern Nevada, and Utah south to southern Arizona, northern Mexico, and extreme western Texas. Winters in western Mexico. STATUS: Common. HABITAT: Found in southwestern deserts, generally wherever there are large mesquites, especially along main watercourses. Also occurs in mountain foothills in streamside cottonwoods and willows. SPECIAL HABITAT REQUIREMENTS: Cavities for nesting. NEST: The only cavity-nesting warbler besides the prothonotary, places nest in four types of cavities; natural cavities in trees (usually mesquite), where the entrance is in a sheltered spot; under loose bark; in aban¬ doned woodpecker holes; and in deserted verdin nests. Generally locates nest 5 to 6 feet above the ground, but ranges from 1 to 15 feet. Occasionally nests in holes in banks, in yuccas, willows, sycamores, or elderberries. FOOD: Forages for insects at middle heights in mesquite, seldom in the treetops or near the ground. REFERENCES: Bent 1953a, Griscom and Sprunt 1979. 394 slorthern Parula RANGE: Breeds from southeastern Manitoba and central Ontario east to New Brunswick south to south-central and southern Texas, the Gulf Coast, and Florida, and west to the eastern edge of the Plains States. Winters in Florida and from Mexico to Central America. STATUS: Common. HABITAT: Primarily associated with swampy woods, especially in the Southeast, where it inhabits mature coniferous and deciduous woodlands where mosslike lichens or Spanish moss are found. In the North, found in swamps and bogs with abundant bearded lichens, in the South, frequents trees draped with Spanish moss. Occasionally occurs in woodlands without moss or lichens. SPECIAL HABITAT REQUIREMENTS: Bearded lichen (Usnea) or Spanish moss (Tillandsia) for nesting material or for nest sites. NEST: Usually suspends nest near the tip of a tree limb that is covered with Usnea or Tillandsia at heights averaging 10 feet, but ranging from 6 to 100 feet above the ground. Conceals nest with moss or lichen, and constructs it of these materials as well. In areas where they are not available, uses other materials. FOOD: Forages in a chickadeelike fashion, gleaning insects and spiders from twigs and foliage. REFERENCES: Bent 1953a, DeGraff et al. 1980, Graber and Graber 1951, Griscom and Sprunt 1979, Johnsgard 1979. 395 Tropical Parula Parula pitiayumi RANGE: Resident from southern Texas in the lower Rio Grande Valley north to near Kingsville, Texas and south locally through Central AmeriCc into South America. Widespread but local in tropics; largely withdraws ir winter from northern limits of range. STATUS: Scarce; once common, the population in southern Texas has declined, probably because of loss of prefered habitat, pesticide pollution, and the greatly expanded population of bronzed cowbirds. HABITAT: Found in dense or open woodlands; inhabits undergrowth, brush and trees along the edges of rivers, low dry woodlands, and semiarid cultivated valleys with scattered trees. Breeds from near sea level to 200 feet in elevation in southern Texas, preferring habitats with trees whose branches are draped with Spanish moss. SPECIAL HABITAT REQUIREMENTS: Air plants (epiphytes) such as Spanish moss for nesting. NEST: Builds nest 8 to 10 feet above the ground in a hollow formed by c mass of Spanish moss or other air plants that hang in trees or bushes. Nests on relatively level land along rivers and streams, in tall brush or timber. FOOD: Mostly eats insects. (Little is known about the diet in the United States.) REFERENCES: Bent 1953a, Griscom and Sprunt 1979, Kaufman in Farrand 1983c, Oberholser 1974b. 396 fellow Warbler lendroica petechia RANGE: Breeds from northwestern and north-central Alaska and northern Yukon to northern Ontario, central Quebec, and southern Labrador south to Mexico, central and northeastern Texas, northern Arkansas, central Georgia, and central South Carolina. Winters from southern California, southwestern Arizona, Mexico, and southern Florida south to South America. Also resident in southern Florida. STATUS: Common; significant population declines in Idaho and the Dakotas, and numbers are declining in many other areas. HABITAT: Prefers moist habitats such as willow- and alder-lined streams and ponds, brushy bogs, and the edges of marshes, swamps, or creeks. Also occurs in dry sites such as hedgerows, roadside thickets, orchards, farmlands, forest edges, and suburban yards and gardens. Generally occurs wherever patches of trees or shrubs grow, but avoids heavy forests. In the Florida Keys, only inhabits coastal mangroves. SPECIAL HABITAT REQUIREMENTS: Small scattered trees or dense shrubbery. NEST: Usually builds nest in an upright fork or crotch of a tree or bush, typically 3 to 8 feet, occasionally up to 40 feet, above the ground. May nest colonially in ideal habitats. Frequently victimized by cowbirds, builds up the nest lining to cover the cowbird eggs. FOOD: Forages for insects and spiders on the limbs of shrubs and trees by gleaning and hawking. REFERENCES: Bent 1953a, Bigglestone 1913, DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979, Schrantz 1943, Tate and Tate 1982, Vickery in Farrand 1983c. 397 Chestnut-sided Warbler Dendroica pensylvanica RANGE: Breeds from east-central Alberta and central Saskatchewan to southern Quebec and New Brunswick south to eastern Colorado, Iowa, central Ohio, and Massachusetts; in the Appalachians south to north- central Georgia and northwestern South Carolina. Winters from Mexico to Panama, casually to South America. STATUS: Common. HABITAT: Inhabits rather open and dry areas having some woody vegetation in the form of shrubs and small trees, preferring the brushy regrowth of clear-cut forests and abandoned pastures and fields. Also frequents second-growth woodland edges and clearings, low shrubbery, briar thickets, brushy hillsides and brooksides, and roadside thickets. SPECIAL HABITAT REQUIREMENTS: Early second-growth moist woodlands with dense vegetation about 3 feet above the ground to provide nest sites and foraging areas. NEST: Conceals nest from 1 to 4 feet above the ground in briar thickets, bushes, saplings, or vines, such as spirea, raspberry, red-osier dogwood, azalea, laurel, gooseberry, meadow-rue, and hazel. FOOD: Consumes insects and spiders gleaned from the foliage of shrubs and other low plants, or caught by flycatching. Occasionally eats a few seeds and berries. REFERENCES: Cripps 1966, Griscom and Sprunt 1979, Lawrence 1948, Tate 1970, Terres 1980, Vickery in Farrand 1983c. 398 Magnolia Warbler Dendroica magnoiia L 4‘A" RANGE: Breeds from northeastern British Columbia, west-central and southern Mackenzie east to north-central Manitoba and south-centra and eastern Quebec, and south to south-central British Columbia, central Saskatchewan, northeastern Minnesota, central Michigan, western Virginia, northwestern New Jersey, and Connecticut. Winters from Mexico to Panama. STATUS: Common. HABITAT: Inhabits open coniferous or mixed coniferous-deciduous woodlands, preferring spruce and fir forests with low trees, and coniferous bogs. Also inhabits dense thickets of spruce and fir, old clearings with small coniferous saplings, second-growth following logging, woodland edges, and coniferous thickets along roadsides. SPECIAL HABITAT REQUIREMENTS: Young conifer stands for nesting. NEST- Conceals nest in a small conifer, in foliage near the tip of a horizontal branch, generally 1 to 8 feet, but typically less than 5 feet, above the ground. Prefers spruce, fir, and hemlock for nesting, but may use hardwoods. FOOD: Tends to forage near the center of a tree for insects, rather than at the outer edges; almost entirely insectivorous. REFERENCES: Bent 1953a, DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979, Petersen in Farrand 1983c. 399 Cape May Warbler Dendroica tigrina RANGE: Breeds from southwestern and south-central Mackenzie and northeastern British Columbia east to central Ontario, southern Quebec and New Brunswick south to central Alberta, southeastern Manitoba, northern Wisconsin, southern Ontario, northeastern New York, and east- central Maine. Winters in central and southern Florida and the West Indies. STATUS: Uncommon. HABITAT: Inhabits fairly open coniferous forests, especially those with a high percentage of mature spruce. Also frequents dense spruce forests with a scattering of tall spires extending above the canopy level, the edges of coniferous forests, especially if birches or hemlocks are present, and in more open land among small trees. Will occasionally proliferate in areas heavily infested with spruce budworms. SPECIAL HABITAT REQUIREMENTS: Tall conifer trees, especially spruce, for nesting. NEST: Builds nest invariably in the uppermost clump of thick foliage near the top of tall conifers, generally invisible from below. (Little is known about the nesting biology of this species, mainly because it tends to nest so high in conifers, and females tend to land near the tree base and move up through the tree rather than fly to the nest.) FOOD: Consumes insects caught by flycatching or gleaning the tips of dense branches and new buds of conifers near the tops of trees. Also eats spiders and occasionally vineyard grapes. REFERENCES: Bent 1953a, DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979, Petersen in Farrand 1983c. 400 Jlack-throated Blue Warbler »endroica caerulescens RANGE: Breeds from western and central Ontario to New Brunswick south to northeastern Minnesota, central Michigan, northeastern Pennsylvania, and southern New England, and in the Appalachians to northeastern Georgia and northwestern South Carolina. Winters primarily from southern Florida to the West Indies, casually in Central and South America. STATUS: Common. HABITAT: Prefers northern hardwood forests with an ample undergrowth of saplings and evergreen or deciduous shrubs. Also inhabits mature coniferous-deciduous forests, especially those with an undergrowth of creeping yew, laurel, hazel, maple, or brushy saplings. In the southern Appalachians, often inhabits dense mountain-laurel thickets. SPECIAL HABITAT REQUIREMENTS: Woodlands with shrubby undergrowth. NEST: Constructs well-concealed nest near the ground, generally not higher than 3 feet above ground, in fallen tree tops, small trees, or shrubs. Partial to nesting in rhododendron, laurel, hemlock, small spruce, fir, and maple. FOOD: Forages in the shrub, subcanopy, and lower canopy layers of forests, gleaning insects from foliage and branches. Mostly consumes insects but occasionally eats seeds and berries. REFERENCES: Bent 1953a, DeGraff et al. 1980, Griscom and Sprunt 1979, Flarding 1931, Johnsgard 1979, Petersen in Farrand 1983c. 401 Yellow-rumped Warbler Dendroica L 4 3 /4" Audubon' RANGE: The northern and eastern (Myrtle) race breeds from western Alaska and central Mackenzie to north-central Labrador south to northern British Columbia, southeastern Saskatchewan, central Michigan, and Massachusetts, and in the Appalachians to eastern West Virginia. The western (Audubon’s) race breeds from central British Columbia and southwestern Saskatchewan south to southern California across to western Texas. The Myrtle race winters from southwestern British Columbia through the Pacific States, southern Arizona, and Colorado, and from Kansas east across the central United States to New England, and south to Panama. The Audubon’s race winters from southwestern British Columbia and Idaho south to Central America. STATUS: Common. HABITAT: Generally inhabits coniferous forests throughout its range, but also found in aspen forests in the Rocky Mountains. In the West, prefers timbered sites with a low percent canopy cover, and forest edges such as those around meadows or lakes. In the East, prefers spruce-fir woodlands, but also frequents young coniferous growth near the edges of woods, mixed woods, and evergreen plantations. SPECIAL HABITAT REQUIREMENTS: Coniferous trees for nesting. NEST: Builds nest well out on a horizontal branch in a conifer tree, screened from above by clumps of needles. Typically locates nest 15 to 20 feet above the ground, but sometimes 4 to 50 feet. FOOD: Eats insects gleaned from foliage and caught by hawking. Also searches for food on the ground. In winter, largely subsists on berries. REFERENCES: Beal and McAtee 1912, Bent 1953a, DeGraff et al. 1980, Griscom and Sprunt 1979, Johnsgard 1979, Kaufman in Farrand 1983c, Verner and Boss 1980. 402 Black-throated Gray Warbler Dendroica nigrescens L 4" RANGE- Breeds from southwestern British Columbia and western A/ashington to northern Utah and southern Wyoming south, primarily in mountains, to northern Baja California, central and southeastern Arizona, sxtreme western Texas, and Mexico. Winters from coastal southern California and southern Arizona south to Mexico. STATUS: Common. HABITAT: In the northern portion of its range, inhabits conifer forests that are open and interspersed with shrubs or forest edges. Farther south, seems to prefer shrubby stands of oaks, pinyon, juniper, and manzanita. Throughout its range, prefers and is perhaps limited to dry slopes. NEST- In Washington, seems to nest exclusively in fir trees, placing nest on horizontal branches 7 to 50 feet above the ground. In habitats further south, builds nests in shrubs such as manzanita, oak, ceanothus, or large white oaks and sycamores. FOOD: Forages among the leaves of shrubs for insects. (Detailed food habits have not been reported.) REFERENCES: Bent 1953a, Griscom and Sprunt 1979, Phillips et al. 1964. 403 Townsend’s Warbler Dendroica townsendi RANGE: Breeds from east-central Alaska, southern Yukon, and coastal British Columbia, as well as parts of Alberta and Saskatchewan, south along the Pacific Coast into northwestern Washington, and inland to central and southeastern Washington, central and northeastern Oregon, northern Idaho, northwestern and south-central Montana, and northwest¬ ern Wyoming. In general, breeding distribution follows that of conifers and mixed coniferous-deciduous forests. Winters in the coastal region of Oregon and California south through the highlands of Mexico and Central America to Costa Rica. STATUS: Common in coniferous forests of the Pacific Northwest. HABITAT: During breeding season, primarily inhabits tall coniferous forests of the Pacific Northwest, most commonly the more mountainous regions, usually near the crowns of tall trees. During migration and winter, moves into humid forests as well as pine-oak associations, open woodlands, and second-growth scrub forests. SPECIAL HABITAT REQUIREMENTS: Tall, coniferous forests of the north Pacific Coast. NEST: Nests in conifers, particularly firs, generally on the limb of the tree and not in the fork or crotch. Constructs nests of bark and slender twigs woven together, usually places them near the crowns of trees. In shorter trees, places nests as close as 10 or 12 feet to the ground. FOOD: Consumes insects found on the needles of conifers as primary diet during spring and summer. (Winter food habits are not well documented.) REFERENCES: Bent 1953a, Griscom and Sprunt 1979. 404 Hermit Warbler )endroica occidentalis RANGE: Breeds in coniferous forests from southern British Columbia south through the coastal ranges and Sierra Nevadas to southern California. Winters in Mexico and Central America. In general, distribution coincides with tall coniferous trees. STATUS: Uncommon and local in summer habitats. HABITAT: Uses very tall conifers, including Douglas-fir and cedar. Prefers scattered groups of tall trees (sometimes 200 or more feet tall) that tower above the rest of the forest. Generally found in the upper canopies of tall trees, and though very active, are often difficult to identify. SPECIAL HABITAT REQUIREMENTS: Tall coniferous trees. NEST: Builds a well-concealed nest, generally supported by needles on the scraggly limbs of conifers, often up to 40 or 50 feet above the ground. Generally forage on the upper portions of trees, while nests are built in the mid-canopy of some of the tall trees. FOOD: Feeds on a variety of insects found in the upper level of coniferous trees, gleaning them from needles and twigs, or hawking flying insects. REFERENCES: Bent 1953a, Griscom and Sprunt 1979. 405 Black-throated Green Warbler Dendroica virens $ RANGE: Breeds from east-central British Columbia and northern Alberta to central Ontario and Newfoundland south to central Alberta, southern Manitoba, central Minnesota, Pennsylvania, and northern New Jersey, and south in the mountains to northern Alabama and Georgia. Winters from southern Texas and southern Florida south to Central America and the West Indies. STATUS: Common. HABITAT: Inhabits open mixed woodlands (especially northern hardwood- hemlock stands), northern coniferous forests with large trees, and larch bogs. Less often, inhabits second-growth hardwoods and pastures with cedars. It occurs rather commonly in pine barrens in Maine and jack pines in Michigan. SPECIAL HABITAT REQUIREMENTS: Some coniferous cover. NEST: Builds a compact, deep cup nest, usually placed on a branch or in a fork of a conifer tree, 3 to 80 feet above the ground; occasionally uses a deciduous tree. FOOD: Largely consumes insects gleaned from leaves and branches, and occasionally hawks flying insects. REFERENCES: Griscom and Sprunt 1979, Harrison 1975, Pitelka 1940, Terres 1982. 406 Golden-cheeked Warbler Dendroica chrysoparia L W RANGE: Breeds from central Texas south to the Edward s Plateau region into Medina and Bexar Counties and west to Real and Kerr Counties. Winters in the highlands of Central America. Occasionally visits the Farallon Islands of California and isolated areas of Florida. STATUS: Uncommon and very local. HABIAT; Prefers small stands of mountain-cedar on rough woody hillsides in canyons or on ridges that separate headwaters of streams, and also inhabits mountain oak, black oak, and live oak thickets on higher grounds as well as oak thickets on the lower flats along the foothills. Populations have been eliminated by removal of “cedar brakes. Mature cedar brakes ranging from several hundred acres to a thousand or more are necessary to insure habitat for this specie. SPECIAL HABITAT REQUIREMENTS: Mountain-cedar in canyons or draws within 1-1/2 miles of water. NEST: Builds nests of cedar bark interspersed with webs, often from spiders. Fastens nests to limbs in the mid-canopy level of mountain- cedars and are difficult to locate because they resemble the bark of the tree. Often parasitized by cowbirds. FOOD: Eats a variety of insects and spiders. REFERENCES: Griscom and Sprunt 1979, Oberholser 1974b, Pulich 1976. 407 Blackburnian Warbler Dendroica fusca RANGE: Breeds from central Alberta east to southern Quebec and Nova Scotia south to southern Manitoba, northeastern Ohio, Pennsylvania, and southeastern New York, and in the Appalachians to South Carolina and northern Georgia. Winters in Central and South America. STATUS: Common. HABITAT: Throughout most of its range, favors mature conifer forests with a few deciduous trees, preferring the upper canopy of tall conifers. Also inhabits climax stands of conifers with sparse understory and with deciduous trees and shrubs around the edges. In the Appalachians, inhabits oak-hickory forests along ridges. SPECIAL HABITAT REQUIREMENTS: Tall coniferous forests; oaks in the Appalachians. NEST: Builds a deeply cupped nest saddled to a horizontal branch of a large tree. Nests in conifers throughout its range but also uses deciduous trees in the South. Usually locates nest high (up to 80 feet) above the ground and well out from the tree trunk. FOOD: Consumes mostly insects gleaned from branches and leaves, but also flying insects caught in the air. Eats some berries when insects are scarce. REFERENCES: Bull and Farrand 1977, Griscom and Sprunt 1979, Lawrence 1953b, Terres 1982. 408 fellow-throated Warbler endroica dominica L W 1ANGE: Breeds from southeastern Kansas, central Ohio, and central jew Jersey south to south-central and eastern Texas, the Gulf Coast, :entral Florida, and the northern Bahama Islands. Wanders north to the 3 re at Lakes and the Maritime Provinces. Winters from southeastern rexas, the Gulf Coast, and South Carolina south to Costa Rica. STATUS: Common in southeastern United States. HABITAT: Generally inhabits large trees along river banks, swamps, and oottomlands, as well as open stands of pines, live oaks, and mixed forests. In the South, prefers forests with abundant Spanish moss. Tends to utilize the upper canopy level of the forests. NEST: In coastal areas, nearly always builds nest in clumps of Spanish moss. In areas with no Spanish moss, saddles nest on a horizontal branch. Locates nests 10 to 100 feet above the ground, generally far out from the tree trunk. FOOD: Is a skillful “fly catcher,” but gleans much of its food from leaves and branches; eats nearly all insects. REFERENCES: Bull and Farrand 1977, Griscom and Sprunt 1979, Harrison 1975, Johnsgard 1979, Sykes in Farrand 1983, Terres 1982. 409 Grace’s Warbler Dendroica RANGE: Breeds from southern Nevada, southern Utah, southwestern Colorado, northern New Mexico, and western Texas south through the mountains of western Mexico to Nicaragua. Winters in northern Mexico; resident from central Mexico southward. STATUS: Locally common in pine-oak forests above 7,000 feet. HABITAT: Typically inhabits pine forests, usually in the upper portions of yellow pines. Sometimes inhabits hemlock and fir woodlands, and occasionally oak thickets; generally from 6,000 to 9,000 feet. SPECIAL HABITAT REQUIREMENTS: Pine forests approximately 7,000 feet in elevation. NEST: Locates nest on limbs of pine trees up to 60 feet above the ground, sometimes in the middle of bunches of pine needles. Builds a very compact nest composed of a variety of grass fibers, hair, vegetable material, and insect webbing, often hidden from view. FOOD: Commonly gleans food (almost entirely insects) from leaves and branches of pines, as well as hemlocks and spruces. Most commonly forages in the very upper canopies of the tree, but sometimes catches flying insects on the wing. REFERENCES: Griscom and Sprunt 1979. 410 Pine Warbler Dendroica pinus IANGE: Breeds from southern Manitoba, western Ontario, southwestern Quebec, and central Maine south to eastern Texas, the Gulf Coast, southern Florida, and the Bahamas. Rare or absent in the upper Mississippi and Ohio River Valleys. Winters in the southeastern United States south to southern Texas, the Gulf Coast, and southern Florida into Mexico. STATUS: Common to rare and local. HABITAT: Inhabits open pine forests and pine barrens, especially jack pine in Minnesota and upland southern pines. Generally avoids tall, moist, and dense coniferous forests. SPECIAL HABITAT REQUIREMENTS: Open pine forests. NEST: Builds nests saddled on horizontal limbs of conifers 8 to 80 (usually 30 to 50) feet above the ground, usually far out from the tree trunk and well concealed in foliage. FOOD: Gleans its food from tree trunks, larger branches, and leaves. In the summer, mostly eats insects and some spiders. In winter, also eats pine seeds, wild fruits and berries, and grass and weed seeds. REFERENCES: Bull and Farrand 1977, Griscom and Sprunt 1979, Harrison 1975. 411 Kirtland’s Warbler Dendroica kirtlandii L 4 3 A" RANGE: Breeds only in extensive tracts of small jack pines in a few counties of northern lower Michigan. Occasionally strays to similar habitats in Wisconsin, southern Ontario, and southern Quebec if not breeding. Winters throughout the Bahamas. STATUS: Endangered. Approximately 1,000 birds in existence. HABITAT: Breeds in very specific habitats: extensive stands (80 +' acres) of young jack pine that are 6 to 20 feet tall and have living pine branches near the ground. Usually moves into burned-over jack pine forests 6 to 13 years after fire and inhabits these young forests for 10 to 12 years. SPECIAL HABITAT REQUIREMENTS: Dense stands of young jack pine. NEST: Tends to nest in loose colonies. Conceals nest under low vegetation (particularly bluestem grass and blueberry) near the base of a small jack pine on flat, dry, porous soil, usually depressed below ground level; about half of nests are parasitized by brown-headed cowbirds. FOOD: Gleans food (mostly insects) from the ground and pine needles. REFERENCES: Griscom and Sprunt 1979, Mackenzie 1977, Mayfield 1960. 412 > rairie Warbler endroica discolor L 4" RANGE: Breeds from eastern Nebraska, central Missouri, northern Illinois, central Wisconsin, northern Michigan, southern Ontario, southeastern New York and New Hampshire south to eastern Texas , the Gulf Coast and southern Florida. Winters from central Florida south to the West Indies and Central America. STATUS: Common. HABITAT: Generally inhabits open brushy lands, often in mixed pine and scrub oak woodlands. Also inhabits southern pine forests, sand dunes, mangroves, and jack pine plains but tends to avoid dense forests. (Suitable habitats have increased on abandoned farms, unmowed orchards, strip mine lands, Christmas tree farms, and burned and grazed woodlands.) SPECIAL HABITAT REQUIREMENTS: Low trees and shrubs; tends to favor areas with some conifers. NEST: Sometimes nests in loose colonies. Attaches well-concealed cup nest to stems and branches of a variety of shrubs and trees, usually 2 to 3 feet above the ground. FOOD: Gleans food from tree leaves, branches, and the ground, but also catches flying insects in the air. Eats mostly insects, but also eats some snails and worms. REFERENCES: Griscom and Sprunt 1979, Nolan 1978. 413 Palm Warbler Dendroica palmarum 6 Western race g RANGE: Breeds from southern Mackenzie and northern Alberta to central Quebec and southern Newfoundland south to northeastern British Columbia, central Alberta, northern Minnesota, southern Quebec, Maine, and Nova Scotia. Winters mostly from north-central Texas to North Carolina south to southern Texas, the Gulf Coast, southern Florida, and islands in the Caribbean. STATUS: Fairly common. HABITAT: Inhabits boggy areas dominated by tamarack, black spruce, and white cedar, and dry, open forests of spruce or jack pine. NEST: Sometimes nests in loose colonies. Usually builds nest on the ground, nearly buried in sphagnum mosses, but may nest in the low branches of conifer saplings. FOOD: Forages on the ground, gleans food from twigs and conifer cones, and also catches flying insects. Mainly eats insects, but also some vegetable matter, especially barberries during winter. REFERENCES: Bent 1953b, Griscom and Sprunt 1979. 414 Jay-breasted Warbler endroica castanea 3ANGE: Breeds from southwestern Mackenzie to north-central Sas- /2" RANGE: Breeds from west-central Mackenzie, northern Alberta, and central Saskatchewan to southern Quebec and Newfoundland; south to eastern Montana, central Texas, Louisiana, Alabama, Georgia, and North Carolina. Winters from southern Texas and Florida through Mexico to South America. STATUS: Common. HABITAT: Generally associated with semiopen upland stands of deciduous or mixed forests; usually not abundant in coniferous forests. Found in mature and second-growth forests, especially those composed of immature or scrubby trees. NEST: Builds nest in a slight depression in the ground, usually at the base of a tree or stump, beside a log, or sometimes in the roots of a fallen tree. FOOD: Mainly eats insects, but it also gleans many spiders and daddy longlegs from tree trunks and larger tree limbs. REFERENCES: Bent 1953a, Forbush and May 1955, Griscom and Sprunt 1979, Smith 1934. 418 \merican Redstart ',etophaga ruticilla 9 RANGE: Breeds from southeastern Alaska, east to Labrador and Newfoundland south to Utah, southeastern Oklahoma, and east Texas to South Carolina. Absent as a breeding bird through most of the Great Plains region. Winters from Baja California, southern Texas, and central Florida south to Brazil. STATUS: Abundant. HABITAT: Prefers open deciduous woodlands with a good understory of shrubs and young trees but is very adaptable. Frequently nests in mixed coniferous-deciduous forests, shade trees and shrubbery around farms, orchards, and willow and alder thickets bordering ponds and streams. NEST: Normally builds its nest 10 to 20 feet above the ground in a crotch or on a horizontal limb of a second-growth deciduous tree. Frequently parasitized by brown-headed cowbirds when nesting outside of woodlands. FOOD: Chiefly eats insects caught in the air or gleaned from leaves and branches. Also eats some spiders, daddy longlegs, and fruits. REFERENCES: Baker 1944, Griscom and Sprunt 1979, Vickery in Farrand 1983c. 419 Prothonotary Warbler Protonotaria citrea d RANGE: Breeds from east-central and southeastern Minnesota, south- central Wisconsin, southern Michigan, southern Ontario, central New York and northern New Jersey south to south-central and eastern Texas, the Gulf Coast and central Florida, and west to eastern Oklahoma, eastern Kansas, and central Oklahoma. Winters rarely in Florida and extreme south; otherwise, migrates to eastern Central America and South America. STATUS: Uncommon. HABITAT: Generally associated with moist, bottomland, or swampy deciduous woods, including woods that are frequently flooded and willow-lined streamsides. SPECIAL HABITAT REQUIREMENTS: Moist woodlands with cavities for nesting. NEST: Nests in natural cavities and old cavities of woodpeckers (especially downy woodpeckers) and chickadees in stumps or snags that are standing in or near water. Will occasionally nest in nest boxes. Usually places nest low, about 5 feet above the ground. FOOD: Primarily eats insects gleaned from tree trunks and branches, shrubs, and fallen logs. REFERENCES: Bent 1953a, Forbush and May 1955, Griscom and Sprunt 1979, Pearson 1936, Walkinshaw 1953. 420 Vorm-eating Warbler 'elmitheros vermivorus ANGE: Breeds from southeastern Nebraska, southeastern Iowa to outhern and east-central Ohio and Massachusetts, south to southeastern (klahoma and northeastern Texas to northern Florida. Winters in the i/est Indies, Mexico, and Central America. TATUS: Locally common to rare. IABITAT: Inhabits wooded hillsides and ravines with medium-sized tands of deciduous trees and undergrowth, often near streams or wampy bogs rimmed by shrubs and vines. PECIAL HABITAT REQUIREMENTS: Dense undergrowth. IEST: Builds nest on the ground at the base of a tree or sapling, sually well concealed under dead leaves. Generally locates nest on a illside or bank of a ravine, but sometimes in a bank cavity or under hrubbery. : OOD: Gleans its food from the ground; mostly eats insects but some piders and a few worms. IEFERENCES: DeGraff et al. 1980, Griscom and Sprunt 1979. 421 Swainson’s Warbler Limnothlypis swainsonii RANGE: Breeds locally from northeastern Oklahoma, southern Missouri southern Illinois, southwestern Indiana, southwestern and eastern Kentucky, southern Ohio, western West Virginia, western and southern Virqinia, and southern Delaware south to east-central Texas, the Gu Coast, and northern Florida. Winters on the Caribbean islands and in southeast Mexico. STATUS: Uncommon. HABITAT: Generally inhabits rich, damp woodlands with deep shade an- dense undergrowth, including wooded swamps and canebrakes of lowlands and, locally, rhododendron thickets of the mountains. SPECIAL HABITAT REQUIREMENTS: Dense underbrush. NEST: Builds a large bulky nest, usually 2 to 6 feet above the ground. I coastal lowlands, commonly nests in cane or palmetto; in highlands, nests in shrubs, small trees, vines, briars, rhododendron, or laurel. Sometimes parasitized by cowbirds. FOOD: Consumes primarily insects and spiders found in leaves on the ground, but occasionally searches for food in low shrubs. REFERENCES: Harrison 1975, Meanley 1966, 1971. 422 Dvenbird Je/'urus aurocapillus RANGE: Breeds from northeastern British Columbia, southern Mackenzie, northern Alberta, across southern Canada to Newfoundland south to eastern Colorado, eastern Oklahoma, northern Arkansas, and the mid- Atlantic States to northern Georgia. Winters in coastal South Carolina, Florida, the Gulf States, coastal Texas, the West Indies, Mexico, and Central America. STATUS: Common. HABITAT: Usually inhabits open, mature, dry, deciduous forests without thick brush and tangles, preferring areas with an abundance of fallen leaves, logs, and rocks. Occasionally inhabits wet or swampy forests; in the North, inhabits jack pine and spruce forests. NEST: Locates nest in a slight depression in the ground. Uses almost any available vegetation to construct an arched nest resembling a dutch oven, with the entrance hole at or near ground level. FOOD: Gleans invertebrate food from the surface of the litter on the forest floor, including insects, small snails, slugs, myriapods, earthworms, and spiders. REFERENCES: DeGraff et al. 1980, Griscom and Sprunt 1979, Hann 1937, Stenger 1958, Vickery in Farrand 1983c. 423 Northern Waterthrush Seiurus noveboracensis RANGE: Breeds from Alaska and southern Mackenzie across Canada to central Labrador and Newfoundland, south to northwestern Washington and east to central Michigan, northeastern Ohio, southeastern West Virginia, Pennsylvania, New York, and Massachusetts. Winters mostly from Mexico to South America and the West Indies. STATUS: Locally common. HABITAT: Generally inhabits thickets along edges of swamps, ponds, and wooded streams with numerous fallen trees. Prefers woodlands and shrubs around standing water rather than moving streams. SPECIAL HABITAT REQUIREMENTS: Cool, shady, wet, brushy areas with open pools. NEST: Builds nests on the ground among fallen trees, at the base of living trees, in cavities of rotten stumps, and under overhanging banks or cuts. FOOD: Gleans food from moist soil and litter, consuming aquatic and terrestrial insects, small crustaceans, mollusks, and some minnows and worms. REFERENCES: Griscom and Sprunt 1979, Johnsgard 1979, Petersen in Farrand 1983c. 424 .ouisiana Waterthrush eiurus motacilla RANGE: Breeds from eastern Nebraska, central Iowa, and east-central Minnesota to central New York and New England, south to eastern tas central Louisiana, central Georgia, and the Carolines. Winters ,n the West Indies, Mexico, and northern South America. STATUS: Uncommon. HABITAT- Favors bottomland forests with moss-covered logs and rank undergrowth along rapidly moving streams. Also sometimes inhabits shrub-grown bogs or areas near swamp pools or lake edges. SPECIAL HABITAT REQUIREMENTS: Woodlands with flowing water. NEST: Typically builds nest on the ground under roots, or in cavities in steep banks along streams. Also nests in cavities of upturned roots o fallen trees over or near water. FOOD: Searches for food along sandy margins of streams where it eats aquatic and terrestrial insects, spiders, small mollusks, killifishes, and snails. REFERENCES: Eaton 1958, Griscom and Sprunt 1979, Petersen in Farrand 1983c. 425 Kentucky Warbler Oporornis formosus L 4>/ 2 " RANGE: Breeds from southeastern Nebraska, southwestern Wisconsin, southern Michigan, central Ohio, southern Pennsylvania, and southeast¬ ern New York south to eastern Texas, the Gulf Coast, central Georgia, and South Carolina. Casual in southwestern States. Winters from Mexico south to northern South America. STATUS: Common. HABITAT: Inhabits shrubby woodland borders and the understory of damp or shady deciduous woods, favoring moist ravines and bottomlands. Often found near water and at low elevations. NEST: Generally builds nest on the ground among plants at the base of shrubs and trees, or under branches of fallen limbs. Occasionally places nest near the ground in shrubs. Commonly victimized by brown-headed cowbirds. FOOD: Gleans most food from leaves on the ground, but occasionally catches insects from low leaves and branches. Mostly eats spiders and insects. REFERENCES: DeGaris 1936, Griscom and Sprunt 1979, Sykes in Farrand 1983c. 426 Connecticut Warbler Oporornis agilis RANGE: Breeds from east-central British Columbia east across Canada to west-central Quebec, and south to southern Manitoba, northern Minnesota, northern Wisconsin, central Michigan, and south-central Ontario. Winters in South America. STATUS: Uncommon and local. HABITAT: Generally inhabits cold, damp black spruce and tamarack bogs, and prefers areas with scattered trees and grassy openings. At the extremes of the breeding range, inhabits well-drained ridges or poplar and aspen woods. NEST: Conceals nest in a mound of moss or beside a clump of dry grass on or near the ground. Usually nests in open forests with widely spaced trees such as aspen and balsam. FOOD: Mostly eats spiders gleaned from the ground or low branches, but also eats insects, their larvae and eggs. REFERENCES: Griscom and Sprunt 1979, Walkinshaw and Dyer 1961, Harrison 1975. 427 Mourning Warbler RANGE: Breeds east of the Rocky Mountains across Canada and the northern United States from northeastern British Columbia to Newfound¬ land south to North Dakota and central New England through the Appalachian Mountains to Virginia. Winters in Central America and northwestern South America. STATUS: Locally common to uncommon. HABITAT: Inhabits shrubby second-growth, dense undergrowth in open woods, shrubby margins of lowland swamps or bogs, and forest clearings or burned areas that have brambles, shrubs, and saplings May occur in partially open coniferous and deciduous woodlands with herb and shrub understories. SPECIAL HABITAT REQUIREMENTS: Extensive stands of saplings or dense shrubs. NEST: Conceals nests in dense herbaceous or shrubby vegetation on or near the ground. Tends to nest in edges along woodland or clearing edges, logging trails, or at the edges of bogs and marshes. FOOD: Generally gleans beetles, lepidopterans, and spiders from the ground or low shrubs. (Detailed food habits have not been studied.) REFERENCES: Cox 1960, DeGraff et al. 1980, DeSante in Farrand 1983c, Griscom and Sprunt 1979. 428 MacGillivray’s Warbler Dporornis tolmiei RANGE: Breeds from southeastern Alaska, southwestern Yukon, northern British Columbia, southern Alberta, northwestern Saskatchewan, and southwestern South Dakota south, primarily in the mountains, to south¬ ern California, central Arizona, and southern New Mexico. Winters from northern Mexico to Panama. STATUS: Common to uncommon. HABITAT: Prefers early successional stages of cutover or burned wood¬ lands or low shrubby habitats. Also inhabits low vegetation such as blackberry, salmonberry, cherry, currant, serviceberry, snowberry, poison oak, ninebark, spirea, and riparian willow and alder. SPECIAL HABITAT REQUIREMENTS: Low vegetation. NEST: Prefers dense, moist, brushy habitat, or areas with tall weeds or ferns for nesting. Builds nest 2 to 5 feet above the ground, attached to several stalks of plants. FOOD: Forages close to the ground in dense thickets, where it gleans insects from the vegetation. (No comprehensive food studies have been made.) REFERENCES: DeSante in Farrand 1983c, Griscom and Sprunt 1979, Terres 1980. 429 Common Yellowthroat Geothlypis trichas RANGE: Breeds from southeastern Alaska to northern Alberta and Newfoundland south to northern Baja California, Mexico, and southern Texas, the Gulf Coast and southern Florida. Winters along the Pacific Coast, from northern California across southern Arizona, central Texas, and southern Arkansas to the Gulf States, and along the Atlantic Coast from New Jersey, Virginia, and Delaware to Florida; also in the Bahamas, West Indies, Mexico, and Central America. STATUS: Common to abundant. HABITAT: Typically inhabits areas with a mixture of dense, lush herba¬ ceous vegetation with small woody plants (mainly shrubs and small trees), in damp or wet situations. Occasionally found in dry thickets or dense undergrowth in open woodlands. SPECIAL HABITAT REQUIREMENTS: Dense growth of low vegetation. NEST: Builds a bulky cup nest of grass, leaves, and bark, well hidden on the ground in a grass tussock or similar vegetation. Occasionally locates nest in shrubs or a tangle of briars up to 3 feet above the ground. FOOD: Gleans insects and spiders from leaves of shrubs, grasses, and forbs. REFERENCES: Griscom and Sprunt 1979, Hofslund 1959, Low and Mansell 1983, Stewart 1953, Terres 1980. 430 Hooded Warbler Wilsonia citrina RANGE: Breeds from southeastern Iowa, northern Illinois, and extreme southern Michigan and Ontario, southern New York, and New England south to eastern Texas, the Gulf of Mexico, and northern Florida. Winters from Mexico to Panama. STATUS: Common. HABITAT: Generally inhabits moist, forested regions of mixed hardwoods of beech, maple, hickory, and oak with dense undergrowth. In the Southeast, also inhabits cypress-gum swamplands. SPECIAL HABITAT REQUIREMENTS: Low, dense, deciduous woody vegetation. NEST: Builds a cuplike nest, usually in a fork of saplings, shrubs, or in herbaceous vegetation, less than 5 feet above the ground. FOOD: Primarily eats insects, and some spiders. Is an expert “flycatcher.” REFERENCES: Armistead in Farrand 1983c, Bent 1953b, Griscom and Sprunt 1979, Odum 1931. 431 Wilson’s Warbler Wilsonia pusilla RANGE: Breeds from northern Alaska, northern Yukon, northern Ontario, southeastern Labrador, and Newfoundland south to southern California, central Nevada, northern Utah, northern New Mexico, central Ontario, northern New England, and Nova Scotia. Winters from southern California and southern Texas to Panama. STATUS: Common. HABITAT: Prefers wet clearings in early stages of regeneration. Also inhabits peat or laurel bogs with scattered young or dwarf spruces and tamaracks, and riparian willow and alder thickets. SPECIAL HABITAT REQUIREMENTS: Shrubby vegetation. NEST: Generally nests on the ground, sometimes in loose colonies. Usually builds nest at the base of a small tree or shrub, often well concealed in a grass hummock. Occasionally, places nest above the ground in low, dense tangles of vegetation. FOOD: Mostly eats insects (about 93 percent of diet) gleaned from the ground and twigs or caught by flycatching. Also eats some spiders and fruit pulp. REFERENCES: Beal 1907, Bent 1953b, Griscom and Sprunt 1979, Petersen in Farrand 1983c, Stewart 1973. 432 Canada Warbler Wilsonia canadensis L 4 3 A" RANGE: Breeds from central Alberta east to southern Quebec and Nova Scotia south to southern Manitoba, central Minnesota, central Michigan, and through the Appalachian Mountains to northern Georgia. Winters in South America. STATUS: Locally common. HABITAT: Inhabits a variety of vegetative types from lowlands to uplands and coniferous to deciduous. Favors shrubby undergrowth in cool, moist, mature woodlands, streamside thickets, and weedy ravines. In the southern highlands, lives in rhododendron thickets. NEST: Builds nest on or near the ground on mossy logs or stumps, in cavities in banks, among roots of fallen trees, or in mossy hummocks. FOOD: Mostly eats insects and spiders caught by flycatching or gleaned from the ground. REFERENCES: Bent 1953b, Griscom and Sprunt 1979, Krause 1965, Petersen in Farrand 1983. 433 Red-faced Warbler Cardellina rubrifrons RANGE: Breeds from central Arizona and southwestern New Mexico south into Mexico to western Durango. Winters in the highlands of Mexico to El Salvador and western Honduras. STATUS: Locally common. HABITAT: Prefers Douglas-fir and Engelmann spruce forests at elevations of 6,500 to 9,000 feet, but also inhabits ponderosa pine, oak, aspen, and riparian stands and seems to favor southern exposures. NEST: Places nest nearly always on the ground, concealed beneath or beside a sheltering log, rock, sapling, or tuft of grass, usually on a well- drained bank or hillside. FOOD: Searches for food through the outer branches of conifer trees and flycatches. Primarily eats insects. (Food habits have not been studied.) REFERENCES: Griscom and Sprunt 1979, Scott and Gottfried 1983. 434 Painted Redstart Vlyioborus pictus RANGE: Breeds from northwestern and central Arizona, southwestern ''Jew Mexico, and western Texas south through the mountains of Central America to Nicaragua; casually in southern California. Winters from northwestern Mexico south through the breeding range. STATUS: Common. HABITAT: Mainly inhabits timbered desert mountain canyons, gulches, and rugged slopes in coniferous and deciduous woodlands, generally near water. Especially favors dense thickets and oaks in secluded nanyons near streams. MEST: Nearly always places nest on the ground under a rock, tree root, nr grass tuft that provides overhead shelter, and usually on a sloping nank or rocky canyon wall near water. -OOD: Gleans its food from leaves, tree trunks, and branches. Also novers while picking insects from tree foliage, and hawks over water. Mostly eats insects. (Food habits have not been reported, however.) REFERENCES: Bent 1953b, Griscom and Sprunt 1979, Marshall and 3alda 1974, Oberholser 1974b. 435 Yellow-breasted Chat Icteria virens L 6V4" RANGE: Breeds from southern British Columbia, North Dakota, souther Minnesota, southern Ontario, Vermont, and New Hampshire south to south-central Baja California, the Gulf Coast, north-central Florida, and Mexico. Winters from southern Texas and southern Florida south throug Central America and western Panama. STATUS: Common. HABITAT: Favors ravine or streamside thickets of vines, briars, small trees, and tall shrubs. Also inhabits forest edges, hedgerows, overgrowr pastures, scrub country, and early successional stages of forest regeneration. SPECIAL HABITAT REQUIREMENTS: Dense shrubs and vines with scattered young trees. NEST: Usually builds nest 2 to 8 feet above the ground in dense small bushes, vines, or briars. May sometimes nest in groups or colonies, bul maintains separate territories. FOOD: Primarily eats insects gleaned from foliage and shrub stems; also eats some berries and fruits. REFERENCES: Bent 1953b, Dennis 1958, Griscom and Sprunt 1979, Petrides 1938, Thompson and Nolan 1973. 436 Olive Warbler RANGE: Breeds from central and southeastern Arizona and southwestern New Mexico through the highlands of Mexico to north-central Nicaragua. Winters throughout the breeding range, except in Arizona and New Mexico, where it moves southward. STATUS: Fairly common. HABITAT: Generally found near the summits of mountains in the South¬ west above 8,000 feet in mixed pine-fir forests; usually observed near the tops of coniferous trees. NEST: Builds a cup-shaped nest, usually placed on the limb of a conifer tree limb near the end, sometimes hidden by pine needles or a cluster of mistletoe; usually high (30 to 80 feet) above the ground. FOOD: Spends considerable time creeping over the branches and twigs of pines searching for insects. (No studies on food habits have been reported.) REFERENCES: Bent 1953a, Griscom and Sprunt 1979. 437 Hepatic Tanager Piranga flava RANGE: Breeds from southeastern California and northwestern Arizona through New Mexico and the Trans-Pecos region of Texas, south through the highlands of Mexico and Central America to central Argentina. Winters from northern Mexico through the breeding range; casually in southern California and southern Arizona in winter. STATUS: Fairly common. HABITAT: Generally favors dense pine and pine-oak woodlands between 5,000 and 7,500 feet in elevation, but also inhabits the more monotypic pine, oak, and pinyon-juniper woodlands near streams. NEST: Builds a flat, saucer-shaped nest, usually in a fork near the end of a horizontal tree branch, 15 to 50 feet above the ground. FOOD: Gleans insects from branches and leaves of oaks and pines. It occasionally hawks for flying insects, and during the summer eats some fruits. (No studies of food habits have been reported.) REFERENCES: Bent 1958, Phillips et al. 1964, Terrill in Farrand 1983c. 438 Summer Tanager Piranga rubra RANGE: Breeds from southeastern California and southern Nevada to central Oklahoma, and from southeastern Nebraska to New Jersey south to the Gulf Coast and northern Mexico. Winters mainly from Mexico to Bolivia; rare winter visitor in southern temperate areas. STATUS: Common. HABITAT: Generally inhabits dry, open woodlands of oaks, pines, and hickories in the Southeast; but only rich bottomland forests at the northern edge of its range. Inhabits low-elevation willows and cottonwoods, and streamside vegetation in canyons in the Southwest. NEST: Builds a flimsy, flat, shallow cup nest on a horizontal limb (often oak) 10 to 35 feet above the ground. FOOD: Eats many bees and wasps, and the larvae from wasp nests. Catches insects in the air and also eats some small fruits. (Food habits have not been thoroughly studied.) REFERENCES: Bent 1958, Fitch and Fitch 1955, Forbush and May 1955, Johnsgard 1979, Potter 1973, Terres 1980. 439 Scarlet Tanager Piranga olivacea cf RANGE: Breeds from southern Manitoba, western Ontario, southern Quebec, and New Brunswick south to eastern North Dakota, central Nebraska, southern Kansas, eastern Oklahoma, central Arkansas, northern Alabama, and northern Georgia; casually in the West. Winters in South America. STATUS: Common. HABITAT: Generally inhabits mature deciduous and mixed deciduous- coniferous woodlands, roadside shade trees, wooded parks, and large shade trees of suburbs. In the Great Plains, primarily inhabits mature hardwood forests of river valleys, hillsides, and valleys. SPECIAL HABITAT REQUIREMENTS: Mature deciduous or mixed woodlands. NEST: Builds a shallow, saucer-shaped nest, usually well out on a horizontal limb of a large tree, usually in a leaf cluster or in a position where it is shaded from above. FOOD: Gleans food from tree tops, shrubs, or the ground. Eats a great variety of insects, slugs, snails, worms, spiders, and millipedes; also some wild fruits. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Terres 1980, Vickery in Farrand 1983c. 440 i/Vestern Tanager D iranga ludoviciana RANGE: Breeds from southeastern Alaska, northern British Columbia, southern Mackenzie, northern Alberta, and central Saskatchewan south to northern Baja California, southern Nevada, southwestern Utah, central and southeastern Arizona, southern New Mexico, and western Texas, and east to eastern Montana, western South Dakota, northwestern Nebraska, central Colorado, and central New Mexico. Winters from Baja California and central Mexico south to Central America. STATUS: Common. HABITAT: Generally inhabits relatively open, mature coniferous forests up to 10,000 feet in elevation. Breeds less frequently in mixed forests and in deciduous forests in the mountains, along rivers, or in gulches and canyons at lower elevations. NEST: Builds a shallow, compact, saucer-shaped nest, saddled in a fork of a horizontal branch well out from the trunk. Usually locates nest in a coniferous tree. FOOD: Consumes insects gleaned from foliage or branches or caught while hawking (about 82 percent of the diet) and fruits (about 18 percent). REFERENCES: Beal 1907 in Bent 1958, Johnsgard 1979, Harrison 1979. 441 Northern Cardina L 7>A" Cardinalis cardinal!; RANGE: Resident from southeastern South Dakota, central Minnesota, northern Wisconsin, and southern Ontario to Massachusetts and Nova Scotia, south to the Gulf Coast and southern Florida. Local in southwest¬ ern Texas and New Mexico, southern Arizona, and southeastern California south throughout the Mexican lowlands. STATUS: Common in most of the eastern range; local in the Southwest. HABITAT: Inhabits forest edges or brushy forest openings, parks, and residential areas with shrubs and low trees, second-growth woods, and open swamps. In the Southwest, inhabits brushy habitats of washes, riparian and residential areas, and the denser desert thorn-scrub. SPECIAL HABITAT REQUIREMENTS: Dense forest understory or dense shrub habitat. NEST: Builds a loose nest in dense shrubbery, small deciduous or coniferous trees, thickets, vines, briar tangles, or mesquite trees, generally less than 10 feet, but may be up to 20 feet, above the ground. FOOD: Primarily eats vegetable matter gleaned from the ground, including grains, miscellaneous weed seeds, and wild fruits. Also eats smaller amounts of insects. REFERENCES: Forbush and May 1955, Harrison 1979, Johnsgard 1979, McAtee 1908, Terrill in Farrand 1980. 442 Pyrrhuloxia Cardinalis sinuatus L 7V 2 RANGE: Resident from Baja California, south-central and southeastern Arizona, southern New Mexico, and southern and western Texas south to central Mexico. STATUS: Fairly common. HABITAT: Generally lives yearlong in thorny thickets, especially at the edges of mesquite along desert arroyos, in thorny shrubs at lower, wide mouths of mountain canyons, and in thickets along streams. In winter, often wanders from thickets to feed in flocks along roads, fencerows, and borders of fields. NEST: Builds a compact nest, usually low (3 to 8 feet above the ground), in a thorny shrub or tree, or in a clump of mistletoe. FOOD: Gleans most of its food from the ground. During the winter, eats cactus fruits, seeds of grasses, weeds, and mesquite, and catkins of cottonwood. In summer, also eats insects. REFERENCES: Harrison 1979, Oberholser 1974b, Terres 1980. 443 Rose-breasted Grosbeak Pheucticus ludovicianui RANGE: Breeds from southern Mackenzie, across southern Canada to Nova Scotia south to north-central North Dakota and Kansas, central Oklahoma, southern Missouri, central Indiana and Ohio to central New Jersey and south along the Appalachians to northern Georgia. Winters from central Mexico to South America; rare in the Southwest. STATUS: Common. HABITAT: Seems to require a combination of large trees, open areas, and thick shrubs or brush. In summer, inhabits second-growth deciduous or mixed woods, borders of swamps and streams, dense growths of small trees and shrubs along edges of woods and pastures, gardens, and parks of towns and villages. SPECIAL HABITAT REQUIREMENTS: Forest edges with dense brush or thick sapling stands. NEST: Builds a flimsy nest, usually in a fork of a deciduous tree or shrub, about 10 to 15 feet above the ground. Occasionally builds nest in conifer trees. FOOD: Consumes about equal amounts of vegetable and animal matter gleaned from leaves, twigs, or from the ground. Vegetable matter includes weed seeds, fruit, and tree seeds and buds; animal portion is nearly all insects. REFERENCES: Forbush and May 1955, Harrison 1975, McAtee 1908, Terres 1980, Vickery in Farrand 1983c. 444 Black-headed Grosbeak Pheucticus melanocephalus L 7W RANGE: Breeds from southern British Columbia east to southern Saskatchewan and central Kansas south to southern Mexico. Winters mainly in western Mexico from southern Baja California and southern Sonora south to Oaxaca. Rare in the Southwest during winter. STATUS: Common. HABITAT: Primarily inhabits relatively open stands of deciduous forests in uplands or floodplains, but also found in or near orchards, brushy woodlands or chaparral, edges or transitions between grasslands and woodlands, riparian groves or thickets, and parks or suburbs with many trees. NEST: Builds a bulky, open-cup nest 4 to 12 feet above the ground in a fork of a variety of shrubs or small trees. Mostly nests (about 80 percent) in deciduous trees and shrubs. FOOD: Gleans food from leaves, stems, or the ground. Mostly (about 57 percent) eats animal material (mainly spiders and insects); also eats vegetable foods (seeds and fruits.) REFERENCES: Bent 1968a, Bevier in Farrand 1983c, Johnsgard 1979, Terres 1980, Weston 1947. 445 Blue Grosbeak Guiraca caerulea RANGE: Breeds from southern California, southern Nevada, southern Colorado, and Nebraska (also north to south-central North Dakota) and southern Ohio and New Jersey south to the Gulf Coast, and central Florida, through Mexico into Costa Rica. Winters from Mexico south to Panama, and in the Bahamas and Cuba. STATUS: Fairly common, but rare and local in the northeastern part of its range. HABITAT: Inhabits weedy pastures, old fields with saplings, forest edges, streamside thickets, hedgerows, swampy thickets, and willows along irrigation ditches. In the West, inhabits mesquite woods. NEST: Builds a compact, rather deep nest 3 to 8 feet above the ground in a low tree, shrub, tangle of vines, or briars, and typically at the edge of an open area. FOOD: Gleans much of its food from the ground, but also searches for food among leaves and branches of trees and shrubs. Consumes insects, snails, spiders, grain, weed seeds, and fruits. REFERENCES: Forbush and May 1955, Johnsgard 1979, Stabler 1959, Terres 1980. 446 Lazuli Bunting °asserina amoena RANGE: Breeds from southern British Columbia to central North Dakota and northeastern South Dakota south to northwestern Baja California, southern Nevada, central Arizona, central New Mexico, and central Texas. Winters from southern Baja California and southern Arizona south to Guerrero and central Veracruz. STATUS: Common to uncommon. HABITAT: Inhabits a variety of habitats from near sea level to 10,000 feet in the Sierras and 8,000 feet in the Rocky Mountains of Colorado. Generally found in diverse habitats with an abundance of shrubs, low trees, and herbaceous vegetation such as thicketed swales and draws of prairies and dry bushy hillsides, wooded valleys, aspen groves, and shrubby banks of mountain streams. In much of the arid West, found in riparian vegetation. SPECIAL HABITAT REQUIREMENTS: Shrubby vegetation. NEST: Builds a coarsely woven cup nest attached to supporting stalks or a fork of a low shrub or vine tangle, usually 2 to 4 feet above the ground. FOOD: Forages on or near the ground in shrubs and occasionally hawks for insects. Consumes animal material (primarily insects which account for over half of the spring and summer diet) and grass and weed seeds. REFERENCES: Bent 1968a, Grinnell and Miller 1944, Harrison 1979, Johnsgard 1979, Terres 1980. 447 Indigo Bunting Passerina cyanea L 4Vi" RANGE: Breeds from southeastern Saskatchewan and northern Minnesota to southern New Brunswick, south to southern New Mexico, central and southeastern Texas, the Gulf Coast, and central Florida; locally in central Colorado, southwest Utah, central Arizona, and southern California. Winters primarily in southern Mexico, Cuba, and the West Indies south to Panama; also in southern Florida and infrequently in coastal Texas. STATUS: Common in eastern range. HABITAT: Generally associated with edges of woods, old burns, open brushy fields, roadside thickets, and brushy ravines. Tends to be more numerous along streams, and avoids deep woods. SPECIAL HABITAT REQUIREMENTS: Forest edges. NEST: Builds a cup nest 1 to 12 feet above the ground in a crotch of a bush, shrub, or low tree, in a tangle of berry vines, or in canebrakes. Frequently parasitized by brown-headed cowbird. FOOD: Forages on or near the ground in shrubs, and eats a variety of foods, including insects, weed seeds, berries, and fruits. REFERENCES: Bent 1968a, DeGraff et al. 1980, Forbush and May 1955, Harrison 1975. 448 Varied Bunting Passerina versicolor RANGE: Breeds in south-central and southeastern Arizona, southwestern and southeastern New Mexico and southern Texas south to Mexico and Guatemala. Winters from southern Texas south throughout the breeding range. STATUS: Local and uncommon. HABITAT: Inhabits mesquite or thorny shrubs, brushy pastures, dense vegetation with few cottonwoods, foothill canyons, and generally hilly and rocky terrain; tends to avoid heavily wooded areas. SPECIAL HABITAT REQUIREMENTS: Thorny shrubs. NEST: Builds a compact cup nest 2 to 10 feet above the ground in the crotch of a shrub, low tree, or vine tangle. FOOD: Presumably eats insects and seeds similar to the diet of the indigo and lazuli buntings. (No definitive studies have been made.) REFERENCES: Bent 1968a, Harrison 1979, Oberholser 1974b, Terres 1980. 449 Painted Bunting Passerina ciris RANGE: Breeds from southeastern New Mexico and southern Missouri south to southern Alabama and into Mexico, also along Atlantic Coast from southeast North Carolina south to central Florida. Winters from southeastern Texas, central Florida, the Bahamas, and Cuba south through Mexico to Panama. STATUS: Locally common. HABITAT: Inhabits open country with brushy and weedy fields, hedges, edges of woods, roadside shrubs, gullies, thickets along streambanks, shelterbelts, and gardens. NEST: Nests in a variety of deciduous shrubs, small trees, and vines. Attaches shallow cup nest to twigs or other supporting vegetation, 3 to 9 feet above the ground, in bushes, low trees, or vine tangles. Raises two and sometimes three broods each year; susceptible to parasitism by brown-headed cowbirds. FOOD: Mostly eats vegetable matter, which is gleaned from the ground or seed heads of grass, but also some animal matter (insects and a few spiders.) REFERENCES: Bent 1968a, Harrison 1979, Johnsgard 1979, Sykes in Farrand 1983c, Terres 1980. 450 a L 5 3 A" Dickcissel Spiza americana im. s RANGE: Breeds from eastern Montana and southern Canada to Massachusetts, south to central Colorado, southern Texas, and central South Carolina. In the eastern portion of the range, breeds sporadically and irregularly. Winters mostly from Mexico to northern South America. Winters locally (in small numbers) in coastal lowlands from southern New England south to Florida and west to southern Texas. STATUS: Common in the Plains, but rare and local in the East; overall population appears to be declining. HABITAT: Generally inhabits grasslands having tall grasses, forbs, or shrubs but also fields planted to such crops as alfalfa, clover, and timothy. Also frequents abandoned or fallow croplands. SPECIAL HABITAT REQUIREMENTS: Dense herbaceous cover and song perches. NEST: Builds a bulky, cup nest on the ground or attached to forks in shrubs, vines, or low trees. Locates nests in a variety of situations such as marshes, hayfields, abandoned or fallow croplands, roadsides, fencerows, and grasslands. FOOD: Gleans most of its food from the ground; eats mainly vegetable materials — weed seeds and grain, and some insects. REFERENCES: Harmeson 1974, Gross 1921, Overmire 1962, Taber 1947, Tate and Tate 1982, Zimmerman 1982. 451 Olive Sparrow Arremonops rufivirgatus RANGE: Resident in south Texas and south locally to Costa Rica. STATUS: Fairly common locally. HABITAT: Generally inhabits thorny shrub habitats such as mesquite, ebony blackbead, anacua, huisache, and retaima in the Rio Grande delta. Farther north, found in streamside cane, briars, willows, ash, and live oaks. NEST: Builds a domed or nearly round nest 2 to 5 feet above the ground in tangles of shrubbery, pricklypear, or low shrubs, usually near the center of the plant. FOOD: Forages on the ground scratching for insects and various seeds. (Little has been reported on food habits.) REFERENCES: Bent 1968a, Harrison 1979, Oberholser 1974b. 452 Sreen-tailed Towhee J /'p/7o chlorurus RANGE: Breeds from southeastern Washington, southern Idaho, southwestern Montana, northwestern and southeastern Wyoming south through the interior mountains to southern California, southern Nevada, and central Arizona to western Texas. Winters from southern California to western and southern Texas south to central Mexico; casual east of the breeding range in fall and winter. STATUS: Fairly common. HABITAT: Generally inhabits relatively arid and brushy foothills with shrubs such as sagebrush, deerbush, snowbrush, wild rose, spirea, manzanita, waxberry, and chokecherry, from 2,500 feet elevation in California to 10,500 feet in Arizona. Tends to breed at higher elevations in the south than in the north. NEST: Builds a large, loosely constructed and deeply cupped nest on the ground or in low shrubs such as sagebrush, waxberry, and snowbrush, usually less than 2 feet above the ground. FOOD: Forages on the ground, and eats berries, weed seeds, and insects. (Food habits studies are limited.) REFERENCES: Bent 1968a, Flarrison 1979, Johnsgard 1979. 453 Rufous-sided Towhee RANGE: Breeds from southern British Columbia to southern Maine south to southern Baja California, Guatemala, northern Oklahoma, eastern Louisiana, and southern Florida. Winters from southern British Columbia, Utah, Colorado, the southern Great Lakes area, and along the Atlantic Coast south throughout the breeding range. STATUS: Common and widespread. HABITAT: Generally inhabits dense brushy fields and pastures, edges of woods, open woodlands, hedgerows, roadside thickets, and clearings. In the West, inhabits sagebrush, willows, and chaparral above the deserts. SPECIAL HABITAT REQUIREMENTS: Dense brushy cover. NEST: Builds a bulky nest, usually in a depression in the ground but sometimes up to 3 feet above the ground in low shrubs such as coffeeberry or sagebrush. Conceals and protects nest with overhanging bushes, logs, vines, or a clump of grass. FOOD: Mostly eats vegetable material (about 70 percent of diet) gleaned from the ground; most important items are acorns, weed seeds and small fruits. Also eats insects, spiders, and snails. REFERENCES: Baumann 1959, Bent 1968a, Davis 1960. 454 3rown Towhee r ipilo fuscus RANGE: Resident from southwestern Oregon, California (except higher mountains, deserts, and extensive forests), Arizona, southeastern Colorado, and west-central Texas south to central and western Mexico. STATUS: Locally abundant. HABITAT: Generally inhabits chaparral, brushland, woodlands, and open habitats such as lawns and gardens - chaparral and suburban gardens in Oregon and California; cholla cactus and pinyon-juniper in Colorado; the lower mountain canyons in Arizona; and low scrub habitats in New Mexico and Texas. SPECIAL HABITAT REQUIREMENTS: Low shrubs. NEST: Builds a bulky nest ranging from ground level up to 35 feet above the ground (usually 3 to 12 feet) in a variety of shrubs and low trees. Usually places nest in the densest part of the foliage, supported by several branches. FOOD: Forages on the ground for seeds and insects. Feeds insects to nestlings during summer; eats predominately weed seeds in winter. Also eats grain and small fruits. REFERENCES: Bent 1968b, Johnsgard 1979, Terres 1980. 455 Abert’s Towhee Pipilo aberti RANGE: Resident from southeastern California, extreme southeastern Nevada, southwestern Utah, central and southeastern Arizona, and southwestern New Mexico south to northeastern Baja California and northwestern Sonora. STATUS: Common. HABITAT: Lives yearlong in shrubby vegetation such as willow, cotton¬ wood, mesquite, and arrowweed, usually near water courses. Less commonly inhabits baccharis and tamarisk, citrus groves, farms, and urban areas. SPECIAL HABITAT REQUIREMENTS: Riparian shrubs. NEST: Builds a bulky nest, usually near the ground in low shrubs such as mesquite, elder, ash, umbrella trees, or arrowweed; occasionally in trees up to 30 feet above the ground. FOOD: Apparently scratches in leaves on the ground, searching for seeds and insects. (No comprehensive study of food habits has been published.) REFERENCES: Bent 1968b, Phillips et al. 1964, Terres 1980. 456 White-collared Seedeater Sporophila torqueola L 3 3 /4" RANGE: Resident from southern Texas (the Rio Grand Valley north to Webb County), and northern Mexico south to Panama. STATUS: Rare and irregular in Texas. HABITAT: Generally prefers open, grassy places including pastures, roadsides, weedy fields, or marshlands covered with tall grasses in the vicinity of low-growing shrubs such as huisache or retaima. NEST: Builds a delicate cup nest 3 to 5 feet above the ground in crotches of weeds or low shrubs. Frequently nests in abandoned weedy fields, often in giant ragweed. FOOD: Generally forages among low shrubs and weeds to glean seeds of grasses and forbs; also eats some insects. (Little has been reported on its food habits.) REFERENCES: Bent 1968a, Oberholser 1974b. 457 Bachman’s Sparrow Aimophila aestivalis l ST¬ RANGE: Breeds from south-central Missouri and central Indiana to central Maryland, south to eastern Texas, the Gulf Coast, and south- central Florida. Winters from eastern Texas, the Gulf States, and southeastern North Carolina south through the breeding range. STATUS: Uncommon and local, with the population down over most of its range. HABITAT: Generally favors brushy hillsides or wooded borders in the northern part of its range, and open pine stands with grasses and scattered shrubs, oaks, or other hardwoods in more southern areas. Inhabits pine barrens in South Carolina, grassy fields in Mississippi, grasslands with scattered young pines and blackberry thickets in Oklahoma, open pine stands in Florida, and limestone glades in Missouri. SPECIAL HABITAT REQUIREMENTS: Dense herbaceous cover inter¬ spersed with, or bordered by, shrubs and trees. NEST: Constructs a well-concealed nest (open or domed) on the ground, usually under a low bush or against a tussock of grass, and often located at the outer edges of grass clumps in slight depressions, with a clear view in front of the nest. FOOD: Forages on the ground primarily for insects and spiders, but also consumes seeds of a variety of plants. REFERENCES: Hardin et al. 1982, Johnsgard 1979, Tate and Tate 1982, Weston in Bent 1968b. 458 Botteri’s Sparrow Jmophila botterii RANGE: Breeds from southeastern Arizona and extreme southern Texas south to Costa Rica. Winters from northern Mexico south throughout the breeding range. STATUS: Rare and local. HABITAT: Only inhabits areas with dense, tall grass, breeding in open grassland and savannah, especially in areas with scattered brush or shrubs. Favors tall grass habitats with mesquite and catclaw in Arizona; prefers salt-grass with some yucca, pricklypear, and mesquite in the coastal prairies of Texas. SPECIAL HABITAT REQUIREMENTS: Open grassland with scattered shrubs or small trees. NEST: Builds nest on the ground among tall grasses, at the base of a tuft of grass, or sometimes under a projecting mat of grass. FOOD: Primarily eats insects but also spiders and seeds. REFERENCES: Cottam and Knappen 1939, Monson in Bent 1968b, Oberholser 1974b, Terrill in Farrand 1983c. 459 Cassin’s Sparrow Aimophila cassinii RANGE: Breeds from southeastern Arizona, New Mexico, central and northeastern Colorado, southwestern Nebraska, and Kansas south into Mexico and Texas. Singing males may appear sporadically from southern California to South Dakota. Winters from southeastern Arizona and western and south-central Texas into Mexico. STATUS: Common. HABITAT: Prefers open grassland and short-grass plains with a few scattered shrubs or small trees. Also frequents mesquite grasslands if the mesquites are small with open areas throughout but will not usually inhabit areas that are entirely grass unless surrounded by a fence for perching. Occasionally occurs in or near mountainous areas, on grassy slopes with scattered yuccas or small oaks. Favors sandy prairies with scattered sage, yucca, cactus, mesquite, and shinnery oaks in Oklahoma. Apparently can breed where no drinking water is available locally. SPECIAL HABITAT REQUIREMENTS: Short-grass plains with scattered shrubs. NEST: May nest either on the ground, or up to 12 inches above the ground in low bushes or among tangled branches of cacti. Typically places ground nests at the foot of small shrubby plants, concealed in weeds or placed in a tuft of grass. FOOD: Eats mainly insects taken from the ground or the grass. In the winter, eats small seeds of weeds and grasses. REFERENCES: Johnsgard 1979, Williams and LeSassier in Bent 1968b. 460 Rufous-winged Sparrow imophila carpalis STATUS: Locally common. HABITAT: Occurs in rather restricted, isolated colonies in open flat grassy areas with scattered thorn bushes, bunch grasses, mesquite, or cholla. Inhabits desert swales with wide grassy bottoms, leguminous brush, and low trees, washes with sandy bottoms but vegetated slopes; creeks bordered by broad-leaved trees, mesquite, grasses, and weeds, and brushy irrigation ditches. SPECIAL HABITAT REQUIREMENTS: Grassy areas with scattered shrubs that are thorny or dense, preferably both. NEST: Nests in the edges of bushes such as hackberry, paloverde, cholla cacti, or mesquite, usually in a crotch or fork of a branch, 5 to 10 feet above the ground. Also will nest in dense clumps of mistletoe. FOOD: During the nesting season, feeds on a variety of insects caught on the wing or gleaned from plant surfaces. During other seasons, presumably eats grass and weed seeds. REFERENCES: Phillips in Bent 1968b, Phillips et al. 1964, Terres 1980, Terrill in Farrand 1983c. 461 Rufous-crowned Sparrow RANGE: Breeds from central California, southwestern Utah, southeastern Colorado, and central Oklahoma south into Mexico. Winters from north¬ eastern New Mexico, northern Texas, and south-central Oklahoma south throughout the breeding range. STATUS: Locally common. HABITAT: Inhabits dry and desertlike habitats, preferring rocky, brushy, relatively arid hillsides with extensive bare areas. Also in rocky glades on the Great Plains; low ridges and foothills covered with scattered shrubs or trees and grass in Arizona; rocky slopes with large boulders, small cedars, and stunted oaks in Oklahoma; and grassy hillsides with scattered rocks and shrubs, especially sagebrush, and coastal scrub in California. SPECIAL HABITAT REQUIREMENTS: Rocky, arid slopes with scattered brush and grass. NEST: Usually builds nest in a small depression on the ground, often near or under a clump of grass, or at the base of a shrub or small tree. Also locates nests up to 2 feet above ground, wedged among dense vehicle growing branches in shrubs and low trees. FOOD: During the nesting season, feeds on a variety of insects and some spiders caught while foraging on or near the ground. In other seasons, eats some seeds. REFERENCES: Cogswell in Bent 1968b, Johnsgard 1979, Phillips in Bent 1968b, Terres 1980, Terrill in Farrand 1983c, Verner and Boss 1980. 462 American Tree Sparrow ipizella arborea RANGE: Breeds from Alaska and northern Yukon across Canada to lorthem Quebec and Labrador, and south to northwestern British Columbia, northern Saskatchewan, and central Quebec. Winters from southern Canada south to northern California, northern and east-central Arizona, north-central Texas, Arkansas, and North Carolina. STATUS: Common. HABITAT: During the nesting season, inhabits scrub conifers, boggy meadows, and wet, hummocky tundra strewn with boulders and interspersed with willow, birch scrub, and other low shrubbery. Occurs as far north as scrubby growth is found, and in sparse forests just below treeline. In winter, frequents open country, weedy fields, brushy pastures, marshes, fencerows, hedgerows, and thickets. SPECIAL HABITAT REQUIREMENTS: Scrubby trees or bushes for nesting. NEST: Usually constructs nest on or near the ground in a depression, in a tussock of grass, or atop a mossy hummock, placed at the base of a tree or shrub, and concealed by grasses. Occasionally, places nest up to 5 feet above ground in dwarf spruce or willow. FOOD: In summer, primarily eats insects plus some plant material, while in winter, primarily eats seeds of weeds and grasses. REFERENCES: Baumgartner 1937a, 1937b; Bent 1968b; DeGraff et al. 1980; Forbush and May 1955. 463 Chipping Sparrow Spizella passerina RANGE: Breeds from east-central and southeastern Alaska and central Yukon to northern Manitoba, southern Quebec, and southwestern Newfoundland south to southwestern and east-central California, central and eastern Texas, the Gulf Coast, and northwestern Florida through the highlands of Mexico to Nicaragua. Winters from central California, northern Texas, Tennessee, and Maryland south in Mexico throughout the breeding range. STATUS: Common. HABITAT: Inhabits gardens, residential areas, farms, orchards, open coniferous and deciduous woodlands, forest edges and clearings, wooded borders of lakes and rivers, mountain meadows, and grassland habitats with scattered trees. Prefers habitats with trees surrounded by an open area with only herbaceous vegetation and some open ground for foraging. In winter, favors weedy fields and dry scrubland. NEST: Builds nest 1 to 25 feet, but usually 3 to 10 feet, above ground in trees, especially conifers, shrubs, or vines. Generally locates nest near the trunk and top of smaller trees, or lower in the branches and farther from the trunk in larger open-grown trees, usually well concealed. FOOD: Generally forages on the ground in open meadows or lawnlike areas gleaning insects and seeds. REFERENCES: Beal and McAtee 1912, DeGraff et al. 1980. DeSante in Farrand 1983c, Forbush and May 1955, Johnsgard 1979, Johnson in Bent 1968b, Stull in Bent 1968b, Walkinshaw 1944. 464 ^lay-colored Sparrow oizella pallida tANGE: Breeds from eastern British Columbia and west-central and southern Mackenzie east to central Ontario, and south to eastern /Vashington, central Montana, eastern Colorado, northern Iowa, central ind southeastern Michigan, and southwestern Quebec. Winters from central Texas to southern Mexico. STATUS: Locally common. HABITAT: Prefers midwestern mixed-grass prairies with scattered low hickets of shrubs such as wolfberry; will inhabit a variety of dry, unculti- zated shrubby habitats, including grasslands with taller shrubs or small trees, brushy hillsides, overgrown clearings and pastures, parklands, brushy woodland edges, burned-over areas, weedy thickets along roads, swamps, fencerows, railroad tracks and fields, shelterbelts, and other early successional disturbed habitats. SPECIAL HABITAT REQUIREMENTS: Open brushland. NEST: Builds nest either on the ground, well hidden in a tuft of grass at the base of a shrub or near a clump of weeds, or up to 4 1/2 feet above ground in a low shrub or small tree. Commonly uses snowberry, rose¬ bushes, serviceberry, and conifers for nesting. FOOD: Feeds primarily on a wide variety of weed and grass seeds, but will also eat insects in spring and summer, and willow catkins and the buds of elms and other trees in spring. REFERENCES: Forbush and May 1955, Fox 1961, Hussong 1946, Johnsgard 1979, Knapton 1978, Root in Bent 1968b, Salt 1966. 465 Brewer’s Spizella breweri Sparrow RANGE: Breeds from southwestern Yukon and northwestern interior British Columbia to southwestern Saskatchewan south, generally east of the Cascades and coast range, to southern California, central Arizona, central Colorado, and southwestern South Dakota. Winters from southern interior California to central Texas south into Mexico. STATUS: Common. HABITAT: Inhabits open, shrub-dominated habitats; arid sagebrush country in the West and scrub balsam-willow habitats in timberline areas of western Canada, as well as bunchgrass prairie with rabbitbrush, dry, brushy mountain meadows, and pinyon-juniper woodlands. SPECIAL HABITAT REQUIREMENTS: Exposed scrub vegetation from desert regions in the south to timberline in Canada. NEST: Builds nest in shrubs, especially sagebrush, almost always located less than 4 feet above the ground. At timberline, locates nest 6 inches above ground in birch trees, well concealed overhead by interlocking branches. Rarely nests on the ground. FOOD: During winter, primarily eats weed seeds. In spring and summer, also consumes insects and spiders. REFERENCES: Paine in Bent 1968b, Reynolds 1981. 466 ield Sparrow izella pusilla (VNGE: Breeds from northwestern and southeastern Montana and >rthern North Dakota to southwestern Quebec and southern New •unswick south to western Kansas, southern Texas, the Gulf Coast, and >uthern Georgia. Winters from Kansas to Massachusetts, and south to exico, the Gulf Coast, and southern Florida. rATUS: Common. ABITAT: Occurs in a variety of habitats that provide low grassy areas id shrubs or low trees, including old fields and pastures overgrown ith briar thickets or deciduous underbrush, brushy fencerows, cut over ne forests and burned-over woodlands wherever briars and brush have (generated, edges of open, unplowed fields, sagebrush flats, forest Iges, and other similar habitats. PECIAL HABITAT REQUIREMENTS: Abandoned fields or other open -eas with low shrubs or trees. EST: Early in the nesting season, usually builds nest on or near the round in weed clumps or tufts of grass; later in nesting season, builds est as high as 4 feet above the ground in shrubs or small trees, ocates nest in a wide range of plant species — grapevines, cinquefoil, lackberry bushes, boxelders, small oaks, and hickories. OOD: Gleans a variety of seeds of weeds and grasses from the ground iroughout the year; in summer, also eats insects (40 percent of diet.) IEFERENCES: Best 1977, 1978, DeGraff et al. 1980, Forbush and May 955, Fretwell 1968, Johnsgard 1979, Vickery in Farrand 1983c, \/alkinshaw in Bent 1968b. 467 Black-chinned Sparrow Spizella atrogularis RANGE: Breeds from south-central California east to southern Nevada and southwest Utah, south to Arizona, New Mexico, western Texas, and Mexico. Winters from coastal California, southern Arizona, New Mexico, and Texas, south into Baja California and Mexico. STATUS: Uncommon. HABITAT: In desert regions, inhabits tall, dense sagebrush or other brushland areas covered with a variety of plant species. Prefers slopes with rocky outcrops and scattered pinyon or juniper trees. In the Far West, inhabits dry chaparral habitat with a variety of shrubs and scrub oak. SPECIAL HABITAT REQUIREMENTS: Chaparral and sage habitat with rocky outcrops. NEST: Generally builds a compact cup nest of dry grasses, often lined with animal hair, typically placed at the base of a shrub or in the lower portions of sage and shrub, occasionally up to 40 inches above the ground. FOOD: Forages through sage and chaparral habitat, presumably taking a variety of seeds, berries, and soft-bodied insects. (Only limited information is available on its food habits.) REFERENCES: Phillips et al. 1964. 468 Vesper Sparrow Pooecetes gramineus L 5 Vi' RANGE: Breeds from southern Mackenzie and central Saskatchewan to southern Quebec and Nova Scotia, south to eastern and southern California, central New Mexico, Kansas, and North Carolina. Winters from central California, central Texas, southern Illinois, and Connecticut south to Mexico, the Gulf Coast, and central Florida. STATUS: Fairly common. HABITAT: Favors sparsely vegetated dry uplands but also occurs in a variety of habitats throughout its range. In the West, inhabits open grass¬ lands and sagebrush flats, pinyon-juniper associations, open meadows and farmlands, and low grassy areas of alpine and subalpine meadows. In the East, inhabits short-grass meadows, pastures, hayfields, country roadsides, prairie edges, blueberry barrens, coastal beachgrass, and, farther north, forest clearings and burned-over areas. SPECIAL HABITAT REQUIREMENTS: Open areas with short herbaceous vegetation and conspicuous song perches. NEST: Builds nest in a depression on the ground, frequently near small patches of bare ground, where the vegetation is low and sparse, or at the base of a dirt clod, clump of weeds, or tussock of grass, often well concealed by surrounding live or dead vegetation. FOOD: Gleans insects and seeds from the ground and from weeds and grasses; also forages on waste grains. REFERENCES: Berger in Bent 1968b, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, King in Bent 1968b, Vickery in Farrand 1983c. 469 Lark Sparrow Chondestes grammacus RANGE: Breeds from western Oregon and Washington, north into southern Canada, east through the Great Plains to the Missouri River, south throughout the Southwest into Mexico. Winters from central California, southern Arizona, and Texas south to Baja California, throughout Mexico, and in parts of the Gulf Coast up through South Carolina. STATUS: Common throughout most of its range. HABITAT: Generally inhabits open prairies and other open lands. In the spring, frequently found along roadsides with grassy vegetation, but prefers open areas with scattered brush and trees. Also inhabits forest edges, cultivated areas, orchards, fields, and savannahs. SPECIAL HABITAT REQUIREMENTS: Dry fields with scattered bushes or trees or open edge. NEST: Builds nest either on the ground or in low trees or bushes. Breeds in the open but retires to the borders of open woodlands or brushy areas after the young have hatched. FOOD: Eats both soft-bodied insects and seeds. REFERENCES: Forbush and May 1955. 470 3lack-throated Sparrow mphispiza bilineata RANGE: Breeds from southeastern Oregon and northern California, east through the Great Basin, south into Baja California and Mexico. Winters from southern Arizona, New Mexico, and Texas, south into Baja California and Mexico. STATUS: Common in parts of its range. HABITAT: Inhabits thinly grassed pastureland with scattered mesquste, yucca, pricklypear, and cholla cacti. Generally found in dry uplands but extends into the depths of Death Valley. SPECIAL HABITAT REQUIREMENTS: Arid areas with scattered shrubs including cactus, sage, and mesquite. NEST: Generally conceals nest near the ground in small bushes or a variety of cactus species. Usually locates nest about 12 inches from the ground, fastened among forking branches of the low shrubs. Builds nest with small twigs and fibers of sage, frequently lining it with the fur of animals found in the area, such as rabbits. FOOD: Eats a variety of insects as well as seeds that are available where it is breeding and wintering. Selects soft-bodied insects. Survives in areas lacking water by eating green vegetation or insects. REFERENCES: Johnsgard 1979. 471 Sage Sparrow Amphispiza belli RANGE: Breeds from central Washington and eastern Oregon, east through the Great Basin into Wyoming and Colorado, south to southern California, Arizona, and New Mexico. Resident from southern California, into Baja California and parts of Arizona. Winters in parts of Arizona, New Mexico, eastern Texas, and northern Mexico. STATUS: Common in parts of its range. HABITAT: Prefers sage habitat where sagebrush, saltbush, and chaparral are found. Often found in and among the bushes in the Great Basin region and other dry desert areas. SPECIAL HABITAT REQUIREMENTS: Sagebrush and chaparral with scattered bushes. NEST: Usually nests in low shrubs in desert regions, but sometimes in small depressions on the ground. Places nest 6 to 18 inches above the ground, often built into the body of the bush so the foundation is firmly placed. Builds nest of sticks and twigs, often lining it with animal hair and dry grass. FOOD: Eats soft-bodied insects, ants, and spiders, often obtained from the ground or low in bushes. In winter, primarily eats small seeds and other plant materials. REFERENCES: Reynolds 1981. 472 IANGE: Breeds from southern Alberta to southwestern Minnesota, and iouth, east of the Rockies, to eastern New Mexico, the Texas Panhandle, md northwestern Missouri; also locally or sporadically in southern Cali- ornia and Utah to west-central Texas. Winters from southern California o north-central Texas south into Mexico and southern Louisiana. STATUS: Common; on the Blue List because of overall declines in Dopulation. HABITAT: On the western Great Plains, inhabits mixed short-grass Drairie and other areas of predominately low growth, but also areas of taller grasses with scattered shrubs and disturbed grasslands. Also inhabits sagebrush, fenced pastures, cultivated or fallow alfalfa or clover croplands, weedy roadsides, meadows, and areas of relatively barren ground. SPECIAL HABITAT REQUIREMENTS: Open habitats with relatively short, herbaceous vegetation. NEST: Builds nest in a depression in the ground, usually well concealed by grasses or other prairie plants, often located near the base of a plant or plant debris. FOOD: Feeds on the ground, taking primarily insects during the summer, especially grasshoppers. In other seasons, eats seeds of weeds and grasses predominately. REFERENCES: Ballard in Farrand 1983c, Baumgartner in Bent 1968b, Forbush and May 1955, Johnsgard 1979, Tate and Tate 1982. 473 Savannah Sparrow Passerculus sandwichensis L 4 3 A" RANGE: Breeds from Alaska and northern Yukon to northern Labrador and Newfoundland, south in coastal regions to west-central California, and in the interior to central California, northern New Mexico, Nebraska, Kentucky, and New Jersey. Winters from southern British Columbia and southern Nevada to southern Kentucky and, east of the Appalachians, from Massachusetts south into Mexico. Resident in coastal southern California. STATUS: Common throughout its range, although the population is down in the western mountains. HABITAT: Inhabits open wet areas with grass or grasslike vegetation. Occurs in hayfields, pastures, coastal and inland marshes, grassy dunes, wet meadow zones of ponds, lakes, and streams, prairies, open grass¬ lands, bogs, open moist areas of mountain parks and meadows, and tundra. SPECIAL HABITAT REQUIREMENTS: Dense grassy or herbaceous vegetation of moderate height. NEST: Places nest in a natural hollow or scratched-out depression in the ground, among thick herbaceous cover, usually well hidden, not only by the dense cover surrounding the nest but also by overhanging vegetation. FOOD: During summer, mostly eats insects, concentrating primarily on beetles and grasshoppers; also consumes some spiders and snails and gleans a wide variety of seeds. REFERENCES: Baird in Bent 1968b, DeGraff et al. 1980, Elliott in Bent 1968b, Forbush and May 1955, Johnsgard 1979, Potter 1972, Taber in Bent 1968b, Tate and Tate 1982, Welsh 1975. 474 Baird’s Sparrow \mmodramus bairdii L 4 >/ 2 " RANGE: Breeds from southeastern Alberta to southern Manitoba south to central and eastern Montana, southern South Dakota, and west- central Minnesota. Winters from southeastern Arizona to north-central Texas south into Mexico. STATUS: Uncommon. Of special concern on the Blue List as population is down on its breeding range. HABITAT: Favors large areas of prairie grassland with tangles of old and new grasses and patches of shrubs such as snowberry, wolfberry, rose, and willow. Also inhabits ungrazed or lightly grazed mixed-grass prairies, moist meadows, tail-grass prairies associated with wetlands, drier rangelands, fallow and stubble fields, and hayfields. May abandon an area after plowing, burning, mowing, or raking. SPECIAL HABITAT REQUIREMENTS: Relatively undisturbed or reclaimed grassy prairie with scattered shrubs. NEST: Nests on the ground, preferably in tall, dense grass or other dense herbaceous vegetation. Places nest in a hollow of a tuft of grass supported by a shrub, well concealed on the ground by overhanging vegetation, or most commonly, in a natural hollow or a shallow excavated depression, with no overhead concealment. FOOD: Forages on the ground for a variety of seeds throughout the year, but consumes many insects in summer. REFERENCES: Cartwright et al. 1937, Johnsgard 1979, Lane in Bent 1968b, Tate and Tate 1982. 475 Grasshopper Sparrow Ammodramus savannarum L 4'/2” RANGE: Breeds from southern interior British Columbia and southern Alberta to southwestern Quebec and southern Maine south to southern California, central Colorado, northern and south-central Texas, central Georgia, and central North Carolina. Winters from central California (rare) and southern Arizona to Tennessee and North Carolina south to Central America. STATUS: Common, but population is declining from the Dakotas and Nebraska east to New York and Maryland. HABITAT: Prefers prairies in the West and cultivated grasslands, especially those with orchardgrass, alfalfa, red clover, and bush clover, in the East. Inhabits mixed-grass, short-grass, and tail-grass prairies, sage prairies, small grain fields and weedy fallow fields. Avoids fields containing more than 35 percent shrubs, but will occupy grassy habitats with some scattered trees. SPECIAL HABITAT REQUIREMENTS: Continuous tall herbaceous cover and conspicuous song perches. NEST: Builds nest in a slight depression on the ground, usually well hidden at the base of a clump of grass or other vegetation, with vegetation arched over the top. May nest singly or in small colonies. FOOD: Gleans food from the ground. Consumes a diet that is 63 percent insects from fall to spring; also includes spiders, snails, and seeds. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Smith 1963, Bent 1968b, Tate and Tate 1980. 476 Henslow’s Sparrow Ammodramus henslowii L 4W RANGE: Breeds from eastern South Dakota and central Minnesota east to northeastern Massachusetts, and south to central Kansas, northern Kentucky, and east-central North Carolina; also locally in eastern Texas. Winters in coastal states from South Carolina south to Florida and west to Texas. STATUS: Uncommon to rare and local. Of special concern on the Blue List due to continued decline in population. HABITAT: Inhabits meadows and neglected weedy or grassy fields, (often with scattered low bushes) generally preferring situations that are low-lying and damp, but occasionally inhabiting dry or cultivated uplands. Also, inhabits broomsedge fields, old pasturelands, and the drier edges surrounding salt marshes. In winter, frequents grassy openings in pine woods and second-growth woodlands. SPECIAL HABITAT REQUIREMENTS: Habitats with dense herbaceous vegetation, ground litter, an intermediate moisture range, and singina perches. a NEST: Nests singly or in loose colonies. Places nest in a depression on e ground, at or near the base of a thick grass clump with the base of the nest just above the ground, or 6 to 20 inches above the ground attached to vertical stems of herbaceous vegetation. Conceals nest under overhanging vegetation. FOOD: During spring, summer, and fall, primarily eats insects (82 Dercent) and some seeds of weeds and grasses (18 percent) gleaned ^EFER ENCES: Forbush and May 1955, Hyde 1939, Robins and Tate 1982. 1971, Tate 477 Le Conte’s Sparrow Ammodramus leconteii L 4VV RANGE: Breeds * rom east 'C®^ l ^ s ^ r ^^ r ^°[^ J ( ^^na^northe , rn^orth nZ ' e and northern Saskatchewan winters from west-central Kansas, so““ and S ° Uth Car0,ma S ° Uth t0 TeX3S ’ the Gulf Coast, and southeastern Georgia. STATUS: Common. HABITAT: Inhabits temp, ‘Ste surrounded by bog rdte S n'fens m in S S forests. Prefers hummocky alkaline wetlands. SPECIAL HABITAT REQUIREMENTS: Moist habitats with dense herbaceous vegetation. NEST: Builds £ ST S3? 5 passes interwoven among stems, ^orally inbense vegetation beneath tangles of old dead rushes, grasses, or sedges. POOD: Primarily eats insects ^—^^“2^00^^^^ seeds is known about the food habits of this REFERENCES: Forbush and May 1955, Easterla 1962, Johnsgard 1979 Murray 1969, Walkinshaw in Bent 1968b. 478 Sharp-tailed Sparrow Ammodramus caudacutus RANGE: Breeds from east-central British Columbia, southern Mackenzie, and central Manitoba south to south-central Alberta, southeastern South Dakota, and northwestern Minnesota; also around James Bay, in southeastern Quebec, and along the Atlantic Coast from eastern Quebec south to North Carolina. Winters along the Atlantic Coast from New York south to Florida, the Gulf Coast west to Texas, and rarely in coastal California. STATUS: Common along the coast, locally common inland. HABITAT: In coastal areas, prefers short grasses in drier sections of salt or brackish marshes, especially where marshhay cordgrass and seashore saltgrass are present. Inland races inhabit freshwater marshes, marshy zones of prairie lakes and ponds, wet meadows, sloughs, and alkaline, hummocky fens. SPECIAL HABITAT REQUIREMENTS: Well-drained sections of wetland with grassy or other herbaceous vegetation. NEST: In coastal areas, builds nest in higher regions of marshes seldom flooded by tides, salt hay meadows, or the zone between marsh and upland. Elevates and conceals nest just above ground, sometimes attached to upright plant stems or placed in thick clumps of grass. Inland, nests are usually sunken in the ground but occasionally elevated. FOOD: Forages on the banks of pools and creeks, gleaning primarily insects and small aquatic animals, but also seeds of grasses and weeds from the ground and low vegetation. Generally feeds throughout the marsh, although in coastal areas, prefers to forage in grass that is dense and matted. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Hill in Bent 1968b, Johnsgard 1979, Low and Mansell 1983, Murray 1969, Woolfenden 1956. 479 Seaside Sparrow Ammodramus maritimus L 5 V 2 " RANGE: Breeds along the Atlantic Coast from Massachusetts south to northern Florida, and along the Gulf Coast from southern Texas east to western Florida. Winters throughout the breeding range to southern Florida, although most northern populations usually withdraw southward. Resident in southern and east-central Florida. STATUS: Locally common, however the dusky race in east-central Florida is on the verge of extinction, and the Cape S'able race in southern Florida is endangered. HABITAT: Inhabits short-grass tidal marshes and meadows (except for the Cape Sable race, which occurs in a narrow band of fresh and brackish marshes in southern Florida). Prefers rank stands of cordgrass along the borders of tidal creeks with muddy bottoms. SPECIAL HABITAT REQUIREMENTS: Tidal salt marshes. NEST: Builds nest above ground in cordgrass or rushes in wetter portions of salt marshes washed by the tide, on the ground in dense vegetation above the high tide mark, or in shrubs such as marsh-elders. Typically locates well-concealed nest up to a foot above ground. Generally, nests in wetter areas than the sharp-tailed sparrow. FOOD: East mostly insects, crustaceans, and small marine life and some seeds of weeds and grasses. Prefers to forage in areas of open mud and smooth cordgrass along the edge of marshes, gleaning its prey from the ground and surrounding vegetation. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Petersen in Farrand 1983c, Sprunt in Bent 1968b, Stimson in Bent 1968b, Woolfenden 1956. 480 RANGE: Breeds from Alaska and the Yukon to northern Quebec and northern Labrador, south to northwestern Washington, in the western mountains to southern California and central Colorado, and east of the Rockies, across central Canada to southern Quebec. Winters from southern Alaska and southern British Columbia south through the Pacific States, and from central Arizona, Kansas and New Brunswick south to Mexico and central Florida. STATUS: Common. HABITAT: Fairly nonspecific in its requirements, needing only dense, shrubby undergrowth. Inhabits a wide variety of habitats throughout its range, including the undergrowth of deciduous or coniferous forests, brushy woodland edges, woodland thickets, chaparral, burns, cut over areas, scrub, riparian woodlands, willow thickets, and montane coniferous scrub. SPECIAL HABITAT REQUIREMENTS: Dense shrubby undergrowth. NEST: Commonly locates nest on the ground, well-concealed by surrounding tangles of vegetation, or in a bush or tree, typically less than 6 feet, but up to 20 feet above the ground. Prefers conifers for nesting. FOOD: Feeds primarily on insects in summer and on seeds of weeds and some grasses in winter. Forages on the ground, scratching in leaf litter under shrubs and in weed patches. REFERENCES: Austin in Bent 1968c, Forbush and May 1955, Terrill in Bent 1968c, Verner and Boss 1980. 481 Song Sparrow Melospiza melodia RANGE: Breeds from Alaska across Canada, south of Hudson Bay, to Newfoundland and south across the northern part of the United States, along the Pacific Coast, and into Mexico. Resident throughout the northern part of the United States and Pacific Coast. Winters coastally from Alaska and Newfoundland south throughout the southern half of the breeding range. STATUS: Common throughout the eastern United States and northern parts of its range. Locally common in the West. HABITAT: Inhabits moist areas with low irregular plants, or deep grassy and brushy vegetation. Particularly common along waterways, seacoasts with marshes of cattails or bulrushes, and along forest edges adjacent to bogs and openings. Also inhabits fencerows, thickets, hedgerows, and gardens. SPECIAL HABITAT REQUIREMENTS: Moist areas with brushy vegetation. NEST: Usually builds nest on the ground, concealed under a tuft of grass or bush, or even a brushpile, but sometimes in shrubs and small trees up to 12 feet above ground, and in tree cavities. FOOD: Eats a wide variety of both vegetable and animal materials. Gleans insects from the ground, leaves, and branches, mostly during the breeding season. REFERENCES: Nice 1937, 1943, Forbush and May 1955. 482 Lincoln’s Sparrow Melospiza lincolnii L 4 3 /4" RANGE: Breeds from western central Alaska across most of Canada, south along the Pacific Coast and the Rocky Mountains in southern California and northern New Mexico, and into the northern Lake States and northern New England. Winters from southern California, southern Arizona, Texas, and New Mexico south throughout Mexico to Costa Rica. Migrates throughout continental North America between its breeding and wintering ranges. STATUS: Common. HABITAT: Prefers bogs, wet meadows, and riparian thickets. Also inhabits hedgerows, fencerows, and the understory of open woodlands, as well as forest edges, clearings, and shrubby areas. SPECIAL HABITAT REQUIREMENTS: Thickets along the edge of fields, waterways, or in wet meadows. NEST: Usually places nest on the ground in a shallow depression. Builds a rather frail structure of leaves, moss, and some grasses. FOOD: Eats both animal and vegetable materials. During breeding season, mostly eats animal material, including a variety of insects, spiders, and millipedes. During winter, mostly eats grain and grass seed. REFERENCES: Forbush and May 1955. 483 Swamp Sparrow Melospiza georgiana RANGE: Breeds from west-central and southern Mackenzie and northern Manitoba across to southern Labrador, and south to northeastern and east-central British Columbia, the Dakotas, northern Illinois, and Maryland. Winters from eastern Nebraska through the Great Lakes region to Massachusetts, south to Texas, the Gulf Coast, and Florida, and west across New Mexico to southeastern Arizona. STATUS: Common. HABITAT: Wetlands with bushes, rank marsh grasses, sedges, and reeds are characteristic habitat. Inhabits brushy wet meadows, sloughs, bogs, swamps, freshwater marshes, along swampy shorelines of lakes or streams, and rarely in coastal brackish meadows. Avoids heavily wooded wetlands. In winter, frequents springs, seeps, and open brooks that have brushy cover nearby. SPECIAL HABITAT REQUIREMENTS: Swampy wetlands with rank emergent vegetation. NEST: Often builds nest among cattail stalks, upon clumps of bent over vegetation, on sedge tussocks, or in bushes, frequently directly over water that may be 2 feet or more deep. Usually places nest about 12 inches above ground or water, preferably in areas with mixed vegetation rather than in pure cattails. FOOD: Feeds mainly on insects in spring and early summer, and on the seeds of marsh plants in late summer and fall. Gleans food while wading in shallow water or from surrounding vegetation. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Low and Mansell 1983, Martin et al. 1951, Petersen in Farrand 1983, Wetherbee in Bent 1968c. 484 White-throated Sparrow Zonotrichia albicollis RANGE: Breeds from southeastern Yukon and west-central and southern Mackenzie to southern Labrador and Newfoundland south to central interior British Columbia, north-central North Dakota, northern Wisconsin, and northern New Jersey. Winters from southeastern Iowa and southern Wisconsin east to Massachusetts, south to Mexico, the Gulf Coast, and Florida, and west across Texas to California. STATUS: Abundant. HABITAT: Inhabits coniferous and northern deciduous forests, favoring semiopen wooded areas with dense undergrowth or brush, including brushy clearings, cutover woodlands, second-growth, forest edges, borders of swamps and bogs, and other shrubby growth. Seldom found far from dense cover. SPECIAL HABITAT REQUIREMENTS: Open woodlands with dense cover in the form of dense woody undergrowth, thickets, or brush. NEST: Usually builds nest on the ground at the edge of a clearing in areas with small trees, clumps of shrubs, and extensive ground cover of herbs, grasses, and often blueberries. Usually locates nest near a large object such as a tree, stump, or log that possibly serves as a lookout perch; conceals nest by surrounding ground vegetation. Occasionally nests above ground in dense bushes, roots of upturned stumps, or in brush heaps. FOOD: Feeds primarily on the seeds of grasses and weeds, and on wild fruits, but consumes a considerable quantity of insects when available. Forages largely on the ground, scratching in leaf litter or gleaning weeds and grasses for its food. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Lowther and Falls in Bent 1968c. 485 Golden-crowned Sparrow Zonotrichia atricapilla RANGE: Breeds from western and north-central Alaska along the coastal region, south-central Yukon, southern British Columbia, and into the extreme northern part of Washington. Winters from southern Alaska and southern British Columbia, mostly west of the Cascades, to northern Baja California. Also occurs in southeastern California and parts of Arizona, Utah, Colorado, and New Mexico. STATUS: Common. HABITAT: Generally inhabits high-elevation thickets and shrubs, and often dwarf conifers found in brushy canyons. In winter, found in dense brush thickets and chaparral, and gardens along the West Coast. SPECIAL HABITAT REQUIREMENTS: Openings in high-elevation coniferous forests, particularly spruce forests. NEST: Builds a nest of bulky material, such as ferns, leaves, and dry grass, on the ground or above ground on horizontal branches of small trees, usually well concealed. FOOD: Eats mainly seeds, shoots from plants, buds, and flowers of grasses and forbs. Also gleans small insects from trees, branches, shrubs, and grass. REFERENCES: Verner and Boss 1980. 486 White-crowned Sparrow RANGE: Breeds throughout Alaska, the northern regions of Canada surrounding the Arctic Ocean-Hudson Bay region, east to the Atlantic, south through the Yukon and British Columbia, through the Rocky Mountains, west to the Pacific Coast and the Sierra Nevadas. Winters throughout most of the United States, except Florida and the northern Great Plains, and in Mexico. STATUS: Abundant. HABITAT: Frequents valleys, brushy hillsides, roadside vegetation, and cultivated fields. In arctic regions, inhabits open stunted tree growth and brushlands. Uses edge habitats and brush piles during winter. SPECIAL HABITAT REQUIREMENTS: Thickets, hedgerows, or edge. NEST: Often builds nest consisting of grassy materials, mosses, and lichens, lined with rootlets or animal hairs, placing it on the ground in a moss or lichen bed, in grassy areas, but sometimes on the lower branches of shrubbery. Most nests are well concealed and difficult to locate. FOOD: Obtains both plant and animal foods by scratching in the soil. Eats the fruiting bodies of mosses, as well as a variety of seeds, and capitalizes on outbreaks of insects. REFERENCES: Beal and McAtee 1912, Forbush and May 1955. 487 Harris’ Sparrow Zonotrichia querula southern Keewatin south to northeastern Saskatchewan and northern Manitoba. Winters primarily along the eastern edge of the Great Plains, although it may be found from southeastern Alaska and southern British Columbia to northern Colorado and central Iowa, and south to California, Texas, and western Tennessee. STATUS: Fairly common. HABITAT: Breeds in the Canadian subarctic, in open clearings, woodland edges, and brushy margins of burned-over areas bordered by spruce forests. During winter, inhabits brushy habitats, preferring woodlot borders and hedgerows. NEST: Builds nest in the ground, usually in mossy hummocks surrounded by water near stunted spruce trees, often under a low shrub on a sheltered southern exposure. May also locate nest in dry clearings under small trees within 100 feet of a lake. FOOD: At all seasons, chiefly eats seeds of weeds, grasses, and wild fruits, but includes insects in diet, especially in summer. Primarily feeds on the ground. REFERENCES: Baumgartner in Bent 1968c, Forbush and May 1955, Kaufman in Farrand 1983c, Semple and Sutton 1932. 488 RANGE: Breeds from Alaska and central Yukon to Labrador and Newfoundland, south to central coastal California, in the mountains to eastern California, central Arizona, and western Texas, southern Alberta, northern and east-central Minnesota, central Michigan, southern New England, and in the Appalachian Mountains to northern Georgia and northwestern South Carolina: also in the Black Hills. Winters from central and south coastal Alaska, coastal British Columbia and across southern Canada south to Mexico, the Gulf Coast, and northern Florida. STATUS: Common to abundant. HABITAT: Occurs from sea level to timberline in a variety of wooded habitats that have openings with dense herbaceous ground cover, including coniferous and deciduous forests, forest edges, woodland clearings, stream borders, open woodlands, brushy cover bordering mountain meadows, and old burns. Avoids deep forest interiors in favor of woodland edges and openings. In winter, prefers weedy fields but also inhabits open woodlands, hedgerows, suburbs, and farmyards. SPECIAL HABITAT REQUIREMENTS: Openings in wooded habitats covered with dense herbaceous vegetation such as grasses or forbs. NEST: Commonly builds nest on the ground near the edge of openings in wooded areas or in a slight depression, usually well concealed under weeds, grasses, fallen logs, tree roots, or other overhead shelter. Occasionally may place nest up to 8 feet above ground in a shrub or tree. FOOD: Forages on the ground, picking up seeds (mostly) and insects. REFERENCES: Barrowclogh in Farrand 1983c, Beal and McAtee 1912, DeGraff et al. 1980, Eaton in Bent 1968b, Hostetter 1961, Martin et al. 1951, Phelps in Bent 1968b, Sprunt in Bent 1968b, Thatcher in Bent 1968b, Whitney in Bent 1968b. 489 Yellow-eyed Junco Junco phaeonotus RANGE: Resident from southern Arizona and parts of New Mexico to central Mexico. STATUS: Locally common. HABITAT: Prefers coniferous forests and pines or oaks that are relatively open, usually above 5,000 feet. Often inhabits areas adjacent to scrub, pastures, and fields. SPECIAL HABITAT REQUIREMENTS: Coniferous and pine-oak forests; edge-type habitat appears to be important. NEST: Usually builds cup-shaped nest on the ground under a clump of grass or log, near a flat stone, or sometimes on the ground under a drooping pine limb or in a thick shrub. FOOD: Feeds primarily on seeds and other vegetable material as well as some insects. Forages on the ground, but occasionally removes animal or vegetable material from the limbs of trees. Occasionally eats fruits. REFERENCES: Bent 1968b. 490 VlcCown’s Longspur >alcarius mccownii L 5W' summer RANGE: Breeds from southeastern Alberta east to north-central North Dakota, and south to northeastern Colorado and northwestern Nebraska. Winters from central Arizona, west-central Kansas, and central Oklahoma south into Mexico. STATUS: Uncommon. HABITAT: Inhabits dry shortgrass prairie, plowed and stubble fields, grazed pastures, dry lake beds, and other sparse, bare, dry grounds on the western plains. SPECIAL HABITAT REQUIREMENTS: Open, dry, sparsely vegetated prairie. NEST: Places nest in a shallow depression on the ground, sometimes in a clump of grass or under a shrub, but usually amid sparse prairie vegetation. Typically builds a nest that is open above, covered only by a few blades of grass, although occasionally it may be concealed by overhanging shrub branches. Rarely, places nest above ground in a shrub. FOOD: Forages on the ground, picking up seeds of weeds and grasses, and insects, especially grasshoppers. REFERENCES: DuBois 1937, Johnsgard 1979, Kaufman in Farrand 1983c, Kraus in Bent 1968c, Mickey 1943. 491 Lapland Longspur Calcarius lapponicus L5%" summer 6 winter 6 RANGE: Breeds from western and northern Alaska and northern Yukon across northern Canada, including many islands to Labrador, and south to south-coastal Alaska and southern Keewatin. Winters from coastal southern Alaska and southern British Columbia across the northern United States to Nova Scotia, and south to southeastern California, Colorado, Arkansas, and Maryland; casually farther south. STATUS: Abundant on breeding grounds; local in interior. HABITAT: Breeds in arctic and subarctic tundra, preferring areas of wide, moist tundra covered with grassy tussocks or hummocks and interspersed with ponds and streams. Inhabits moist sedge meadows, swampy flats, marshes, and moist grasslands, preferring sedge tussocks or ridges covered with low willows, or dwarf birch intermixed with heath. Avoids extremely wet areas unless elevated sites are available. In winter, found in stubble fields, barren grounds, beaches, prairies, or open weedy meadows. SPECIAL HABITAT REQUIREMENTS: Moist tundra on relatively level terrain with elevated sites such as tussocks, hummocks, or low ridges, and concealing vegetation. NEST: Usually places nest in a depression in the side of a small hummock or tussock, protected by overhanging grass, sedge, or leafy twigs of shrubs. Uses drier, elevated sites for nesting rather than low, wet, surrounding terrain. FOOD: In summer, eats mostly insects and the seeds of a variety of plants; in winter, feeds mainly on seeds of weeds and grasses. Gleans food from the ground. REFERENCES: DeGraff et al. 1980, Jehl 1968, Williamson in Bent 1968c. 492 Smith’s Longspur 'alcarius pictus summer 6 RANGE: Breeds in east-central Alaska and adjacent British Columbia, and from northern Alaska east across northern Canada to extreme northern Ontario. Migrates through the northern Great Plains. Winters from Kansas and Iowa south to Oklahoma, Texas, and Louisiana. STATUS: Uncommon. HABITAT: Inhabits dry, grassy, hummock areas in tundra, often where there are dry sedge meadows dominated by dwarf birch and scattered black spruce. Prefers areas with a perch site, often using small spruces. Winters in grassy and weedy fields, prairies, and along edges of open areas such as airport runways and roadways. SPECIAL HABITAT REQUIREMENTS: Small isolated trees in tundra areas for perches. NEST: Builds a nest, commonly unprotected from above, in a small depression in a relatively dry, flat hummock, usually on a hummock ridge or at the base of a small tree or shrub. FOOD: Eats a variety of foods during summer; largely plant material, principally seeds (90 percent in the first part of June.) Also eats inverte¬ brates when available, including ants, spiders, and beetles. Commonly eats immature insects in the latter part of the summer. REFERENCES: Jehl 1968. winter 493 Chestnut-collared Longspur Calcarius ornatus RANGE: Breeds from southern Alberta, Saskatchewan, and Manitoba, east of the Rockies to northeastern Colorado and western Kansas. Winters from northern Arizona, central and northern New Mexico, eastern Colorado, and central Kansas south into Mexico. Occurs rarely in California. STATUS: Common. HABITAT: Inhabits shortgrass plains and prairies (cultivation has reducec breeding habitat.) In winter, tends to congregate on cultivated fields, and along edge areas, fencerows, and roadways. SPECIAL HABITAT REQUIREMENTS: Shortgrass prairie. NEST: Builds nest on the ground, usually in a small depression in grass in uncultivated grassland with occasional low ridges and shallow areas. Seems to prefer nest sites in moist areas. In flooded meadows, places nest on an elevated site. FOOD: Feeds primarily on grass seed and some insects, which are gleaned from the ground. During winter, feeds almost entirely on plant material. REFERENCES: Forbush and May 1955. 494 Snow Bunting D lectrophenax nivalis RANGE: Breeds in Arctic tundra from northern Alaska to Prince Patrick and northern Ellesmere islands, south to extreme northwestern British Columbia, east-central Mackenzie, central and southeastern Keewatin, and northern Labrador. Winters from west-central and southern Alaska and southern Canada south to California, Colorado, Missouri, and North Carolina, casually farther south. STATUS: Common. HABITAT: Breeds in rough, rocky Arctic tundra with scattered vegetation. Prefers stony beaches, rocky escarpments, and cliffs, occurring less commonly in grassy tundra. In winter, inhabits open country, along lake shores, beaches, and roadsides, and grassy, weedy, or stubble fields. SPECIAL HABITAT REQUIREMENTS: Rocky areas with sparse vegetation. NEST: Nearly always places nest in a hole or cranny, in a variety of natural and artificial sites, often a foot or more back in narrow rock crevices under loose rocks on the ground. May place nest in a depression or in cracks in the ground if other sites are lacking but rarely exposed on open ground. FOOD: In winter, primarily eats seeds of weeds and grasses gleaned from the tips of plants and the surface of the snow. Adds insects and spiders to the diet during summer. REFERENCES: DeGraff et al. 1980. Forbush and May 1955, Parmelee in Bent 1968c, Sutton and Parmelee 1954b. 495 Bobolink Dolichonyx oryzivorus RANGE: Breeds from southern interior British Columbia across southern Canada and central Ontario south to eastern Oregon, central Colorado, central Illinois, and central New Jersey. Winters in South America. STATUS: Locally common, although numbers are decreasing in the Northeast due to a decline in agriculture; extending range in the West because of irrigation. HABITAT: Prefers large open fields of tall grass, alfalfa, clover, or grain crops, but also inhabits wet meadows, ungrazed to lightly grazed mixed- grass prairies, and fallow fields. During migration, frequents marshes and grain fields. SPECIAL HABITAT REQUIREMENTS: Large expanses of grassland or forb cover. NEST: Builds nest on the ground, usually in a hollow scraped in the ground or in a natural depression, rarely above ground attached to plant stems. Always locates nest in dense stands of tall vegetation such as hay, alfalfa, clover, or thick growths of weeds. FOOD: Prefers to forage in cultivated grain fields, gleaning insects, seeds of weeds and grasses, and waste grain from standing vegetation and on the ground. REFERENCES: Beal 1900, Bent 1958, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979. 496 L 7 1 / HANGE: Breeds from the southern tip of Alaska, and Yukon down to lorthern Washington, across the northern part of the United States and Canada including Idaho, Montana, Wyoming, the Great Lakes, and New England. Resident in the rest of the United States south into Baja Halifonia, Mexico, and Central America. Northern birds migrate southward in winter. Red-winged Blackbird Agelaius phoeniceus STATUS: Abundant. HABITAT: Breeds in marshes and agricultural areas, usually where there ire wetlands and also along the edge of hayfields, old fields, and lastures. Prefers areas with trees nearby and where habitat edges are ibundant; often perches on old erect vegetation. Is extremely territorial, aartitioning territories into areas of several hundred square feet, thus efficiently limiting the numbers that can exploit a resource. Flocks in the winter and moves throughout fields and marshy areas. SPECIAL HABITAT REQUIREMENTS: Standing vegetation along open fields and marshes. NEST: Nests in a deep narrow cup of grass, reeds, and weed rootlets, usually attached to emergent vegetation (particularly cattails) up to 12 feet above ground. Also nests in weeds and brush patches, croplands such as alfalfa and cereal grains, even upland areas of mixed chaparral. FOOD: Consumes a diet consisting of both vegetable and animal material, including a variety of grains and seeds, insects, spiders, mites, and snails. Often descends in large numbers on cultivated fields, eating a great deal of the grain. REFERENCES: Albers 1978, Beal 1900, Case and Hewitt 1963, Lowe and Mansell 1983, Mott et al. 1972, Orians 1961, Payne 1969. 497 Tricolored Blackbird Agelaius tricolor L 7V .2 RANGE: Breeds from southern Oregon east of the coast range south through interior California along the Pacific Coast from central California to northwest Baja California. Resident from northern California south throughout breeding range and adjacent agricultural areas. Some north¬ ern birds are migratory. STATUS: Common. HABITAT: Commonly breeds in freshwater marshes of cattail, tule, bulrush, and sage. Roosts in the strips along marshes between rice fields. Feeds and roosts in dense flocks, ranging from 4 to over 20,000 ir a colony, throughout the year. In winter, moves through marshes, open cultivated lands, and pastures. SPECIAL HABITAT REQUIREMENTS: Cattail or tule marshes. NEST: Builds nest of cattails, sedges, grasses, or other aquatic vegeta¬ tion gathered from the surface or in shallow water, and attached to cattails or twigs in shrubs and blackberry thickets, usually near water. Prefers live emergent vegetation for nesting. FOOD: Gleans food from the ground and low vegetation; eats insects, spiders, and occasionally small tadpoles and snails. In winter, eats rice and a variety of grain crops. REFERENCES: Payne 1969. 498 Eastern Meadowlark Sturnella magna o southern New Brunswick, south through the Eastern United States to rexas, the Gulf Coast, Central America, and Florida, and west to south¬ western South Dakota, central Nebraska and central Arizona. Winters : rom central Arizona, Kansas, central Wisconsin, New England, and Mova Scotia south throughout the breeding range, although casually farther north. STATUS: Common, although there are widespread declines in the Eastern States. HABITAT: Prefers pastures, but also occurs in other grass-dominated habitats such as hayfields, grassy meadows, tallgrass prairies, open fields of corn, alfalfa, and clover, and weedy orchards. Prefers moist meadows and lowlands at the western edge of its range, where distribu¬ tion overlaps that of the western meadowlark. SPECIAL HABITAT REQUIREMENTS: Open grasslands with elevated singing perches such as fences, poles, or lone trees. NEST: Builds nest on the ground in a natural depression or scrape, well concealed by a canopy of vegetation bent over the nest, preferably in cover 10 to 20 inches high. FOOD: Gleans food from the ground and low vegetation. During summer, mainly eats insects; in winter, primarily eats seeds of weeds and grasses, and waste grain, except in southerly states where insects are still available. REFERENCES: Bent 1958, Johnsgard 1979, Lanyon 1957, Roseberry and Klimstra 1970, Tate and Tate 1982. 499 RANGE: Breeds from central British Columbia, north-central Alberta, Saskatchewan, and central Canada, south throughout most of the Western United States. Found in most areas west of the Mississippi. Resident throughout the Pacific, southern Rocky Mountain, and southerr Great Plain States. Winters also in Oklahoma, Texas, and along the coasts of Mexico and Baja California. STATUS: Common. HABITAT: Typically inhabits grasslands, savannahs, cultivated fields, and pastures, preferring open fields with perch sites such as fences, old logs, or dead trees. SPECIAL HABITAT REQUIREMENTS: Open grasslands. NEST: Builds nest in a shallow depression on dry ground in open grassland, often in grass or a small grass tuft, sometimes in rocky areas Usually uses grasses for nest material. FOOD: Gleans food from the ground or low vegetation. Eats both anima and vegetable material; about 70 percent animal material, mainly beetles, but also a variety of other insects and invertebrates; and a variety of grains and seeds. REFERENCES: Lanyon 1957. 500 Yellow-headed Blackbird Xanthocephalus xanthocephalus L 8>/2" RANGE: Breeds from southern and central Canada, throughout the western part of the United States, west of the Mississippi River. Winters from southern California, Arizona, New Mexico, and Texas into Mexico. STATUS: Common. HABITAT: Inhabits freshwater marshes of cattails, bulrushes, and reeds, generally over water. Winters in open cultivated fields, pastures, and marshes. SPECIAL HABITAT REQUIREMENTS: Marshy vegetation. NEST: Generally nests in colonies. Builds a woven basketlike cup nest of marsh vegetation lined with fine grass and attaches it to reeds and cattails 1 to 3 feet above water or sometimes in willows in wet areas. FOOD: Eats both vegetable and animal material gleaned from the ground, mostly vegetative material, which includes seeds and leaves of grasses and forbs, and grain crops. REFERENCES: Beal 1900, Forbush and May 1955, Lowe and Mansell 1983, Willson 1966. 501 Rusty Blackbird Euphagus carolinus L 8" RANGE: Breeds from western and north-central Alaska and northern Yukon to southern Keewatin, northern Quebec and central Labrador south to southwestern and south-coastal Alaska, central interior British Columbia, central Manitoba, northern New England, and northeastern New York. Winters from south-coastal Alaska and southeastern British Columbia to southern Ontario and southern New England south to Texas, the Gulf Coast, and northern Florida, and west to central Colorado. STATUS: Fairly common. HABITAT: Almost always found near water, breeding in boggy spruce woods, along swampy wooded shores of lakes, streams, tree-bordered marshes, beaver ponds, and swamps; and on wooded islands in lakes. Rarely occurs in fields with other blackbirds. In winter, does not stay so close to water, occurring also in open woodland, scrub, pastures, weedy gardens, and cultivated land. SPECIAL HABITAT REQUIREMENTS: Wooded wetlands during the breeding season. NEST: Nests in dense growths of evergreens, especially second-growth spruce or balsam, 2 to 20 feet, but typically less than 10 feet above the ground. Also uses dead trees or clumps of deciduous bushes such as buttonbush or sweetgale along streamsides for nesting. FOOD: Forages by gleaning insects, weed seeds, waste grain, and wild fruit from the ground in pastures, fields, or grassy edges of wetlands, or by wading in shallow water. REFERENCES: Beal 1900, Bent 1958, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Kaufman in Farrand 1983c, Kennard 1920. 502 Brewer’s Blackbird iuphagus cyanocephalus RANGE: Breeds from southwestern and central British Columbia to southern Ontario, south throughout the northern United States; resident in the Pacific Coast, Great Plains, and Rocky Mountain states. Winters from southern British Columbia, east-central Montana, and the northern portions of the Gulf States, south to Baja California, northern Mexico, southern Texas, the Gulf Coast, and Florida. STATUS: Common. HABITAT: Prefers to be near water in habitats such as riparian wood¬ lands, aspen groves, parklands, cultivated lands, and marshes; often found around human habitation. Uses bulrushes and pines for roosting and daytime resting places and displays from the tops of pine trees. In winter, frequents pastures and fields. SPECIAL HABITAT REQUIREMENTS: Marshlike areas. NEST: Nests singly or in loose colonies on the ground or in trees and shrubs 20 to 30 feet above the ground. Places the cup-shaped nest usually at or near the end of a branch. FOOD: Consumes a diet of about 68 percent vegetative and 32 percent animal material gleaned mostly from the ground. Commonly follows plows searching for insects. REFERENCES: Beal 1900, Terres 1980, Williams 1952. 503 Great-tailed Grackle Quiscalus mexicanus RANGE: Resident from southern California, southern Nevada, southeast¬ ern Colorado, Kansas, and southwestern Missouri south through Mexico to South America. STATUS: Common. HABITAT: Inhabits a variety of climatic regions and uses a diversity of plant types for nesting. Basically needs standing water and open ground for foraging; does not occur in forests or at great distances from water. In deserts and prairie country, only occurs near water courses or irri¬ gated agricultural areas where trees are available. SPECIAL HABITAT REQUIREMENTS: Partially open situations with scattered trees and water. NEST: Nests colonially; builds a bulky cup nest of grass, sticks, and seaweed or Spanish moss that is lined with grass and rootlets. Locates nest 2 to 60 feet above ground in a variety of tree species or may build it in reeds over water when trees are unavailable. FOOD: Gleans its food from the ground, occasionally wading in shallow water. Eats grains, berries, mollusks, crustaceans, insects, small fish, young birds, and eggs. REFERENCES: Lowe and Mansell 1983, Pruitt 1975, Selander and Giller 1961. 504 Boat-tailed Grackle Quiscalus maior RANGE: Resident along the Atlantic Coast from New York to Florida, and west along the Gulf Coast to Texas. STATUS: Common. 9 BOAT-TAILED GRACKLE L 16" GREAT-TAILED GRACKLE L 12-16" HABITAT: Prefers open coastal marshes near large bodies of brackish or salt water, but also inhabits large tidal rivers short distances inland, city parks, and farmland. In Florida, inhabits inland freshwater habitats on the peninsula as well as coastal marshes. Seldom occurs more than a few miles inland. SPECIAL HABITAT REQUIREMENTS: Coastal fresh or saltwater wetlands. NEST: Nests in colonies in cattails, bulrushes, marsh-grass, bushes, or trees, from 6 inches to 80 feet above the ground. Usually builds nest in vegetation growing in water or in branches hanging over water Prefers to nest in marsh vegetation, often selecting marsh habitat over trees near water. FOOD: Finds most of its food on the ground by probing in mud and litter or turning over shells or stones. Commomly forages in marshy areas, mudflats, and the margins of ponds, lakes, and streams. Eats insects, fish, frogs, crayfish, mollusks, young birds and eggs, rice, waste grains, and berries. REFERENCES: Beal 1900, Low and Mansell 1983, Mcllhenny 1937, Pruitt 1975, Selander and Giller 1961, Tutor 1962. 505 Common Grackle Quiscalus quiscula L 10-12" RANGE: Breeds from northeastern British Columbia and southern Mackenzie to southwestern Newfoundland, south to central and southeastern Texas, the Gulf Coast and southern Florida, and west to eastern Wyoming, central Colorado, and central and southeastern New Mexico. Winters from Kansas, Iowa, the southern Great Lakes region, New England, and Nova Scotia south to southeastern New Mexico, southern Texas, the Gulf Coast, and Florida. STATUS: Abundant. HABITAT: Prefers open habitat (especially agricultural) with scattered trees or open woodlands, forest edge, and nearby human habitation. Frequents city parks, swamps, brushy or reedy marshes, and cultivated lands, especially in migration. NEST: Builds nest in a variety of trees and often has local preferences for tree species. Nests colonially, building nests 7 to 35 feet above the ground (average 20 feet); most are well concealed in dense masses of foliage. FOOD: Forages for food in cultivated fields, shrubs, and shallow water. Eats nuts, weed seeds, small bulbs, eggs and young of other birds, and fish; can be a pest when large numbers descend on cultivated crops. REFERENCES: Beal 1900, Forbush and May 1955, Jones 1969, Maxwell 1970. 506 Bronzed Cowbird Molothrus aeneus RANGE: Resident from extreme southeastern California, southern Arizona, New Mexico, and Texas south through central Mexico to Panama. STATUS: Locally common. HABITAT: Inhabits mostly open country with occasional tree patches or large tall shrubs. Prefers humid, hot climate, often in areas where cattle are grazed and is common in areas of human habitation. SPECIAL HABITAT REQUIREMENTS: Open areas with scattered trees or shrubs. NEST: Builds no nest; generally lays its eggs in the nests of other birds, preferably those nesting in brush, semi-open to open ranch, farm, and residential areas. FOOD: Eats weed and grass seeds, grain, and insects that are gleaned from the ground. Also eats insects from the skin of livestock. Commonly feeds and roosts in large flocks. REFERENCES: Oberholser 1974b. 507 Brown-headed Cowbird Molothrus ater L 6 Vi" RANGE: Breeds from southeastern Alaska, northern British Columbia, and southern Mackenzie east to southern Quebec and southern Newfoundland, and south to Mexico, the Gulf Coast, and central Florida. Winters from northern California, central Arizona, the Great Lakes region, and New England south to Mexico, the Gulf Coast, and southern Florida. STATUS: Common. HABITAT: Prefers habitats where low or scattered trees are interspersed with grassland vegetation. Originally occupied open grasslands and avoided unbroken forestlands, but due to agriculture, cattle grazing, and deforestation, occupies a much expanded range. Now found in open coniferous and deciduous woodlands, forest edges, brushy thickets, agricultural land, and suburban areas. SPECIAL HABITAT REQUIREMENTS: Habitats with open grassy spaces. NEST: Builds no nest; lays its eggs in the nests of over 100 species of birds, particularly tyrant flycatchers, finches, vireos, and warblers. FOOD: Gleans weed seeds, which form over half of the diet, as well as grass seeds, waste grain, and insects from the ground. Commonly forages in pastures searching for insects stirred up by cattle. REFERENCES: Beal 1900, Bent 1958, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Mayfield 1965. 508 Orchard Oriole RANGE: Breeds from southeastern Saskatchewan, southern Manitoba, and central Minnesota east to northern Massachusetts, south to Mexico, Texas, the Gulf Coast and central Florida, and west to eastern Colorado. Winters from Mexico to South America, casually to southern Texas, rarely in coastal California. STATUS: Locally common, but populations are decreasing from Kansas to Alabama. HABITAT: Prefers orchards and open country with a few scattered trees. Also breeds in residential areas, farmlands, shelterbelts, woodland margins, and lightly wooded river bottoms. At times, it may inhabit marshes and bordering trees. Heavily wooded or dense forests are avoided. SPECIAL HABITAT REQUIREMENTS: Open woodlands or open areas with scattered trees. NEST: Suspends semipendulous nest, well concealed by dense foliage, from a fork or crotch of a variety of trees and shrubs 4 to 70 feet, typically 10 to 20 feet, above the ground. Commonly nests in trees also supporting Eastern kingbird nests. FOOD: Gleans insects, which form over 90 percent of the diet, from leaves of trees and shrubs. Also eats fruits. REFERENCES: Bent 1958, DeGraff et al. 1980, Dennis 1948, Forbush and May 1955, Johnsgard 1979, Tate and Tate 1982. 509 Hooded Oriole Icterus cucullatus RANGE: Breeds from northern coastal and central California, southern Nevada, central Arizona, and western Texas south into northern Mexico. Resident in southern Baja California and throughout mainland Mexico. STATUS: Common. HABITAT: Inhabits palm trees, mesquite, dry shrubs, and some deciduous and riparian woodlands; often found around ranches and towns. NEST: Usually constructs a large, hanging nest with a variety of grasses, Spanish moss, thin branches, as well as dry vegetables, hair, and other local materials woven together, and suspends it from the limbs of trees or cacti. FOOD: Eats a variety of insects, along with flower nectar, fruit, and other plant materials. REFERENCES: Phillips et al. 1964, Terres 1980. 510 Altamira Oriole Icterus gularis RANGE: Resident from the lower Rio Grande Valley in extreme southern Texas south through Mexico to Central America. STATUS: Rare. HABITAT: Inhabits open woodlands, trees along fields and streams, scattered groves in pastures, and hillsides. NEST: Fastens conspicous pensile pouch nest, 1 to 2 feet long, to the ends of slender, strong, flexible terminal twigs, 12 to 35 feet above the ground. Usually places nest in eborty blackbead, mesquite, or willow. FOOD: Forages among the leaves of trees for insects and fruits; also eats caterpillars, spiders, small figs, and berries. REFERENCES: Bent 1958, Oberholser 1974b, Sutton and Pettingill 1943, Terres 1980. 511 Audubon’s Oriole Icterus graduacauda RANGE: Resident in southern Texas and Mexico. STATUS: Uncommon. HABITAT: Inhabits dense forests along stagnant water courses or old stream beds, occurring in mesquite, hackberry, ebony blackbead, or huisache with a thick undergrowth of shrubs or small trees. Also frequents thickets in forest openings. NEST: Attaches half-pensile nest to upright terminal branches, twigs, and leaves 6 to 14 feet above the ground. Places nest in dense cover, usually mesquite. FOOD: Forages at midlevel in trees in dense woods for insects and small fruits. REFERENCES: Bent 1958, Kaufman in Farrand 1983c, Oberholser 1974b. 512 Northern Oriole Icterus galbula ullock's race RANGE: Breeds from southern interior British Columbia and central Alberta to central Maine and central Nova Scotia, south to southern Texas, Mexico, the central Gulf States, central North Carolina, and Delaware. Winters along the Gulf Coast and from Mexico to South America. STATUS: Common. HABITAT: In the East, inhabits orchards, deciduous forest edges, wooded river bottoms, upland forests, partially wooded suburban areas, parks, and shelterbelts. In the west, prefers semiarid mesquite groves and deciduous trees bordering streams or irrigation ditches in open country, prairie, or cultivated areas. SPECIAL HABITAT REQUIREMENTS: Tall deciduous trees for nesting. NEST: Usually attaches pendant nest by its rim to the tip of a long drooping branch, 9 to 70 feet, but typically 25 to 30 feet above the ground. Most frequently uses large trees, especially elms and cotton¬ woods growing in the open, but will use a wide variety of deciduous trees throughout its range. FOOD: Primarily gleans insects from leaf and twig surfaces; also eats a few spiders and some wild and cultivated fruit. REFERENCES: Bent 1958, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Typer in Bent 1958. 513 Scott’s Oriole Icterus parisorum L 7 " 1st year <5 RANGE: Breeds from southern California, Nevada, Utah, western Colorado, central New Mexico, and western Texas south into Mexico. Winters from southern California to southern Mexico. STATUS: Common. HABITAT: Prefers pinyon-juniper woodlands of montane semidesert areas, live oak-yucca associations, and sycamores and cottonwoods in canyons. Also uses joshua-tree habitat. SPECIAL HABITAT REQUIREMENTS: Yucca, pinyon-juniper, or oak trees in arid areas. NEST: Constructs a cup-shaped nest with fibers of local grasses, weeds, and other vegetative material and suspends it from branches of almost any available tree, including joshua-trees and yucca plants. FOOD: Probes flowers for nectar and small insects; also eats other insects, berries, and cactus fruits. REFERENCES: Phillips et al. 1964, Terres 1980. 514 RANGE: Breeds above timberline from Alaska to southwestern Alberta and south through the Cascades, Sierra Nevada, and the Rocky Moun¬ tains to east-central California, central Utah, and north-central New Mexico. In winter, descends to lower elevations (4,000 to 7,000 feet) but remains in the same general geographic area. STATUS: Common. HABITAT: Prefers barren, rocky or grassy areas and cliffs among glaciers or above timberline for breeding habitat. In winter, descends to lower elevation open habitat such as fields, cultivated lands, brushy areas, and areas of human habitation. SPECIAL HABITAT REQUIREMENTS: Talus or cliffs for nesting. NEST: Nests in the cracks or holes of cliffs that offer protection. Builds a bulky nest of grasses and dry stalks of various herbaceous materials, lining it with fine grass and feathers. FOOD: Consumes a diet that consists of vegetable material (more than 75 percent), mainly small seeds of alpine plants. Occasionally eats leaves and fruiting capsules and also consumes insects that are frozen in snow, as well as small insects found on the ground or in vegetation. REFERENCES: French 1959, Johnson 1965. 515 Pine Grosbeak Pinicola enucleator RANGE: Breeds from western and central Alaska and northern Yukon east to northern Quebec and northern Labrador, south to central California and northern New Mexico, and, east of the Rockies, to northern Alberta, southern Ontario, and central Maine. Winters from central Alaska and southern Mackenzie east to Labrador and south throughout the breeding range; in invasion years occurs farther south. STATUS: Locally common. HABITAT: Inhabits coniferous forests near timberline in the western mountains, and northern spruce-fir forests up to treeline in Canada and Alaska. Also occurs in spruce stands bordering bogs, barren areas with clumps of dwarf spruce and tamarack, and mixed coniferous and deciduous woodlands. Prefers stands with large trees and low to intermediate canopy cover, usually near the edge of an open area or along a forest border. SPECIAL HABITAT REQUIREMENTS: Coniferous forest of spruce-fir or pine. NEST: Constructs nest in thick foliage near the end of a horizontal bough in conifers, often spruce, sometimes in underbrush. May place nest low in a conifer or up to 35 feet above the ground. FOOD: Primarily eats buds and seeds of a variety of coniferous and deciduous trees, picked and gleaned from trees or from the ground. Also forages on some spiders and insects in spring and summer. REFERENCES: Bent 1968a, Blake in Bent 1968a, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979, Taber in Bent 1968a, Verner and Boss 1980. 516 Purple Finch Oarpodacus purpureus 9 RANGE: Breeds from northern British Columbia, southern Yukon, and northern and central Alberta east to central Ontario and Newfoundland south to Baja California, and, east of the Great Plains, to central Minnesota, northern Ohio, West Virginia, and southeastern New York. Winters from southwestern British Columbia south to Baja California, and from southern Manitoba east to Newfoundland, south to Texas, the Gulf Coast, and Florida. STATUS: Common. HABITAT: Generally inhabits coniferous forest edge, open mixed woodlands, and evergreen plantations. Also occurs in conifers in parks and residential areas. In winter, commonly congregates around houses with feeding stations, roosting in dense evergreens or thickets. NEST: In the East, places nest on a horizontal branch of a conifer, especially spruce, but occasionally in a deciduous tree or shrub. In the West, commonly nests in both deciduous and coniferous trees, prefer¬ ably near water. Often locates nest near tree tops, up to 40 feet above the ground. FOOD: During winter and spring, primarily eats seeds, while in late spring and summer adds insects and wild and cultivated fruits to diet. Gleans much of its food from the branches of trees and shrubs. REFERENCES: Bent 1968a, DeGraff et al. 1980, Forbush and May 1955, Johnsgard 1979. 517 Cassin’s Finch Carpodacus cassinii RANGE: Resident in western United States, in the mountains of the Pacific States, Idaho, and Montana, south into New Mexico and Arizona. Extends into Canada to breed, south into Mexico in winter. STATUS: Common in western conifers. HABITAT: Inhabits conifer forests, up to lodgepole pine type, with low to intermediate canopy cover; frequents forest edges more than interiors. In winter, moves to shrubby, bushy areas, as well as open areas with scattered trees. SPECIAL HABITAT REQUIREMENTS: Conifer forests. NEST: Nests in large conifers, near the ends of limbs quite high above the ground. Usually constructs nest of twigs, wood stems, rootlets, and lichen, and lined with hair and rootlets and sometimes bark. FOOD: Primarily eats vegetable material, preferring buds, berries, and seeds, particularly those of conifers. Also eats some animal material, but only incidentally. REFERENCES: Samson 1976, Verner and Boss 1980. 518 House Finch Carpodacus mexicanus RANGE: Breeds in disjunct distribution throughout most of the western United States, south into Texas and Mexico, and in the northeastern part of the United States, north of the Carolinas, into New England. Winters throughout the breeding range in the west and south into the Appalachian Mountains. STATUS: Abundant. Expanding range in the East. HABITAT: Inhabits rural, suburban, and urban yards, parks, farms, and open woodlands, as well as wooded areas with large openings, chaparral, and arid shrublands. NEST: Builds nests in a variety of sites, including tree cavities, and in dense outer foliage of trees and shrubs. In the East, generally chooses conifers, locating nest about 20 feet above the ground. Usually constructs nest with a variety of local grass and other available vegetation. FOOD: Forages on the ground for a variety of vegetative material, in¬ cluding seeds and fruits. Also forages in domestic fruit trees, sometimes doing considerable damage. Consumes widely varing kinds of foods, depending on local availability; includes some insects. REFERENCES: DeGraff et al. 1980, Evenden 1957, Terres 1980. 519 Red Crossbill Loxia curvirostra RANGE: Resident from southern Alaska to Newfoundland, through southern Canada, and south to the northern Great Lakes area. Also frorr the Pacific states over to the Rocky Mountains, south into parts of Mexico. Wanders irregularly during the nonbreeding season. STATUS: Locally common in conifer forests. HABITAT: Inhabits coniferous and mixed coniferous-deciduous forests, humid pine-oak associations, and lowland pine savannah. Prefers forests with low canopy cover and is highly nomadic in response to changes in conifer seed production. SPECIAL HABITAT REQUIREMENTS: Conifer forests with abundant seed production. NEST: Builds saddled nest well out on a branch or in a cluster of conifer leaves 5 to 80 feet above the ground. Nests during all months when suitable seed crops are available, but mostly from February through June. FOOD: Feeds primarily on seeds of conifer trees, plus seeds of deciduous trees in summer. Also eats insects during spring and summer. REFERENCES: Forbush and May 1955, Griscom 1937, Snyder 1954. 520 i/Vhite-winged Crossbill .oxia leucoptera L 5 3 A' RANGE: Resident from northeastern Alaska, south to the Pacific Northwest, throughout most of southern Canada, the eastern coast of Canada, and northern New England. STATUS: Irregular. HABITAT: Inhabits boreal forests, leaving them only when food is scarce. SPECIAL HABITAT REQUIREMENTS: Conifer forests. NEST: Builds a deep, saucer-shaped nest composed of lichens and twigs on branches of conifer trees. Frequently locates nest in a spruce tree, sometimes relatively close to the ground (2 to 70 feet). May breed during any month of the year, but mostly from January to May. FOOD: Consumes a variety of foods, but seems to prefer seeds of Norway spruce and hemlock. Also eats insects, seeds of other conifers and deciduous trees, and some fruits. REFERENCES: Forbush and May 1955, Terres 1980. 521 Common Redpoll Carduelis flammea RANGE: Breeds in the northern Arctic region from northern Alaska to Baffin Island, and south to northern British Columbia, and east to Newfoundland. Resident in southern part of the breeding range. Winters from Alaska and northern Saskatchewan east to Newfoundland and south to about mid-United States. STATUS: Irregular. HABITAT: Inhabits open fields with scattered small spruces or other trees, or small shrubs in the north circumpolar region. In winter, forms large flocks and moves erratically, occasionally to the edge of forests. SPECIAL HABITAT REQUIREMENTS: Open field sites with perch areas NEST: Generally places nest 3 to 7 feet above the ground, either in a dwarf tree or shrub or on the ground in open tundra. Conceals nest in vegetation, generally near lakes or ponds. FOOD: Gleans seeds from birches, alders, pines, willows, and shrubs. Also forages on the ground for seeds of grasses, forbs, and trees, and for insects. Moves during winter to find sources of food. REFERENCES: Forbush and May 1955, Grinnell 1943. 522 Hoary Redpoll ■arduelis hornemanni L5" 6 RANGE: Breeds in western and northern Alaska, northern Yukon, lorthern and east-central Mackenzie, southern Victoria Island, Keewatin, lortheastern Manitoba, Southhampton Island, and northern Quebec. Mso on Ellesmere, Bylot, and northern Baffin Islands, and in northern Greenland. Winters in the breeding range (except extreme northern areas) and south, irregularly, to southern Canada, and Montana to aorthern Illinois, New York, and New England. HABITAT: Breeds in shrubby areas, including sparse, low vegetation in apen tundra. In winter, inhabits open areas, fields, and open woodlands. MEST: Builds nest with dry grass, rootlets, and willow down or feathers; places it in low shrubs such as willow or dwarf birch and often over water. FOOD: Mostly eats plant materials, especially seeds, but some insects and larvae in summer. Forages on the ground, and gleans food from branches and leaves of shrubs. REFERENCES: Bent 1968. 523 Pine Siskin Carduelis pinus L 4W RANGE: Breeds from central Alaska south through the Western United States and east across Canada to Newfoundland. Winters throughout Eastern and Southern United States from New England, the Great Lakes, and the Northern Plains States, south into Mexico. Resident in the southern Rocky Mountains, southwestern deserts, and the Pacific Coast. STATUS: Irregularly common in large flocks. HABITAT: Generally inhabits coniferous forests, and prefers those with low to intermediate canopy cover. Less numerous in second-growth alders, aspen, and broadleaf trees along the fringes of coniferous forests. Very social, forms flocks with other species. SPECIAL HABITAT REQUIREMENTS: Coniferous forests. NEST: Generally locates nest on a conifer branch, often in the shelter of another small branch and usually 15-35 feet above the ground. FOOD: An opportunistic feeder that gleans food from foliage, bark of conifers, and from the ground. Also eats seeds, some berries, and a wide variety of insects. REFERENCES: Forbush and May 1955, Weaver and West 1943, Terres 1980. 524 Lesser Goldfinch Oarduelis psaltria RANGE: Resident from southwestern Washington, western Oregon, northern California, northern Utah, northern Colorado, northwestern Oklahoma, and central and southern Texas south to Baja California and South America. STATUS: Common. HABITAT: Generally inhabits scattered trees, woodland edge, second growth, open fields, pastures, and human habitation. Often found in drier foothill regions, in the deserts, and up to 7,300 feet in elevation and usually near water. SPECIAL HABITAT REQUIREMENTS: Scattered trees and nearby water. NEST: Saddles nest on a branch of a shrub or tree. Generally locates nest in dense foliage, 2 to 30 feet above the ground. Sometimes nests in loose colonies. FOOD: Often forages in large flocks throughout the year, gleaning food from or near the ground; rarely feeds in tree crowns. Eats seeds, fruits, flowers, and other plant material, as well as small quantities of insects. REFERENCES: Coutlee 1968, Linsdale 1957, Verner and Boss 1980. 525 Lawrence’s Goldfinch Carduelis lawrencei 2 L 4VV winter RANGE: Breeds from central California south to southern California, northwestern Baja California, and western Arizona Winters from north- central California, central Arizona, southwestern New Mexico, and western Texas south to northern Baja California, southern Arizona, and northern Sonora. STATUS: Locally common. HABITAT: Inhabits oak woodlands, chaparral, riparian woodlands, pinyon-juniper associations, and weedy areas in arid regions, usually near water. SPECIAL HABITAT REQUIREMENTS: Scattered oak trees and water. NEST: Prefers to nest in lichen-infested blue oaks that grow close together on dry slopes. Also nests in sycamores, and will often nest in small colonies. Locates nests about 20 feet above the ground in the forks of tree branches, usually 3 to 6 feet from the top of a main branch FOOD: Primarily eats seeds, gleaning them from foliage and the ground Mostly eats weed seeds, but also shrub seeds and insects. REFERENCES: Coutlee 1968, Linsdale 1957, Verner and Boss 1980. 526 American G C — 'arduelis tristis winter <5 RANGE: Breeds from southern British Columbia and north-central Mberta east to central Ontario and southwestern Newfoundland, and south to California, southern Colorado, northeastern Texas, central Mabama, and South Carolina. Winters from southern British Columbia, the northern United States, southern Ontario, and Nova Scotia south to Mexico, the Gulf Coast, and southern Florida. STATUS: Common. HABITAT: Frequents habitats with thistles or cattails, such as open weedy fields, farmyards, swamps, pastures with scattered trees, and forest edges. In the West, inhabits riparian areas, especially those with willows present along streams, ditches, and ponds. SPECIAL HABITAT REQUIREMENTS: Open weedy fields and scattered woody growth for nesting. NEST: Usually builds nest in a cluster of upright branches or on a horizontal limb of a wide variety of trees or shrubs, typically 5 to 15 feet above the ground. Delays nesting until there is an abundant supply of seeds, particularly those of composites and thistles, to feed the young. FOOD: Primarily eats seeds from a variety of plants, especially those in the composite family, as well as some insects in spring and summer, fruits, and succulent vegetation. Gleans food from the tips of weed stalks, the ground, and plants. REFERENCES: Coutlee 1968, DeGraff et al. 1980, Forbush and May 1955, Holcomb 1969, Johnsgard 1979, Stokes 1950, Tyler in Bent 1968a. 527 Evening Grosbeak Coccothraustes vespertinus RANGE: Breeds from southwestern and north-central British Columbia t< Nova Scotia and south, in the mountains, to central California, west- central and eastern Nevada, southeastern Arizona, southern New Mexico, the Mexican highlands, and, east of the Rocky Mountains, to Minnesota, Michigan, and Massachusetts. Winters throughout the breeding range sporadically south to southern California, southern Arizona, the Gulf Coast, and central Florida. STATUS: Locally abundant. HABITAT: Favors coniferous forests (primarily spruce and fir), throughou most of its range, often extending into areas where trees are quite sparse and into mixed forests. In winter, forms large flocks and may move downslope to oak or pine-oak habitats, parks, and around towns. SPECIAL HABITAT REQUIREMENTS: Conifers. NEST: Usually places nest on a horizontal limb of a conifer, 20 to 100 feet above the ground. Builds a shallow cup, usually in a dense cluster of leaves near the end of a branch. FOOD: Eats seeds, fruits, and buds from a variety of trees and shrubs. In summer, the diet includes insects. Picks vegetable matter from the branches and the ground; gleans insects from branches or hawks them in the air. REFERENCES: Forbush and May 1955, Verner and Boss 1980. 528 House Sparrow >asser domesticus L 5'A” HANGE: First introduced from Europe in 1850, became established several years later. Now resident throughout most of North America, up o the edge of the tundra and south into Central America. STATUS: Abundant. HABITAT: Prefers to stay in the neighborhood of human dwellings; avoids large forests or other habitat far from human habitation. During winter, roosts in sheltered places such as open sheds and unoccupied Duildings, under eaves or thick vines, or in a variety of holes and :revices. NEST: Constructs nest with grass, string, and any locally available material, in cavities, crevices, open areas under eaves, on ledges of buildings, and in trees or shrubs. Nests in colonies in some areas. FOOD: Consumes a wide variety of food, most of which is gleaned from the ground; also eats fruits from trees. Eats a variety of insects, vege¬ tables, fruits, seeds, grains, and garbage. Is considered a pest around farms and feedlots, where it eats livestock feeds. REFERENCES: DeGraff et al. 1980, Forbush and May 1955, Summers- Smith 1958, 1963. 529 Eurasian Tree Sparrow Passer montanus RANGE: Introduced from Europe and established in St. Louis, Missouri, in 1870; now also found in east-central Missouri and western Illinois. STATUS: Locally common. HABITAT: Occurs in residential areas, farmlands, fields, and open woodland around St. Louis, where it was introduced in 1870. It is far les agressive than its relative the house sparrow, but has slowly expanded its range to western Illinois. NEST: Nests in natural cavities, crevices, and woodpecker holes, but appears to be a weak competitor for available nesting sites. FOOD: Feeds primarily on the ground on weed seeds, insects, and waste grain. (Little information is available on food habits.) REFERENCES: Bent 1958, Scott et al. 1977, Van Benthuysen in Farrand 1983c. 530 BIRD/COVER-TYPE MATRICES We have attempted to summarize bird use of the major forest types, grasslands, and deserts of the United States. We have indicated bird use )f wetland and other open habitats commonly encountered within forests, ind conversely, in wooded habitats within deserts and grasslands, but lave not emphasized coastal habitats. This publication is primarily ntended to serve as a reference for forest and rangeland managers. Information provided in the following matrices was compiled from nany sources. The 20 habitat matrices for the eastern and western orest types are based upon the forest cover type groups in “Forest :over types of the United States and Canada,’’ F. H. Eyre, editor, Society M American Foresters, Washington, D.C., 1980. The habitat matrices ^resented for the Great Plains are based upon regions as described in ‘Land resource regions and major land resource areas of the United States,’’ Soil Conservation Service, U.S. Department of Agriculture Handbook 296, Washington, D.C., 1981. Nonforest habitat types within these broad forest cover types and Plains regions, and their use by birds in both the breeding and non- breeding seasons, were developed from the following: “Management of southern forests for nongame birds,” R. M. DeGraaf, technical coordinator, General Technical Report SE-14, Southeastern Forest Experiment Station, Forest Service, U.S. Department of Agriculture, Asheville, NC, 1978 “Nongame bird habitat management in the coniferous forests of the western United States,” R. M. DeGraaf, technical coordinator, General Technical Report PNW-64, Pacific Northwest Forest and Range Experiment Station, Forest Service, U.S. Department of Agriculture, Portland, OR, 1978 “Management of north-central and northeastern forests for nongame birds,” R. M. DeGraaf, technical coordinator, General Technical Report NC-51, North Central Forest Experiment Station, Forest Service, U.S. Department of Agriculture, St. Paul, MN, 1979 “Management of western forests and grasslands for nongame birds,” R. M. DeGraaf, technical coordinator, General Technical Report INT-86, Intermountain Forest and Range Experiment Station, Forest Service, U.S. Department of Agriculture, Ogden, UT, 1980 “New England wildlife: habitat, natural history, and distribution,” R. M. DeGraaf and D. D. Rudis, General Technical Report NE-108, Northeastern Forest Experiment Station, Forest Service, U.S. Department of Agriculture, Broomall, PA, 1986 “Guide to bird habitats of the Ozark Plateau,” K. E. Evans and R. A. Kirkman, General Technical Report NC-68, North Central Forest Experiment Station, Forest Service, U.S. Department of Agriculture, St. Paul, MN, 1981 531 “Bird-habitat relationships on southeastern forest lands,” P. B. Hamel, H. E. LeGrand, Jr., M. R. Lennartz, and S. A. Gauthreaux, Jr., General Technical Report SE-22, Southeastern Forest Experiment Station, Forest Service, U.S. Department of Agriculture, Asheville, NC, 1982 “Birds of the Great Plains,” P. A. Johnsgard, University of Nebraska Press, Lincoln, NB, 1979 “Wildlife habitats in managed forests—the Blue Mountains of Oregon and Washington,” J. W. Thomas, technical editor, Agriculture Handbook 553, Forest Service, U.S. Department of Agriculture, Washington, DC, 1979 “California wildlife and their habitats; western Sierra Nevada,” J. Verner and A. S. Boss, technical coordinators, General Technical Report PSW-37, Pacific Southwest Forest and Range Experiment Station, Forest Service, U.S. Department of Agriculture, Berkeley, CA, 1980; and many unpublished reports provided by USDA Forest Service regional offices and colleagues. Eastern Forest Cover Type Groups Eastern White-Red-Jack Pine. Eastern white pine ( Pinus strobus ), red pine ( Pinus resinosa), or jack pine ( Pinus banksiana) composes most of the stocking. The northeastern and Lake States pine types all occur as essentially pure stands, usually on lighter soils. Eastern white pine occurs from the Canadian Maritime Provinces, across the Lake States to Manitoba, and down the Appalachian Mountains to Georgia. It is shade- tolerant and also occurs as a scattered tree in other types, and on many soils. Red pine is most extensive in the Lake States and southern Ontario, and also extends east to New England, Quebec, and the Mari¬ time Provinces, where it usually occurs on small outwash areas, rocky slopes, or hilltops. It is shade-intolerant, and occurs in even-age stands. Jack pine is mainly found in the Lake States; it characteristically origi¬ nates after fire and is a short-lived, intolerant pioneer on dry, sandy soils Red Spruce-Balsam Fir. Red spruce (Picea rubens) or balsam fir (Abies balsamea) composes most of the stocking. These species frequently occur together from the Maritimes and adjacent Quebec, northern New England, New York, and the Appalachians. Either may be pure or compose a majority of the stocking; paper birch (Betula papyrifera), aspen (Populus tremuloides and P. grandidentata), red maple (Acer rubrum ), eastern white pine, and northern white cedar ( Thuja occidentalis) are common associates. Red spruce is long-lived and shade-tolerant; disturbance creates conditions favorable for establishment of balsam fir. 532 Longleaf Pine-Slash Pine. Longleaf pine ( Pinus palustris) or slash ine (P elliottii) composes a majority of the stocking. This type occurs on ie Gulf and Atlantic coastal plains from Louisiana to South Carolina, on range of sites from sandy ridges to poorly drained flatwoods. ixcluding fire allows slash pine to become established, and hardwoods md shrubs commonly proliferate. Where longleaf pine stands are treated ^ith prescribed fire, an open understory results. Loblolly Pine-Shortleaf Pine. Loblolly ( Pinus taeda) and shortleaf (P. ichinata) pines together compose the majority of the stocking. Loblolly line predominates except on drier sites; the type occurs from Delaware ;outh along the Atlantic coastal plain and Piedmont to Florida and west ilong the Gulf coastal plain to east Texas. Typically found on moist sites, t spreads to drier sites if fire is controlled. The type is succeeded by jpland oaks. Oak-Pine. Upland oaks and pines (usually loblolly or shortleaf) each :omprise 25 percent of the stocking. Oak and pine types generally occur ; rom east Texas to Georgia on upland sites on the Gulf coastal plain and Piedmont, and north in smaller areas through the Appalachians to include table mountain pine (P. pungens)- oak, Virginia pine (P virginiana)- oak, and pitch pine (P. resinosa)- oak types. Oak-Hickory. Upland oaks and hickories ( Carya spp.) compose most of the stocking, and pines constitute less than 25 percent of the stocking. Oak-hickory forests occur across a wide geographic range from Texas, Missouri, and Iowa to southern New England, with many oak and other hardwood species involved under various physiographic conditions. Oak-Gum-Cypress. In these bottomland forests of the lower Mississippi River Valley and those of its major tributaries from the Ohio River south, tupelo ( Nyssa ), blackgum (A/, sylvatica ), sweetgum (, Liquidambar styraciflua), oak (Quercus), or bald cypress ( Taxodium distichum ), singly or in combination, compose most of the stocking; pines contribute less than 25 percent of the stocking. Elm-Ash-Cottonwood. Elm ( Ulmus ), ash ( Fraxinus ), cottonwood (Populus deltoides), or red maple compose most of the stocking in these forests. Common associates in river bottoms (especially the Missouri River drainage) are sycamore (Platanus) and willow (Salix). On uplands, including those in Lake States, western New York, and southern New England, common associates are red maple ( Acer rubrum) and American beech ( Fagus grandifolia). Northern Hardwoods. Sugar ( Acer saccharum) or red maple, American beech, or yellow birch ( Betula alleghaniensis), singly or in combination, compose most of the stocking. The northern hardwood 533 type group varies geographically in its composition. It extends from the Maritimes through Wisconsin and south through the central Appalachians. Sugar maple is characteristic of the type group. Beech is absent in much of its western extent and on wetter sites in the East, where red maple and yellow birch also become common. Balsam fir and red spruce are common associates in the Northeast, aspen is common throughout; northern red oak ( Quercus rubra), white ash (Fraxinus americana), eastern white pine, paper birch, and eastern hemlock ( Tsuga canadensis) are commonly associated in the central and southern parts of the range, where the type is often called mixed woods. Aspen-Birch. Quaking and bigtooth aspens or paper birch compose a majority of the stocking. Aspen and paper birch are transcontinental in distribution. Both are pioneer types that establish after fire and clearcutting. Aspen is unique in that almost all stands regenerate from root suckers. The type is short-lived and is succeeded on dry sites by red pine, red maple, or oaks; on intermediate sites by white pine; on moist fertile sites by northern hardwoods; and on the wettest sites by balsam fir. Paper birch is succeeded by spruce-fir in the northern parts of its range and by northern hardwoods and eastern hemlock on well- drained, fertile sites elsewhere. Western Forest Cover Type Groups Douglas-fir. Douglas-fir (Pseudotsuga menziesii) composes most of the stocking. Common associates are western hemlock ( Tsuga heterophylla), western redcedar ( Thuja plicata), true firs (Abies), redwood (Sequoia sempervirens), ponderosa pine (Pinus ponderosa), and larch (Larix). The type group predominates in the Pacific Northwest, but also occurs (decreasing southward) throughout the Rocky Mountains south to northern New Mexico. Hemlock-Sitka Spruce. Western hemlock and/or Sitka spruce (Picea sitchensis) compose most of the stocking. Common associates include Douglas-fir, silver fir (Abies amabilis), and western redcedar. The type comprises the coastal forests of Washington and Oregon. Redwood. Redwood (Sequoia sempervirens) composes most of the stocking. The type is restricted to the California fog belt, extending from southernmost Oregon south along the Pacific Coast to the Santa Lucia Mountains. The type extends inland to the reaches of coastal fogs. Common associates are Douglas-fir, grand fir (Abies grandis), and tanoak (Lithocarpus densiflorus). Ponderosa Pine. Ponderosa pine composes most of the stocking. Common associates in the western part of the range (California, Oregon) 534 iclude Jeffrey (P. jeffreyi), and sugar (P. lambertiana) pines; to the north, 'ouglas-fir and incense-cedar ( Libocedrus decurrens ); to the east, limber 3 flexilis), Arizona (P. ponderosa var. arizonica ), and Chihuahua (P. )iophylla var. chihuahuana) pines; and throughout, white fir (Abies oncolor). The type is generally distributed to the west, north, and east f the Great Basin and the deserts of the Southwest. Western White Pine-Larch. Western white pine ( Pinus monticola) omposes most of the stocking. The type attains its best development in orthern Idaho and northwestern Montana. Common associates include je stern redcedar, larch, white fir, Douglas-fir, lodgepole pine (P. contorta), ind Engelmann spruce (Picea engelmannii). Such admixtures produce he “mixed conifer” type, as it is known locally. Western larch ( Larix )ccidentalis) comprises a plurality of the stocking in some areas between he Columbia River in eastern Washington and the west slopes of the tocky Mountains in Montana. Common associates are Douglas-fir, grand ir, western redcedar, and western white pine. Lodgepole Pine. Lodgepole pine composes most of the stocking; he mid-elevation type occurs to 11,000 feet in the Rocky Mountains, to 11,500 feet in California, and to 6,000 feet in Oregon and Washington. Best development is on moist, sandy, or gravelly loam. Common associates are subalpine fir ( Abies lasiocarpa), western white pine, Engelmann spruce, aspen, and larch. Fir-Spruce. The true firs, Engelmann spruce, or Colorado blue spruce ( Picea pungens) compose most of the stocking. Common associates are lodgepole pine and, at high elevation, mountain hemlock (Tsuga mertensiana). Aspen-Hardwoods. Aspen (Populus tremuloides) or red alder ( Alnus rubra) compose a majority of the stocking. The aspen type is the most common and extensive hardwood type in the western United States. It occurs primarily at middle elevations on a variety of sites in the Rocky Mountain cordillera, where it is usually succeeded by interior Douglas-fir. Aspen is usually first to dominate burns and other disturbed areas, where it produces even-aged stands. Where conifer seed sources are absent, aspen may exist as a virtual climax, where it vegetatively reproduces repeatedly, developing into all-aged stands. All western aspen communities have an herbaceous understory, commonly forbs, but sometimes grasses and sedges. In the northern portion of the type range in the West, willows, common bearberry, and buffaloberry are common understory shrubs. Farther south, snowberry, chokecherry, and western serviceberry are more common. Red alder is essentially coastal and the most important hardwood of the Pacific Northwest; best growth is on moist, rich, loamy bottomlands. 535 Chaparral. Chaparral consists of heavily branched, dwarfed trees or shrubs, commonly evergreens, whose canopy at maturity covers at least 50 percent of the ground. Common constituent plants include oaks (Quercus ), Mountain-mahogany ( Cercocarpus ), silktassel (Garrya), ceanothus (Ceanothus), manzanita ( Arctostaphylos ), and chamise (Adlenostoma). Pinyon-Juniper. Pinyon pines (primarily P edulis , P cembroides, P monophylla) and junipers (primarily Juniperus osteosperma , J. deppeana , and J. monosperma) compose most of the stocking. This type is widely distributed throughout the semiarid West, usually on dry, shallow, rocky soils of mesas, benches, and canyon walls. Eastern Open, Wetland, Plains, Deserts, and Other Nonforest Habitats Field, Glade, Orchard. Primarily grass, hayfields, abandoned agricultural land, and fruit orchards with grassy ground cover. Pasture, Wet or Sedge Meadow. Agricultural lands that are too wet. steep, or rocky for crops; meadows dominated by grasses or sedges (Carex spp.) with soils that are saturated or seasonally flooded. Fresh Marsh, Pond. Palustrine and lacustrine wetlands, permanently flooded, containing emergents such as cattails (Typha), bullrushes ( Scirpus ), rushes (Juncus), and floating-leaved plants: spatterdock ( Nuphar ) and water lily (Nymphaea). Wooded Swamp, Bog, Shrub Swamp. Palustrine, forested wetlands, either needle-leaved evergreen or broad-leaved deciduous; dominant plants are Atlantic white-cedar (Chamaecyparis thyoides ), black spruce ( Picea mariana), or red maple. Wooded swamps are seasonally or permanently flooded; bogs are permanently flooded. Lake, Stream, River. Stratified lacustrine wetlands; permanently flowing watercourses of any width. Sand Pine, Scrub Oak. Southeastern and southern woodlands on droughty, infertile, coarse-textured or sandy soils that support any of the scrub oaks (0. laevis, 0. incana, Q. marilandica , or 0. stellata var. margaretta) or sand pine (Pinus clausa). Pocosins. Bay-swamp, and pond pine ( Pinus serotina) woodlands with boggy soils in which broadleaf evergreens predominate: black gum (Nyssa sylvatica var. biflora ), persea (Persea barbonia), magnolia (Magnolia virginiana), gordonia ( Gordonia lasianthus) and associates, of pond pine, Atlantic white-cedar. 536 Everglades, Mangroves. Palustrine wetlands in southern Florida nat are semi-permanently flooded, dominated by saw grass ( Cladium imaicense ); estuarine, intertidal wetlands dominated by mangrove Rhizophora). Alpine Tundra, Krummholtz. Elevated slopes above timberline :haracterized by low, shrubby, slow-growing woody plants and a ground :over of boreal lichens, sedges, and grasses; the transition zone from ;ubalpine forest to alpine tundra characterized by dwarfed, wind-sheared rees. 3reat Plains Habitats Gulf Prairies and Marshes. Moderate to tall dense open grasslands jominated by seacoast bluestem ( Andropogon littoralis) and coastal jacahuista ( Spartina spartinae). East Texas Prairies, Cross Timbers, Pineywoods, and Post Oak Savanna. Open grassy savanna to dense brushland occurring in north- central Texas; pine-hardwood forest and grazing lands of east Texas; midgrass prairie dominated by little bluestem (Andropogon scoparius) and shin oak ( Quercus mohriana). South Texas Shrub-Grassland. Vegetation ranges from desert grass- shrub vegetation in west Texas to mixed oak savanna in the eastern Edwards Plateau region to open grassland on the Rio Grande plain. Southern Plains. Open to moderately dense short grasslands occurring from southeastern Colorado and central Oklahoma south through eastern New Mexico and Texas panhandle. Natural vegetation is characterized by grama ( Bouteloua ) and buffalo grass ( Buchloe). Central Plains. Grasslands ranging from short grasses in the West to tall grasses in the East. Includes the region from southeastern Wyoming and northeastern Colorado east through Indiana. Northern Plains. Plains region from north-central Montana and northwestern Minnesota south to southeastern Wyoming and northwestern Iowa. Area supports grassland vegetation dominated by western wheatgrass (Agropyron smithii) and needlegrass ( Stipa ) in the West and by little bluestem in the East. Wetland and Riparian Habitats. All Great Plains wetland and riparian habitats, including marshes, ponds, lakes, streams, stock ponds, and woodlands associated with wetlands. 537 Shelterbelts and Woodlots Planted bands of trees that serve as windbreaks to protect fields or farmsteads; other wooded areas surrounded by agricultural lands. Pine-Oak, Brushy Woodland, Badlands-Juniper. Includes dry woodlands of pines and oaks with or without a brushy understory. Badlands-juniper region is largely devoid of vegetation but may have scattered junipers on suitable sites. Southwestern and Western Nonforested Habitats Relict Conifer Forest, Madrean Evergreen Woodland Warm- temperate forests and woodlands in the Southwest. Relict conifer forests consist of small populations of cypress ( Cupressus ) and closed-cone pines, bishop pine ( Pinus muricata), and knobcone pine (P. attenuata), restricted to canyons and suitable slopes along drainages. Madrean evergreen woodland is composed primarily of evergreen oaks but includes madrean pines. Aspen. Mid-elevation sites in the Great Basin consisting of pure or nearly pure stands of quaking aspen. Great Basin Shrubsteppe. Open to dense stands of shrubs and low trees, including big sagebrush ( Artemisia tridentata), saltbush ( Atriplex confertifolia), greasewood ( Sarcobatus vermiculatus), or creosote bush {Larrea divaricata). Sonoran Desert Scrub. Open to dense vegetation of shrubs, low trees, and succulents dominated by paloverde ( Cercidium microphyllum), pricklypear ( Opuntia spp.), and giant saguaro (Cereus giganteus). Chihuahuan Desert Scrub. Open stands of creosote bush and large succulents (Ferocactus pringlei , Echinocactus ptatyaconthus) in southern New Mexico and southwest Texas. Mohave Desert Scrub. Located between the Great Basin desert scrub and the Sonoran desert scrub, it is intermediate between them, sharing plant species of both but containing the endemic arboreal leaf succulent, Joshua tree (Yucca brevifolia). Desert Riparian Deciduous Woodland, Marsh. Woodlands, especially of cottonwoods, that occur where desert streams provide sufficient moisture for a narrow band of trees and shrubs along the margins. 538 Annual Grasslands, Farms. Grasslands dominated by wild oat Avena spp.), ripgut brome (Bromus rigidus), soft chess (Bromus mollis), >ur clover ( Medicago hispida), and filaree (Erodium spp.) with less than 5 >ercent woody cover. River, Riparian Woodland, Subalpine Marsh. Occurs at elevations vhere stream conditions provide sufficient permanent moisture for emergent plants, or for a narrow band of deciduous trees and shrubs; at ow elevation characterized by cottonwood and sycamore, at mid- slevation by white alder ( Alnus rhombifolia) and bigleaf maple (Acer vacrophyllum), and at high elevation by willow. Mountain and Alpine Meadows. Sedges ( Carex) and grasslike plants ( Heleocharis , Scirpus) above treeline. 539 Table 1 —Birds occurring in eastern forest types Bird Species Double-crested Cormorant Anhinga American Bittern Least Bittern Great Blue Heron Great Egret Snowy Egret Little Blue Heron Cattle Egret Green-backed Heron Black-crowned Night-Heron Yellow-crowned Night-Heron Glossy Ibis White-faced Ibis Wood Duck American Black Duck Gadwall Ring-necked Duck Common Goldeneye Bufflehead Hooded Merganser Common Merganser Black Vulture Turkey Vulture Osprey American Swallow-tailed Kite Missippi Kite Bald Eagle Sharp-shinned Hawk Cooper’s Hawk Northern Goshawk Red-shouldered Hawk Broad-winged Hawk Short-tailed Hawk Red-tailed Hawk Golden Eagle American Kestrel Merlin Peregrine Falcon Spruce Grouse CD - cp x: C/5 CD CD l6 Q) _>* Q) O O 03 O r~ c X) § Q_ c n 0 _l O _l X X X X X X X X X X X X X XX X X X X X X X X X X X X X X X X X XX X X X X X X X X X X X X X X X X X X X X X X X a> c CL jk (0 o X X X X X X X X X X X X X if) (/) 0 k— Q. > k_ O O E O 0 JZ O) A CC 03 O O X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX XXX X X X X XXX XXX X X XXX X X XXX X X XXX XXX (Continued) 540 xxx x x Elm-ash-cottonwood Table 1 —Birds occurring in eastern forest types—continued 3ird Species Ruffed Grouse Wild Turkey Northern Bobwhite yellow Rail King Rail Virginia Rail Sora Purple Gallinule Common Moorhen Limpkin Solitary Sandpiper Spotted Sandpiper Upland Sandpiper Common Snipe American Woodcock Rock Dove Mourning Dove Common Ground-Dove Black-billed Cuckoo Yellow-billed Cuckoo Common Barn-Owl Eastern Screech-Owl Great Horned Owl Northern Hawk-Owl Burrowing Owl Barred Owl Great Gray Owl Long-eared Owl Boreal Owl Northern Saw-whet Owl Common Nighthawk Chuck-will’s-widow Whip-poor-will Chimney Swift Ruby-throated Hummingbird Belted Kingfisher Red-headed Woodpecker Red-bellied Woodpecker Yellow-bellied Sapsucker Downy Woodpecker 541 (Continued) Table 1 —Birds occurring in eastern forest types—continued Bird Species Hairy Woodpecker Red-cockaded Woodpecker Three-toed Woodpecker Black-backed Woodpecker Northern Flicker Pileated Woodpecker Olive-sided Flycatcher Eastern Wood-Pewee Yellow-bellied Flycatcher Acadian Flycatcher Alder Flycatcher Willow Flycatcher Least Flycatcher Eastern Phoebe Great Crested Flycatcher Eastern Kingbird Purple Martin Tree Swallow Northern Rough-winged Swallow Bank Swallow Cliff Swallow Barn Swallow Gray Jay Blue Jay American Crow Fish Crow Common Raven Black-capped Chickadee Carolina Chickadee Boreal Chickadee Tufted Titmouse Red-breasted Nuthatch White-breasted Nuthatch Brown-headed Nuthatch Brown Creeper Carolina Wren Bewick’s Wren House Wren Winter Wren Marsh Wren XXX XXX XXX X X XXX X X XXX XXX XXX XXX X X XXX XXX XXX XXX XXX XXX XXX X XXX XXX XXX XXX X X X XXX XXX X XXX XXX X 542 (Continued) Elm-ash-cottonwood Table 1 —Birds occurring in eastern forest types—continued Bird Species Solden-crowned Kinglet Ruby-crowned Kinglet Blue-gray Gnatcatcher Eastern Bluebird Veery Gray-cheeked Thrush Swainson’s Thrush Hermit Thrush Wood Thrush American Robin Gray Catbird Northern Mockingbird Brown Thrasher Water Pipit Bohemian Waxwing Cedar Waxwing Northern Shrike Loggerhead Shrike European Starling White-eyed Vireo Solitary Vireo Yellow-throated Vireo Warbling Vireo Philadelphia Vireo Red-eyed Vireo Bachman’s Warbler Blue-winged Warbler Golden-winged Warbler Tennessee Warbler Orange-crowned Warbler Nashville Warbler Northern Parula Yellow Warbler Chestnut-sided Warbler Magnolia Warbler Cape May Warbler Black-throated Blue Warbler Yellow-rumped Warbler Black-throated Green Warbler Blackburnian Warbler CD C Q. o ■6 O) £ 5 X X X X X X X X X X XXX X XXX X X X X X X X X X X XX XX X X X X X X X X X X XXX X X X X X X X X X X X X X XXX x XX X X X X X X X X X XX XX X X X X X X XX XX X X XX X X X X X X X X X X X X XX XX X X X X X X XXX XXX X X XXX X XXX X X X X X X X X X X X X X X X X X X X X X XXX X X X X X X X X X X X X X X XX x X X X X X X X X X X X XX X X X X X X X X X X X X X X X X x XX x XX X X X X X X X X X X X X X X X x XX X x XX X X X X X X X x X (Continued) 543 Elm-ash-cottonwood Table 1 —Birds occurring in eastern forest types—continued Bird Species Yellow-throated Warbler Pine Warbler Prairie Warbler Palm Warbler Bay-breasted Warbler Blackpoll Warbler Cerulean Warbler Black-and-white Warbler American Redstart Prothonotary Warbler Worm-eating Warbler Swainson’s Warbler Ovenbird Northern Waterthrush Louisiana Waterthrush Kentucky Warbler Mourning Warbler Common Yellowthroat Hooded Warbler Wilson’s Warbler Canada Warbler Yellow-breasted Chat Summer Tanager Scarlet Tanager Northern Cardinal Rose-breasted Grosbeak Blue Grosbeak Indigo Bunting Painted Bunting Dickcissel Rufous-sided Towhee Bachman’s Sparrow American Tree Sparrow Chipping Sparrow Lark Sparrow Fox Sparrow Song Sparrow Lincoln’s Sparrow Swamp Sparrow White-throated Sparrow X X X X XXX XXX X XXX X X X X X X XXX XXX X XXX X X XXX XXX XXX XXX X X XXX XXX 544 (Continued) Elm-ash-cottonwood Table i —Birds occurring in eastern forest types—continued Bird Species i/Vhite-crowned Sparrow Dark-eyed Junco Red-winged Blackbird Rusty Blackbird Brewer’s Blackbird CO CO 2 Q_ >> o E 3 U) cc O X X Common Grackle Brown-headed Cowbird Orchard Oriole Northern Oriole Pine Grosbeak X X X X X X X X X X X X X X X Purple Finch House Finch Red Crossbill White-winged Crossbill Pine Siskin X X X X X X XX XX XX X X X X X X X x American Goldfinch Evening Grosbeak X X X X X X X 545 xx xxx xxx Elm-ash-cottonwood Table 2—Birds occurring in eastern nonforest habitats (exclusive of Great Plains) Bird Species Common Loon Pied-billed Grebe Horned Grebe Red-necked Grebe Double-crested Cormorant Anhinga American Bittern Least Bittern Great Blue Heron Great Egret Snowy Egret Little Blue Heron Cattle Egret Green-backed Heron Black-crowned Night-Heron Yellow-crowned Night-Heron Glossy Ibis Fulvous Whistling-Duck Canada Goose Wood Duck Green-winged Teal American Black Duck Mottled Duck Mallard Northern Pintail Blue-winged Teal Northern Shoveler Gadwall American Wigeon Redhead Ring-necked Duck Greater Scaup Lesser Scaup Common Goldeneye Bufflehead Hooded Merganser Common Merganser Red-breasted Merganser Ruddy Duck Black Vulture _ O CD JZ ■O s sz o l— CD O) ~o CD CD *D C o Q_ 0) O -Q Q. 0 > CO O -Q 3 CD > 2 w U)~0 C o 03 o E E D ZH o O E CL - O CD E 5 03* CD CD j£ CD § E E CO CD "O ~o k_ O TD 03 £ CD CO cd ro CD C CD c 3 U) CD O E ■g s CO Q. C CD o "co 0 - 3 *o sz -O -Q CD Z£ 03 —1 *o 0)0 C "O CD LL CD 03 CL CO CD E CD CD LL O 3 II c co (f) O O Q_ Q) Q- > o LU a < X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX (Continued) 546 Table 2—Birds occurring in eastern nonforest habitats (exclusive of Great Plains)—Continued Bird Species T3 i_ 03 r~ CD CD T3 _Q 3 O CD O 0 -C (0 E 03 £ Q. E E CO k_ O CO T3 J? £ CO k_ 03 CO 03 03 CD c CO 03 CD O E "O CD co to Q. c CO " d CD Ll 3 CO 03 Q_ ~o 03 CD ■o 0 u c CD O -Q 0 03 O o C/3 o CL d > .a o o' O 0 .cl C/3 E 03 $ CL E E 03 o C/3 "O 03 £ C/3 03 C/3 03 0 0 C C/3 CD 03 O E ~o 0 C/3 CO CL C C/3 ~o a> Ll 3 CO 03 CL “O 03 0 E .c C/3 0 LL ~o o o 5 -Q -C C/3 0' 03 ■Q C 03 c n o o o Q_ C/3 0 > 2 » CD U C O 03 O E S "O ® 1 -o — CO 0)0 <5 Q. > o LU i: XXX X X X X X X X X X X X X XXX X X XXX XXX X X X X X X XXX XXX XXX x X XXX X X X X x X X X x X X x X x X X X X X X X X X X X X X X XXX X X XXX X X X x X X X X X X X X X X X X X X X X X X XXX XXX XXX X X X X X X (Continued) 548 Alpine tundra, krummholtz Table 2—Birds occurring in eastern nonforest habitats (exclusive of Great Plains)—Continued N 3ird Species Eastern Phoebe Great Crested Flycatcher lA/estern Kingbird Eastern Kingbird Gray Kingbird Scissor-tailed Flycatcher Horned Lark Purple Martin Tree Swallow Northern Rough-winged Swallow Bank Swallow Cliff Swallow Barn Swallow Gray Jay Blue Jay Scrub Jay American Crow Fish Crow Common Raven Black-capped Chickadee Carolina Chickadee Boreal Chickadee Tufted Titmouse Red-breasted Nuthatch White-breasted Nuthatch Brown-headed Nuthatch Brown Creeper Carolina Wren Bewick’s Wren House Wren Winter Wren Sedge Wren Marsh Wren Golden-crowned Kinglet Ruby-crowned Kinglet Blue-Gray Gnatcatcher Eastern Bluebird Veery Gray-cheeked Thrush Hermit Thrush ■a 03 -C o CD CD "D CD (/) TD c o Q_ CD O _Q Q. a) > k_ zc 03 O JD 3 o a> O Q) JZ CO 1 E E CO o in CD c -o 03 ^ o O ^ Q- - E 0 O 2 CD TD E 3 JD c o 13 0 O O 0 0 C Q. JD 0 c 0 o E £ _ TD 0 (Tj 0 ^ JD — 0 CD (J 0 ■Q C 3 0 C C o a) cl o 0 o > o CO CL LU ic < X X X XXX X XXX XXX XXX X X XXX XXX X X x X X x X X XXX X x X X XXX X X XXX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX XX X x (Continued) 551 Table 2 —Birds occurring in eastern nonforest habitats (exclusive of Great Plains)—Continued Bird Species Western Meadowlark Rusty Blackbird Brewer’s Blackbird Boat-tailed Grackle Common Grackle Brown-headed Cowbird Orchard Oriole Northern Oriole Purple Finch House Finch Red Crossbill Common Redpoll Pine Siskin American Goldfinch House Sparrow N CD - "D CD 03 03 f~ ~o o O E Q_ _ 0) CD -C C/3 § E E 03 03 "D 03 ^ 0) 03 w « 03 £ 1 E ■o 5 CD CO - 3 "O -C *0 -Q 03 03 _l ~o 03 LL 2| CL E C/3 03 LL O 13 > if) 03 o _Q 3 u_ O if) CD C Q. U c 03 c n O if) JZ 03 > E 2 O CL LU iz < 552 Table 3—Birds occurring in Great Plains habitats iird Species CO CD .C co co E "O c CO co 0) *i_ co CL "5 CD CO TD £ C O CO ° «r m "2 .® 8 '« I, CL C 2 >* o3 Q. CD > C 03 CO pr CO s , 0 ) 01 10 ^ s ° 05 £ 00 to E o UJ 5= Ql A 3 k_ x: 05 05 (0 X .05 3 O CO X X X X X XXX XXX XXX XXX XXX XXX XXX X X XXX x X X XXX XXX XXX X X XXX XXX X x X x X X X X X X X habitats—continued tS c CO CO C/5 (/) Q_ C C/5 C *- 03 C 03 ~o Q- 03 Q- c 03 C Q_ s "cij "O C/5 c -s sz JZ 03 £ 3 C at -Q o CD o > CO CO o z 5 -c X X X X XXX X X X XXX X X X XXX XXX XXX X XXX XXX XXX XXX XXX XXX X X X X X X X XXX X X X X X X X X X X X XXX X X XXX XXX XXX XXX XXX X XXX XXX XXX X XXX XXX XXX XXX X X X X X X X X XXX XXX (Continued; 554 Shelterbelts and woodlots Table 3—Birds occurring in Great Plains habitats—continued Sird Species Scaled Quail fellow Rail Slack Rail tapper Rail (ing Rail /irginia Rail Sora 3 urple Gallinule Common Moorehen \merican Coot Sandhill Crane Whooping Crane Slack-bellied Plover Snowy Plover 3 iping Plover ~o c if) o c 03 if) O 03 TD "O 03 e O c 6 C/3 TD sz _Q c CO O O £ o £ TD C 03 03 05 O £ > 03 C c 03 3 SZ if) if) C if) if) C 03 Q- "k_ TD c 03 >> sz if) 03 Q. 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Q. c 0 0 C -C L_ 0 o O z X X X X X X X X X X X X X X X XXX X X X X XXX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X X X X X X X X X X X c w I_ to CL T3 c CO X X X X X X X XXX XXX X X X XXX XXX XXX X X X X XXX X XXX XXX X X X X X X X X X X X X XXX X (Continued) 556 Shelterbelts and woodlots Table 3—Birds occurring in Great Plains habitats—continued 5ird Species fellow-bellied Sapsucker .adder-backed Woodpecker Downy Woodpecker Hairy Woodpecker Hed-cockaded Woodpecker Northern Flicker bleated Woodpecker Glive-sided Flycatcher A/estern Wood-Pewee Eastern Wood-Pewee Acadian Flycatcher Willow Flycatcher .east Flycatcher Eastern Phoebe Say’s Phoebe Vermilion Flycatcher Ash-throated Flycatcher Great Crested Flycatcher Brown-crested Flycatcher Couch’s Kingbird Cassin’s Kingbird Western Kingbird Eastern Kingbird Scissor-tailed Flycatcher Horned Lark Purple Martin Tree Swallow Violet-green Swallow Northern Rough-winged Swallow Bank Swallow Cliff Swallow Barn Swallow Blue Jay Scrub Jay Black-billed Magpie American Crow Fish Crow Chihuahuan Raven Common Raven Black-capped Chickadee C/5 05 SI c c 03 k_ JZ CO CO c CO CO c ( r) 0) Cl ( f) a) c > CO 0) 03 03 CL c 03 03 CL \_ 03 i— 03 X Q. (0 03 X ~o c c a3 CL 03 C CD CL H GO a> JD O GO 3 CO (0 sz 3 c k— "5 03 E o O 03 o CD O a LU Q. c n a) CO o ~z. X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X X X X X X X XXX X X X X X X X X X X X X XXX X X X X X X X X X X X X X X XXX X X X X X X X X X c <5 CO Q. TD c (0 ~0 c/5 C/5 o "O o o 5 T3 c CO T5 c « "O o o >. 1- -C CD c/5 a 05 05 XI CO si C/5 CO ■a ? § Si _ CO Q- X5 X X X X XXX XXX XXX X X X X X X X XXX X X X X XXX X X XXX XXX XXX XXX X X X X X X X X X X X XXX XXX X XXX XXX X XXX (Continued) 557 Table 3—Birds occurring in Great Plains habitats—continued Bird Species Carolina Chickadee Tutted Titmouse Verdin Bushtit Red-breasted Nuthatch White-breasted Nuthatch Brown-headed Nuthatch Brown Creeper Cactus Wren Rock Wren Canyon Wren Carolina Wren Bewick’s Wren House Wren Winter Wren Sedge Wren Marsh Wren Golden-crowned Kinglet Ruby-crowned Kinglet Blue-gray Gnatcatcher Black-tailed Gnatcatcher Eastern Bluebird Veery Swainson’s Thrush Hermit Thrush Wood Thrush American Robin Gray Catbird Northern Mockingbird Sage Thrasher Brown Thrasher Long-billed Thrasher Curve-billed Thrasher Water Pipit Sprague’s Pipit Cedar Waxwing Bohemian Waxwing Northern Shrike Loggerhead Shrike European Starling in a> .c in to E "O c CO in gj L. to CL a x X X X X X X X X X X X X X X X X X X X X X X X in T3 g to c ° c m S c 2 >•« Q. > C CO m = m to _ x - -X ,a> to to to to to E o UJ = Cl X X X X X X X X X X X X X X X X X X X X X X X X X _Q i= CO CO c if) CO CO CL X (® ■O c c k— CO CD JZ CO JZ 3 CO =3 o CO o ( f ) O) (f) CO if) c c CO CO CL CL c 2 CD C SI k_ CD o o z X X X X X X X X X X X X X X X XXX X X X X XX X X X X X X X X X X X X X X XX X X XXX X X X X X X X X X X X X X X X X X X X X X ■o X X X X X X XXX X X X X X XXX X X X XXX XXX X X X X X X X X X XXX XXX XXX XXX X X X XXX X X XXX XXX (Continued) 558 Shelterbelts and woodlots Table 3—Birds occurring in Great Plains habitats—continued lird Species Vhite-eyed Vireo Jell’s Vireo Slack-capped Vireo Solitary Vireo fellow-throated Vireo Varbling Vireo Bed-eyed Vireo JIue-winged Warbler Drange-crowned Warbler 'lorthern Parula fellow Warbler fellow-rumped Warbler Bolden-cheeked Warbler fellow-throated Warbler Cerulean Warbler Black-and-white Warbler American Redstart 3 rothonotary Warbler /Vorm-eating Warbler Bwainson’s Warbler Dvenbird Northern Waterthrush Louisiana Waterthrush Kentucky Warbler Common Yellowthroat Hooded Warbler Wilson’s Warbler Yellow-breasted Chat Summer Tanager Scarlet Tanager Northern Cardinal Pyrrhuloxia Rose-breasted Grosbeak Black-headed Grosbeak Blue Grosbeak Lazuli Bunting Indigo Bunting Painted Bunting Dickcissel Olive Sparrow co a) .c CO (/) *o O 03 l_ to ° to E CO ' "g -C _Q T3 CD o CO eye 3 k_ C 2 sz to to to .3? CO ~ CO CO Q - co to CO * -3 , CD CO CO i2? c ^ CO Q. ^ a3 ° £ « "5 to -£? co co E o 3 CO o 2 0 LU = D. CO CD X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX XXX XXX X a <5 re (l ) (J) Q_ c cn ro r- ns CD ° £ 05 — m -Q m "3 V} X XXX X X X X X X XXX X X X X XXX XXX XXX X X X XXX X X X X X XXX X X XXX XXX X X XXX XXX XXX X X X X X X XXX XXX XXX X c CO CO Southern plains Central plains Northern plains Wetland and rip. habitats X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X (Continued 560 x x Shelterbelts and woodlots Table 3—Birds occurring in Great Plains habitats—continued ird Species usty Blackbird rewer’s Blackbird ireat-tailed Grackle oat-tailed Grackle ommon Grackle rown-headed Cowbird irchard Oriole lorthern Oriole urple Finch louse Finch led Crossbill lommon Redpoll ine Siskin esser Goldfinch .merican Goldfinch ivening Grosbeak louse Sparrow if) C 05 w o c n (0 to * T3 5 X - , 03 C ^ CD Q_ "5 ^ 5 ° CO £ 00 to E o £ to ■5 to 0 £5 O LU = CL C/3 O) X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X XXX XXX c .2 ng-eared Owl lort-eared Owl ureal Owl arthern Saw-whet Owl ammon Nighthawk ammon Poorwill hip-poor-will ack Swift lux’s Swift 'hite-throated Swift lack-chinned Hummingbird nna’s Hummingbird alliope Hummingbird road-tailed Hummingbird ufous Hummingbird lien’s Hummingbird elted Kingfisher awis’ Woodpecker corn Woodpecker allow-bellied Sapsucker ed-breasted Sapsucker /illiamson’s Sapsucker adder-backed Woodpecker uttall’s Woodpecker >owny Woodpecker lairy Woodpecker i/hite-headed Woodpecker hree-toed Woodpecker Jack-backed Woodpecker lorthern Flicker 'ileated Woodpecker )live-sided Flycatcher Western Wood-Pewee Villow Flycatcher tammond’s Flycatcher )usky Flycatcher Sray Flycatcher Vestern Flycatcher Slack Phoebe in (U Q c CO k_ o c o co 3 b in CD in Cl> Q c CO 3 x: cO 3 .C !c O if) if) if) O E ~o c 03 03 _o 15 if) 5 6 CD 4- 03 *- Q. > 03 O CD "O if) O CD O 3 C c §5 c -O 3 03 O CD Q S < ix in 2 E X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X (Continued) 567 Relict conifer forests, madrean evergreen woodlands Table 5—Birds occurring in southwestern and western nonforest habitats—continued Bird Species <1) Q_ Q. 0) 00 _Q 3 to a) O c to 3 -C to 3 -C O if) 3 o W E if) *o -Q P c/> 3 c _co CD if) k_ O (/) O CD a) ~o w c co (A TD c CO o o $ if) CD .£= if) c Q. co CO 0) o a> c E if) if) CO c CO CO E ■D c CO k_ 0) a CD > aj o CL C ~ J5 a> ~o (f) o a> o k_ U) *C0 c c CO a. CD > CD C Q. CO JD c 05 c o if) o "O 03 CD c o o o 0 s o s < DC E a: Common Merganser Red-breasted Merganser Ruddy Duck Black Vulture Turkey Vulture California Condor Osprey Black-shouldered Kite Bald Eagle Northern Harrier Sharp-shinned Hawk Cooper’s Hawk Northern Goshawk Common Black-Hawk Harris' Hawk Gray Hawk Red-shouldered Hawk Swainson’s Hawk Zone-tailed Hawk Red-tailed Hawk Ferruginous Hawk Rough-legged Hawk Golden Eagle American Kestrel Merlin Peregrine Falcon Prairie Falcon Gray Partridge Chukar Ring-necked Pheasant Blue Grouse White-tailed Ptarmigan Ruffed Grouse Sage Grouse Sharp-tailed Grouse Wild Turkey Montezuma Quail Northern Bobwhite Scaled Quail Gambel’s Quail X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X (Continued) 568 Tabie 5—Birds occurring in southwestern and western nonforest habitats—continued rd Species alifornia Quail ountain Quail lapper Rail irginia Rail ora ommon Moorhen merican Coot andhill Crane /hooping Crane illdeer lountain Plover lack-necked Stilt merican Avocet ireater Yellowlegs i/illet ipotted Sandpiper Ipland Sandpiper ong-billed Curlew Garbled Godwit east Sandpiper .ong-billed Dowitcher Common Snipe Vilson’s Phalarope : ranklin’s Gull ting-billed Gull California Gull Herring Gull Caspian Tern : orster’s Tern Slack Tern Hock Dove Band-tailed Pigeon A/hite-winged Dove Vlourning Dove nca Dove Common Ground-Dove Black-billed Cuckoo Yellow-billed Cuckoo Greater Roadrunner Common Barn-owl _Q in in 0 Q. 3 3 O E CL _Q O 3 CO 0 3 (J ) _Q TO m V) 3 O 0 * -Q 3 -C= (r > O c n CD in 0 (/) 0 Q O in i— CD 0 T3 C CO sz in CO E in to c _C0 m c in CO 0) O c c CO 3 SZ in CD Q CO Q- in to c in CO CT> CD CO CO 0 CO CO CO 0 o c o 3 SZ zz > CO o i— 0 in 0 TO o o 3 C C 0 CO o O < X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X in to c _cO TO o 0 C O £ Q. Sm CO ro TO c E C CO 1c ^ Q_ c 03 in £ o C TO 5) 3 03 > ■§ O 0 DC E X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X X (Continued) 569 Relict conifer forests, madrean evergreen woodlands Table 5—Birds occurring in southwestern and western nonforest habitats—continued Bird Species Flamulated Owl Eastern Screech-Owl Western Screech-Owl Whiskered Screech-Owl Great Horned Owl Northern Pygmy-Owl Ferruginous Pygmy-Owl Elf Owl Burrowing Owl Spotted Owl Great Gray Owl Long-eared Owl Short-eared Owl Northern Saw-whet Owl Lesser Nighthawk Common Nighthawk Common Poorwill Whip-poor-will Black Swift Vaux’s Swift White-throated Swift Broad-billed Hummingbird Violet-crowned Hummingbird Blue-throated Hummingbird Magnificent Hummingbird Black-chinned Hummingbird Anna’s Hummingbird Costa’s Hummingbird Calliope Hummingbird Broad-tailed Hummingbird Rufous Hummingbird Allen's Hummingbird Elegant Trogon Belted Kingfisher Lewis’ Woodpecker Acorn Woodpecker Gila Woodpecker Red-naped Sapsucker Yellow-bellied Sapsucker Red-breasted Sapsucker 0 Q. .Q 3 CO 3 o to E to ■O Q. o o 3 k. c 0 to JD 3 3 O to (0 L_ 0 CO -Q 3 k. O ■g o 0 TJ CO 0 sz CO 3 to ~o _C0 "O o o CO 0 0 c Q. -C k_ 0 CO 0 o CO 0 C. 03 CO E c 3 CO £ c -C CO CO CO T3 C 0 c CO k_ CO CO p c (f) Q 03 0 03 CO CO "k_ CO 03 CD c 03 3 _£= 03 3 Q 0 > Q. L. k- ■Q c _co 03 To CO g. k_ 0 c Q. c To CO £ o To O r~ 03 0 ■O 3 i-T c *o 0 c CO o c 0 3 CO o !E o 0 o c > -Q o 0 O CO O Q £ < ir 3 CO E X X X X X X X X X X X X X V) Jr (D 0 Z- 0 0 > 0 v. ■c c cz o c O CO — to ^ U ±z-C — m c a> w o IE5 X X X X X X X X X X X X X X X X X X X X X X X X X X X XX X X XX X X X XX XX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X (Continued) 570 Table 5—Birds occurring in southwestern and western nonforest habitats—continued Bird Species Williamson’s Sapsucker Ladder-backed Woodpecker Nuttall’s Woodpecker Downy Woodpecker Hairy Woodpecker a> CL Q. OJ to .Q 3 k_ SI in if) 03 CO CO CD o .o 3 <5 in o. a. 05 <0 -Q 3 k_ -C (fi c (fi 0 CD -Q 3 O (/) 0 (n 0 O c 03 03 a; O o c= o c n _Q 3 k_ o (0 ai 01 ai a c as 3 -C (0 3 -C O 01 3 O -Q u (fi 0 0 3 k_ O (fi o 0 "O c 0 sz (fi (C (ji ~o c _0 "O 0 0 £ 0 0 .c 0 C Q. 0 0 "0 *“ Q. 0 O 03 0 ■Q O k-T 0 C *o O (fi 0 O o c c 0 > JD O O 0 0 Q < ir 0 E 0 0 0 c 0 k_ 0 O) w_ ^0 0 > 'E 0 c C 0 0 ,. 0 0 •c © 0 CE E XX X X X X X X X X X X X X X X X X X X X Bushtit Red-breasted Nuthatch White-breasted Nuthatch Pygmy Nuthatch Brown Creeper X X X X X X X X X X X Cactus Wren X X X Rock Wren X X X X Canyon Wren X X X X Bewick’s Wren X X X X X X House Wren X X X X X X X Winter Wren X X Marsh Wren X X X X American Dipper X X Golden-crowned Kinglet X Ruby-crowned Kinglet X X Blue-gray Gnatcatcher X X X X X Black-tailed Gnatcatcher X X X Western Bluebird X X X X X Mountain Bluebird X X X X X X X Townsend's Solitaire X X X X Veery X Swainson’s Thrush X X Hermit Thrush X X X X American Robin X X X X X X X X Varied Thrush X X (Continued) 572 Table 5—Birds occurring in southwestern and western nonforest habitats—continued Bird Species CD _Q 3 Q_ Q. 3 _Q 3 O if) tn -Q O if) 03 O _C0 ■O 75 3 _Q_ 03 if) o C LU o' CD o C > 3 Q £ < □: U) CD C Q. 03 T3 C CO c To o c T3 3 03 o CD E Wrentit Gray Catbird Northern Mockingbird Sage Thrasher Bendire’s Thrasher Curve-billed Thrasher California Thrasher Crissal Thrasher Le Conte’s Thrasher Water Pipit Bohemian Waxwing Cedar Waxwing Phainopepla Northern Shrike Loggerhead Shrike European Starling Bell’s Vireo Gray Vireo Solitary Vireo Hutton’s Vireo Warbling Vireo Orange-crowned Warbler Nashville Warbler Virginia’s Warbler Lucy’s Warbler Yellow Warbler Yellow-rumped Warbler Black-throated Gray Warbler Townsend’s Warbler Hermit Warbler Grace’s Warbler Ovenbird MacGillivray’s Warbler Common Yellowthroat Wilson’s Warbler Red-faced Warbler Painted Redstart Yellow-breasted Chat Olive Warbler Hepatic Tanager X X X X XX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X XX X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X X (Continued) 573 Relict conifer forests, madrean evergreen woodlands Table 5—Birds occurring in southwestern and western nonforest Bird Species Summer Tanager Western Tanager Northern Cardinal Pyrrhuloxia Rose-breasted Grosbeak Black-headed Grosbeak Blue Grosbeak Lazuli Bunting Indigo Bunting Varied Bunting Green-tailed Towhee Rufous-sided Towhee Brown Towhee Abert’s Towhee Botteri’s Sparrow Cassin’s Sparrow Rufous-winged Sparrow Rufous-crowned Sparrow American Tree Sparrow Chipping Sparrow Clay-colored Sparrow Brewer's Sparrow Black-chinned Sparrow Vesper Sparrow Lark Sparrow Black-throated Sparrow Sage Sparrow Lark Bunting Savannah Sparrow Grasshopper Sparrow Fox Sparrow Song Sparrow Lincoln’s Sparrow Swamp Sparrow White-throated Sparrow Golden-crowned Sparrow White-crowned Sparrow Harris’ Sparrow Dark-eyed Junco Yellow-eyed Junco habitats—continued k_ O 03 0 SZ U 03 k_ 03 "O c C 03 J? 03 .9 E 03 03 V— 03 0 S w 0 Q Q- "o c k_ 03 0 03 0 > u. — 03 0 TD 3 03 O C O 0 O c a s < X X X X X X X X X X X X X XXX X X XXX X XXX XXX X X X X X X X XXX X XXX X X X XXX XXX X 5-S •2 E !c *“ Q. <5 > -O be ot X X X X X X X X X X X X X X X X X X X X X X X X X X X 0 03 C Q. 03 0 c 0 0 0 o 03 ~o c 0 0 0 0 O .E | S o C o o 5 l 0 — ■6 c c *o 3 0 o O 0 *0 ^ E cr eJ X X X X X X X X X X X X (Continued) 574 Table 5 —Birds occurring in southwestern and western nonforest habitats—continued iird Species ® 3 Q- *- CL n O a) ■§ w w tr S « 8 t r ® “ $ Q C CO c n 3 O 3 P (0 O CD a) jz "O (/) c S E ,9-*u *- c tr — CD "O _ CD 5 =J * CO CL ~ o -= c -o CD 5 3 CO F > O CD < cc « 2 E yicCown’s Longspur .apland Longspur Chestnut-collared Longspur Bobolink Bed-winged Blackbird fricolored Blackbird Eastern Meadowlark A/estern Meadowlark fellow-headed Blackbird Busty Blackbird 3rewer’s Blackbird Sreat-tailed Grackle Common Grackle 3ronzed Cowbird 3rown-headed Cowbird Hooded Oriole Northern Oriole Scott’s Oriole Rosy Finch 3 ine Grosbeak D urple Finch Cassin’s Finch House Finch Red Crossbill Common Redpoll Pine Siskin Lesser Goldfinch Lawrence’s Goldfinch American Goldfinch Evening Grosbeak House Sparrow X X X X X X X X X X X X X X X X X X X XXX X X X X X X X X X X X X X X X X X X X X X X X X X X X XXX X X XXX X X X XXX X X X X XXX X XXX X X X X X XXX X X X X X X X XXX X X XXX X X X X X X X X X X X X X X X 575 Relict conifer forests, madrean evergreen woodlands LITERATURE CITED Aber, J.D. 1979. 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Auk. 99:292-298. 611 Index to Birds and Families Family Gaviidae Common Loon Gavia immer . 13 Family Podicipedidae Pied-billed Grebe Podilymbus podiceps .14 Horned Grebe Podiceps auritus .15 Red-necked Grebe Podiceps grisegena .16 Eared Grebe Podiceps nigricollis .17 Western Grebe Aechmophorus occidentalis .18 Family Pelecanidae American White Pelican Pelecanus erythrorhynchos .19 Family Phalacrocoracidae Double-crested Cormorant Phalacrocorax auritus .20 Family Anhingidae Anhinga Anhinga anhinga .21 Family Ardeidae American Bittern Botaurus lentiginosus .22 Least Bittern Ixobrychus exilis .23 Great Blue Heron Ardea herodias .24 Great Egret Casmerodius albus .25 Snowy Egret Egretta thula .26 Little Blue Heron Egretta caerulea .27 Cattle Egret Bubulcus ibis .28 Green-backed Heron Butorides striatus .29 Black-crowned Night-Heron Nycticorax nycticorax .30 Yellow-crowned Night-Heron Nycticorax violaceus .31 Family Threskiornithidae Glossy Ibis Plegadis falcinellus .32 White-faced Ibis Plegadis chihi .33 Family Anatidae Fulvous Whistling-Duck Dendrocygna bicolor .34 Black-bellied Whistling-Duck Dendrocygna autumnalis .35 Tundra Swan Cygnus columbianus .36 Trumpeter Swan Cygnus buccinator .37 Greater White-fronted Goose Anser albifrons .38 Snow Goose Chen caerulescens .39 Ross’ Goose Chen rossii .40 Canada Goose Branta canadensis .41 Wood Duck Aix sponsa .42 612 Green-winged Teal Anas crecca .43 American Black Duck Anas rubripes .44 Mottled Duck Anas fulvigula .45 Mallard Anas platyrhynchos .46 Northern Pintail Anas acuta .47 Blue-winged Teal Anas discors .48 Cinnamon Teal Anas cyanoptera .49 Northern Shoveler Anas clypeata .50 Gadwall Anas strepera .51 American Wigeon Anas americana .52 Canvasback Aythya valisineria .53 Redhead Aythya americana .54 Ring-necked Duck Aythya collaris .55 Greater Scaup Aythya marila .56 Lesser Scaup Aythya affinis .57 Harlequin Duck Histrionicus histrionicus .58 White-winged Scoter Melanitta fusca .59 Common Goldeneye Bucephala clangula .60 Barrow’s Goldeneye Bucephala islandica .61 Bufflehead Bucephala albeola .62 Hooded Merganser Lophodytes cucullatus .63 Common Merganser Mergus merganser .64 Red-breasted Merganser Mergus serrator .65 Ruddy Duck Oxyura jamaicensis .66 Family Cathartidae Black Vulture Coragyps atratus .67 Turkey Vulture Cathartes aura .68 California Condor Gymnogyps californianus .69 Family Accipitridae Osprey Pandion haliaetus .70 American Swallow-tailed Kite Elanoides forficatus .71 Black-shouldered Kite Elanus caeruleus .72 Snail Kite Rostrhamus sociabilis .73 Mississippi Kite Ictinia mississippiensis .74 Bald Eagle Haliaeetus leucocephalus .75 Northern Harrier Circus cyaneus .76 Sharp-shinned Hawk Accipiter striatus .77 Cooper’s Hawk Accipiter cooperii .78 Northern Goshawk Accipiter gentilis .79 Common Black-Hawk Buteogallus anthracinus .80 Harris’ Hawk Parabuteo unicinctus .81 Gray Hawk Buteo nitidus .82 Red-shouldered Hawk Buteo lineatus .83 Broad-winged Hawk Buteo platypterus .84 Short-tailed Hawk Buteo brachyurus .85 613 Swainson’s Hawk Buteo swainsoni .86 White-tailed Hawk Buteo albicaudatus .87 Zone-tailed Hawk Buteo albonotatus .88 Red-tailed Hawk Buteo jamaicensis .89 Ferruginous Hawk Buteo regalis .90 Rough-legged Hawk Buteo lagopus . 91 Golden Eagle Aquila chrysaetos .92 Family Falconidae American Kestrel Falco sparverius . 93 Merlin Falco columbarius . 94 Peregrine Falcon Falco peregrinus . 95 Prairie Falcon Falco mexicanus .96 Family Cracidae Plain Chachalaca Ortalis vetula . 97 Family Phasianidae Gray Partridge Perdix perdix .98 Chukar Alectoris chukar . 99 Ring-necked Pheasant Phasianus colchicus .100 Spruce Grouse Dendragapus canadensis .101 Blue Grouse Dendragapus obscurus .102 White-tailed Ptarmigan Lagopus leucurus .103 Ruffed Grouse Bonasa umbellus .104 Sage Grouse Centrocercus urophasianus .105 Greater Prairie-Chicken Tympanuchus cupido .106 Lesser Prairie-Chicken Tympanuchus pallidicinctus .107 Sharp-tailed Grouse Tympanuchus phasianellus .108 Wild Turkey Meleagris gallopavo .109 Montezuma Quail Cyrtonyx montezumae .110 Northern Bobwhite Colinus virginianus .Ill Scaled Quail Callipepla squamata .112 Gambel’s Quail Callipepla gambelii . 113 California Quail Callipepla californica .114 Mountain Quail Oreortyx pictus .115 Family Rallidae Yellow Rail Coturnicops noveboracensis .116 Black Rail Laterallus jamaicensis .117 Clapper Rail Rallus longirostris .118 King Rail Rallus elegans .119 Virginia Rail Rallus limicola .120 Sora Porzana Carolina .121 Purple Gallinule Porphyrula martinica .122 Common Moorhen Gallinula chloropus .123 American Coot Fulica americana .124 614 Family Aramidae Limpkin Aramus guarauna .125 Family Gruidae Sandhill Crane Grus canadensis .126 Whooping Crane Grus americana .127 Family Charadriidae Black-bellied Plover Pluvialis squatarola .128 Lesser Golden-Plover Pluvialis dominica .129 Snowy Plover Charadrius alexandrinus .130 Semipalmated Plover Charadrius semipalmatus .131 Piping Plover Charadrius melodus .132 Killdeer Charadrius vociferus .133 Mountain Plover Charadrius montanus .134 Family Recurvirostridae Black-necked Stilt Himantopus mexicanus .135 American Avocet Recurvirostra americana .136 Family Scolopacidae Greater Yellowlegs Tringa melanoleuca .137 Lesser Yellowlegs Tringa flavipes .138 Solitary Sandpiper Tringa solitaria .139 Willet Catoptrophorus semipalmatus .140 Spotted Sandpiper Actitis macularia .141 Upland Sandpiper Bartramia longicauda .142 Long-billed Curlew Numenius americanus .143 Hudsonian Godwit Limosa haemastica .144 Marbled Godwit Limosa fedoa .145 Ruddy Turnstone Arenaria interpres .146 Red Knot Calidris canutus .147 Sanderling Calidris alba .148 Semipalmated Sandpiper Calidris pusilla .149 Western Sandpiper Calidris mauri .150 Least Sandpiper Calidris minutilla .151 White-rumped Sandpiper Calidris fuscicollis .152 Baird’s Sandpiper Calidris bairdii .153 Pectoral Sandpiper Calidris melanotos .154 Stilt Sandpiper Calidris himantopus .155 Buff-breasted Sandpiper Tryngites subruficollis .156 Short-billed Dowitcher Limnodromus griseus .157 Long-billed Dowitcher Limnodromus scolopaceus .158 Common Snipe Gallinago gallinago .159 American Woodcock Scolopax minor .160 Wilson’s Phalarope Phalaropus tricolor .161 Red-necked Phalarope Phalaropus lobatus .162 615 Family Laridae Franklin’s Gull Larus pipixcan .163 Bonaparte’s Gull Larus Philadelphia .164 Ring-billed Gull Larus delawarensis .165 California Gull Larus californicus .166 Herring Gull Larus argentatus .167 Caspian Tern Sterna caspia .168 Common Tern Sterna hirundo .169 Forster’s Tern Sterna forsteri . 170 Least Tern Sterna antillarum . 171 Black Tern Chlidonias niger . 172 Family Alcidae Marbled Murrelet Brachyramphus marmoratus . 173 Family Columbidae Rock Dove Columba livia . 174 White-crowned Pigeon Columba leucocephala . 175 Red-billed Pigeon Columba flavirostris .176 Band-tailed Pigeon Columba fasciata . 177 White-winged Dove Zenaida asiatica .178 Mourning Dove Zenaida macroura . 179 Inca Dove Columbina inca .180 Common Ground-Dove Columbina passerina .181 Family Cuculidae Black-billed Cuckoo Coccyzus erythropthalmus .182 Yellow-billed Cuckoo Coccyzus americanus .183 Mangrove Cuckoo Coccyzus minor .184 Greater Roadrunner Geococcyx californianus .185 Smooth-billed Ani Crotophaga ani .186 Groove-billed Ani Crotophaga sulcirostris .187 Family Tytonidae Common Barn-Owl Tyto alba . 188 Family Strigidae Flammulated Owl Otus flammeolus .189 Eastern Screech-Owl Otus asio . 190 Western Screech-Owl Otus kennicottii .190 Whiskered Screech-Owl Otus trichopsis . 191 Great Horned Owl Bubo virginianus .192 Snowy Owl Nyctea scandiaca . 193 Northern Hawk-owl Surnia ulula . 194 Northern Pygmy-Owl Glaucidium gnoma . 195 Ferruginous Pygmy-Owl Glaucidium brasilianum .196 Elf Owl Micrathene whitneyi .197 Burrowing Owl Athene cunicularia .198 Spotted Owl strix occidentalis .199 Barred Owl strix varia .200 616 Great Gray Owl Strix nebulosa .201 Long-eared Owl Asio otus .202 Short-eared Owl Asio flammeus .203 Boreal Owl Aegolius funereus .204 Northern Saw-whet Owl Aegolius acadicus .205 Family Caprimulgidae Lesser Nighthawk Chordeiles acutipennis .206 Common Nighthawk Chordeiles minor .207 Common Pauraque Nyctidromus albicollis .208 Common Poorwill Phalaenoptilus nuttallii .209 Chuck-will’s-widow Caprimulgus carolinensis .210 Whip-poor-will Caprimulgus vociferus .211 Family Apodidae Black Swift Cypseloides niger .212 Chimney Swift Chaetura pelagica .213 Vaux’s Swift Chaetura vauxi .214 White-throated Swift Aeronautes saxatalis .215 Family Trochilidae Broad-billed Hummingbird Cynanthus latirostris .216 White-eared Hummingbird Hylocharis leucotis .217 Buff-bellied Hummingbird Amazilia yucatanensis .218 Violet-crowned Hummingbird Amazilia violiceps .219 Blue-throated Hummingbird Lampornis clemenciae .220 Magnificent Hummingbird Eugenes fulgens .221 Lucifer Hummingbird Calothorax lucifer .222 Ruby-throated Hummingbird Archilochus colubris .223 Black-chinned Hummingbird Archilochus alexandri .224 Anna’s Hummingbird Calypte anna .225 Costa’s Hummingbird Calypte costae .226 Calliope Hummingbird Stellula calliope .227 Broad-tailed Hummingbird Selasphorus platycercus .228 Rufous Hummingbird Selasphorus rufus .229 Allen’s Hummingbird Selasphorus sasin .230 Family Trogonidae Elegant Trogon Trogon elegans .231 Family Alcedinidae Belted Kingfisher Ceryle alcyon .232 Green Kingfisher Chloroceryle americana .233 Family Picidae Lewis’ Woodpecker Melanerpes lewis .234 Red-headed Woodpecker Melanerpes erythrocephalus .235 Acorn Woodpecker Melanerpes formicivorus .236 617 Gila Woodpecker Melanerpes uropygialis .235 Golden-fronted Woodpecker Melanerpes aurifrons . 231 Red-bellied Woodpecker Melanerpes carolinus .235 Yellow-bellied Sapsucker Sphyrapicus varius .24( Red-breasted Sapsucker Sphyrapicus ruber .24' Williamson’s Sapsucker Sphyrapicus thyroideus . 24c Ladder-backed Woodpecker Picoides scalaris .245 Nuttall’s Woodpecker Picoides nuttallii .24' Downy Woodpecker Picoides pubescens .245 Hairy Woodpecker Picoides villosus .24( Strickland’s Woodpecker Picoides stricklandi .245 Red-cockaded Woodpecker Picoides borealis .24 i White-headed Woodpecker Picoides albolarvatus .245 Three-toed Woodpecker Picoides tridactylus .25( Black-backed Woodpecker Picoides arcticus .25' Northern Flicker Colaptes auratus .255 Pileated Woodpecker Dryocopus pileatus .255 Ivory-billed Woodpecker Campephiius principalis .25' Family Tyrannidae Northern Beardless-Tyrannulet Camptostoma imberbe .255 Olive-sided Flycatcher Contopus borealis .255 Greater Pewee Contopus pertinax .255 Western Wood-Pewee Contopus sordidulus .255 Eastern Wood-Pewee Contopus virens .255 Yellow-bellied Flycatcher Empidonax flaviventris .26( Acadian Flycatcher Empidonax virescens .26' Alder Flycatcher Empidonax alnorum .265 Willow Flycatcher Empidonax traillii .265 Least Flycatcher Empidonax minimus .26' Hammond’s Flycatcher Empidonax hammondii .265 Dusky Flycatcher Empidonax oberholseri .265 Gray Flycatcher Empidonax wrightii .265 Western Flycatcher Empidonax difficilis .265 Buff-breasted Flycatcher Empidonax fulvifrons .265 Black Phoebe Sayornis nigricans .27C Eastern Phoebe Sayornis phoebe .271 Say’s Phoebe Sayornis saya .275 Vermilion Flycatcher Pyrocephalus rubinus .275 Dusky-capped Flycatcher Myiarchus tuberculifer .27^ Ash-throated Flycatcher Myiarchus cinerascens .275 Great Crested Flycatcher Myiarchus crinitus .276 Brown-crested Flycatcher Myiarchus tyrannulus .277 Great Kiskadee Pitangus sulphuratus .276 Sulphur-bellied Flycatcher Myiodynastes luteiventris .27S Tropical Kingbird Tyrannus melancholicus .28C Couch’s Kingbird Tyrannus couchii .281 618 Cassin’s Kingbird Tyrannus vociferans .282 Thick-billed Kingbird Tyrannus crassirostris .283 Western Kingbird Tyrannus verticalis .284 Eastern Kingbird Tyrannus tyrannus .285 Gray Kingbird Tyrannus dominicensis .286 Scissor-tailed Flycatcher Tyrannus forficatus .287 Rose-throated Becard Pachyramphus aglaiae .288 Family Alaudidae Horned Lark Eremophila alpestris .289 Family Hirundinidae Purple Martin Progne subis .290 Tree Swallow Tachycineta bicolor .291 Violet-green Swallow Tachycineta thalassina .292 Northern Rough-winged Swallow Stelgidopteryx serripennis .293 Bank Swallow Riparia riparia .294 Cliff Swallow Hirundo pyrrhonota .295 Cave Swallow Hirundo fulva .296 Barn Swallow Hirundo rustica .297 Family Corvidae Gray Jay Perisoreus canadensis .298 Steller’s Jay Cyanocitta stelleri .299 Blue Jay Cyanocitta cristata .300 Green Jay Cyanocorax yncas .301 Scrub Jay Aphelocoma coerulescens .302 Gray-breasted Jay Aphelocoma ultramarina .303 Pinyon Jay Gymnorhinus cyanocephalus .304 Clark’s Nutcracker Nucifraga Columbiana .305 Black-billed Magpie Pica pica .306 Yellow-billed Magpie Pica nuttalli .307 American Crow Corvus brachyrhynchos .308 Northwestern Crow Corvus caurinus .309 Fish Crow Corvus ossifragus .310 Chihuahuan Raven Corvus cryptoleucus .311 Common Raven Corvus corax .312 Family Paridae Black-capped Chickadee Parus atricapillus .313 Carolina Chickadee Parus carolinensis .314 Mexican Chickadee Parus sclateri .315 Mountain Chickadee Parus gambeli .316 Siberian Tit Parus cinctus .317 Boreal Chickadee Parus hudsonicus .318 619 Chestnut-backed Chickadee Parus rufescens .319 Bridled Titmouse Parus wollweberi .320 Plain Titmouse Parus inornatus .321 Tufted Titmouse Parus bicolor .322 Family Remizidae Verdin Auriparus flaviceps .323 Family Aegithalidae Bushtit Psaltriparus minimus .324 Family Sittidae Red-breasted Nuthatch Sitta canadensis .325 White-breasted Nuthatch Sitta carolinensis .326 Pygmy Nuthatch Sitta pygmaea .327 Brown-headed Nuthatch Sitta pusilla .328 Family Certhiidae Brown Creeper Certhia americana .329 Family Pycnonotidae Red-whiskered Bulbul Pycnonotus jocosus .330 Family Troglodytidae Cactus Wren Campylorhynchus brunneicapillus ... 331 Rock Wren Salpinctes obsoletus .332 Canyon Wren Catherpes mexicanus .333 Carolina Wren Thryothorus ludovicianus .334 Bewick’s Wren Thryomanes bewickii .335 House Wren Troglodytes aedon .336 Winter Wren Troglodytes troglodytes .337 Sedge Wren Cistothorus platensis .338 Marsh Wren Cistothorus palustris .339 Family Cinclidae American Dipper Cinclus mexicanus .340 Family Muscicapidae Golden-crowned Kinglet Regulus satrapa .341 Ruby-crowned Kinglet Regulus calendula .342 Blue-gray Gnatcatcher Polioptila caerulea .343 Black-tailed Gnatcatcher Polioptila melanura .344 Eastern Bluebird Sialia sialis .345 Western Bluebird Sialia mexicana .346 Mountain Bluebird Sialia currucoides .347 Townsend’s Solitaire Myadestes townsendi .348 Veery Catharus fuscescens .349 620 Gray-cheeked Thrush Catharus minimus .350 Swainson’s Thrush Catharus ustulatus .351 Hermit Thrush Catharus guttatus .352 Wood Thrush Hylocichla mustelina .353 American Robin Turdus migratorius .354 Varied Thrush Ixoreus naevius .355 Wrentit Chamaea fasciata .356 Family Mimidae Gray Catbird Dumetella carolinensis .357 Northern Mockingbird Mimus polyglottos .358 Sage Thrasher Oreoscoptes montanus .359 Brown Thrasher Toxostoma rufum .360 Long-billed Thrasher Toxostoma longirostre .361 Bendire’s Thrasher Toxostoma bendirei .362 Curve-billed Thrasher Toxostoma curvirostre .363 California Thrasher Toxostoma redivivum .364 Crissal Thrasher Toxostoma dorsale .365 Le Conte’s Thrasher Toxostoma lecontei .366 Family Motacillidae Water Pipit Anthus spinoletta .367 Sprague’s Pipit Anthus spragueii .368 Family Bombycillidae Bohemian Waxwing Bombycilla garrulus .369 Cedar Waxwing Bombycilla cedrorum .370 Family Ptilogonatidae Phainopepla Phainopepla nitens .371 Family Laniidae Northern Shrike Lanius excubitor .372 Loggerhead Shrike Lanius ludovicianus .373 Family Sturnidae European Starling Sturnus vulgaris .374 Family Vireonidae White-eyed Vireo Vireo griseus .375 Bell’s Vireo Vireo bellii .376 Black-capped Vireo Vireo atricapillus .377 Gray Vireo Vireo vicinior .378 Solitary Vireo Vireo solitarius .379 Yellow-throated Vireo Vireo flavifrons .380 Hutton’s Vireo Vireo huttoni .381 Warbling Vireo Vireo gilvus .382 621 Philadelphia Vireo Vireo philadelphicus .383 Red-eyed Vireo Vireo olivaceus .384 Black-whiskered Vireo Vireo altiloquus .385 Family Emberizidae Bachman’s Warbler Vermivora bachmanii .386 Blue-winged Warbler Vermivora pinus .387 Golden-winged Warbler Vermivora chrysoptera .388 Tennessee Warbler Vermivora peregrina .389 Orange-crowned Warbler Vermivora celata .390 Nashville Warbler Vermivora ruficapilla .391 Virginia’s Warbler Vermivora virginiae .392 Colima Warbler Vermivora crissalis .393 Lucy’s Warbler Vermivora luciae .394 Northern Parula Parula americana .395 Tropical Parula Parula pitiayumi .396 Yellow Warbler Dendroica petechia .397 Chestnut-sided Warbler Dendroica pensylvanica .398 Magnolia Warbler Dendroica magnolia .399 Cape May Warbler Dendroica tigrina .400 Black-throated Blue Warbler Dendroica caerulescens .401 Yellow-rumped Warbler Dendroica coronata .402 Black-throated Gray Warbler Dendroica nigrescens .403 Townsend’s Warbler Dendroica townsendi . .404 Hermit Warbler Dendroica occidentalis .405 Black-throated Green Warbler Dendroica virens .406 Golden-cheeked Warbler Dendroica chrysoparia .407 Blackburnian Warbler Dendroica fusca .408 Yellow-throated Warbler Dendroica dominica .409 Grace’s Warbler Dendroica graciae .410 Pine Warbler Dendroica pinus .411 Kirtland’s Warbler Dendroica kirtlandii .412 Prairie Warbler Dendroica discolor .413 Palm Warbler Dendroica palmarum .414 Bay-breasted Warbler Dendroica castanea .415 Blackpoll Warbler Dendroica striata .416 Cerulean Warbler Dendroica cerulea .417 Black-and-white Warbler Mniotilta varia .418 American Redstart Setophaga ruticilla .419 Prothonotary Warbler Protonotaria citrea .420 Worm-eating Warbler Helmitheros vermivorus .421 Swainson’s Warbler Limnothlypis swainsonii .422 Oven bird Seiurus aurocapillus .423 Northern Waterthrush Seiurus noveboracensis .424 Louisiana Waterthrush Seiurus motacilla .425 Kentucky Warbler Oporornis formosus .426 Connecticut Warbler Oporornis agilis .427 622 Mourning Warbler Oporornis Philadelphia .428 MacGillivray’s Warbler Oporornis tolmiei .429 Common Yellowthroat Geothlypis trichas .430 Hooded Warbler Wilsonia citrina .431 Wilson’s Warbler Wilsonia pusilla .432 Canada Warbler Wilsonia canadensis .433 Red-faced Warbler Cardellina rubrifrons .434 Painted Redstart Myioborus pictus .435 Yellow-breasted Chat Icteria virens .436 Olive Warbler Peucedramus taeniatus .437 Hepatic Tanager Piranga flava .438 Summer Tanager Piranga rubra .439 Scarlet Tanager Piranga olivacea .440 Western Tanager Piranga ludoviciana .441 Northern Cardinal Cardinalis cardinalis .442 Pyrrhuloxia Cardinalis sinuatus .443 Rose-breasted Grosbeak Pheucticus ludovicianus .444 Black-headed Grosbeak Pheucticus melanocephalus .445 Blue Grosbeak Guiraca caerulea .446 Lazuli Bunting Passerina amoena .447 Indigo Bunting Passerina cyanea .448 Varied Bunting Passerina versicolor .449 Painted Bunting Passerina ciris .450 Dickcissel Spiza americana .451 Olive Sparrow Arremonops rufivirgatus .452 Green-tailed Towhee Pipilo chlorurus .453 Rufous-sided Towhee Pipilo erythrophthalmus .454 Brown Towhee Pipilo fuscus .455 Abert’s Towhee Pipilo aberti .456 White-collared Seedeater Sporophila torqueola .457 Bachman’s Sparrow Aimophila aestivalis .458 Botteri’s Sparrow Aimophila botterii .459 Cassin’s Sparrow Aimophila cassinii .460 Rufous-winged Sparrow Aimophila carpalis .461 Rufous-crowned Sparrow Aimophila ruficeps .462 American Tree Sparrow Spizella arborea .463 Chipping Sparrow Spizella passerina .464 Clay-colored Sparrow Spizella pallida .465 Brewer’s Sparrow Spizella breweri .466 Field Sparrow Spizella pusilla .467 Black-chinned Sparrow Spizella atrogularis .468 Vesper Sparrow Pooecetes gramineus .469 Lark Sparrow Chondestes grammacus .470 Black-throated Sparrow Amphispiza bilineata .471 Sage Sparrow Amphispiza belli .472 Lark Bunting Calamospiza melanocorys .473 Savannah Sparrow Passerculus sandwichensis .474 623 Baird’s Sparrow Ammodramus bairdii .475 Grasshopper Sparrow Ammodramus savannarum .476 Henslow’s Sparrow Ammodramus henslowii .477 Le Conte’s Sparrow Ammodramus leconteii .478 Sharp-tailed Sparrow Ammodramus caudacutus .479 Seaside Sparrow Ammodramus maritimus .480 Fox Sparrow Passerella iliaca .481 Song Sparrow Melospiza melodia .482 Lincoln’s Sparrow Melospiza lincolnii .483 Swamp Sparrow Melospiza georgiana .484 White-throated Sparrow Zonotrichia albicollis .485 Golden-crowned Sparrow Zonotrichia atricapilla .486 White-crowned Sparrow Zonotrichia leucophrys .487 Harris’ Sparrow Zonotrichia querula .488 Dark-eyed Junco Junco hyemalis .489 Yellow-eyed Junco Junco phaeonotus .490 McCown’s Longspur Calcarius mccownii .491 Lapland Longspur Calcarius lapponicus .492 Smith’s Longspur Calcarius pictus .493 Chestnut-collared Longspur Calcarius ornatus .494 Snow Bunting Plectrophenax nivalis .495 Bobolink Dolichonyx oryzivorus .496 Red-winged Blackbird Agelaius phoeniceus .497 Tricolored Blackbird Agelaius tricolor .498 Eastern Meadowlark Sturnella magna .499 Western Meadowlark Sturnella neglecta .500 Yellow-headed Blackbird Xanthocephalus xanthocephalus ....501 Rusty Blackbird Euphagus carolinus .502 Brewer’s Blackbird Euphagus cyanocephalus .503 Great-tailed Grackle Quiscalus mexicanus .504 Boat-tailed Grackle Quiscalus major .505 Common Grackle Quiscalus quiscula .506 Bronzed Cowbird Molothrus aeneus .507 Brown-headed Cowbird Molothrus ater .508 Orchard Oriole Icterus spurius .509 Hooded Oriole Icterus cucullatus .510 Altamira Oriole Icterus gularis .511 Audubon’s Oriole Icterus graduacauda .512 Northern Oriole Icterus galbula .513 Scott’s Oriole Icterus parisorum .514 Family Fringillidae Rosy Finch Leucosticte arctoa .515 Pine Grosbeak Pinicola enucleator .516 Purple Finch Carpodacus purpureus .517 Cassin’s Finch Carpodacus cassinii .518 House Finch Carpodacus mexicanus .519 624 Red Crossbill Loxia curvirostra .520 White-winged Crossbill Loxia leucoptera .521 Common Redpoll Carduelis flammea .522 Hoary Redpoll Carduelis hornemanni .523 Pine Siskin Carduelis pinus .524 Lesser Goldfinch Carduelis psaltria .525 Lawrence’s Goldfinch Carduelis lawrencei .526 American Goldfinch Carduelis tristis .527 Evening Grosbeak Coccothraustes vespertinus .528 Family Passeridae House Sparrow Passer domesticus .529 Eurasian Tree Sparrow Passer montanus .530 625 990 Agricultural Chartbook It’s Easy To Order Another Copy! Just dial 1-800-999-6779. Toll free (in the United States and Canada). All other areas please dial 301 - 725 - 7937 . Ask for 1990 Agricultural Chartbook (AH-689) The cost is $11.00 per copy. For non-U.S. addresses (including Canada), add 25 percent. Charge your purchase to your VISA or MasterCard, or we can bill you. Or send a check or purchase order (made payable to ERS-NASS) to: ERS-NASS P.O. Box 1608 Rockville, MD 20849-1608. We’ll fill your order by first-class mail. jUN 1 1990 C *3 FEDERAL DEPOSITORY Washington, DC 20005-4788 April 1990 intents 4 6 12 16 21 25 3 Economic Picture of U.S. Agriculture International and National Economic Outlook Farm Financial Conditions Crop Production Agricultural Trade Meat, Livestock, and Dairy Production Food Prices 26 Background Charts for U.S. Agriculture 27 28 29 30 33 34 35 36 37 39 40 41 42 43 44 45 47 48 49 51 53 54 55 56 58 59 60 61 62 63 Farm Population Farm Labor Regional Farm Structure Income Farm Debt Food and Fiber System Agriculture and the General Economy Assets and Finance Finance and Inputs Inputs Costs and Returns Land Use Land Values Land Use Changes Wetlands Irrigation Timber Products Forests and Timber Conservation Farmer Cooperatives Population Personal Income Poverty Employment Food and Fiber System Employment Nonmetro Versus Metro Employment Industry Employment Farm Credit Banking Earnings 64 Economic Indicators 66 Consumer Prices 67 Food Marketing Costs 68 Food Consumption 69 Women’s Diet 71 Family Economics 74 Food Assistance 75 Child Nutrition and Food Distribution 76 Producer Subsidy Equivalents 77 World and U.S. Trade 78 U.S. Trade 85 World Production 89 World Trade 93 Livestock 96 Dairy 97 Poultry 100 Aquaculture 101 Rice and Other Grains 102 Wheat 103 Coarse Grains 104 Soybeans 105 Fibers 107 Vegetables 108 Fruit 109 Fruit and Tree Nuts 110 Tree Nuts and Tropical Products 111 Sugar 113 Tobacco 115 Transportation 117 Index 1990 Agricultural Chartbook Committee Economics Management Staff Sara L. Wampler, Editor Susan DeGeorge, Graphic Coordinator Data Services Center Barbara Barnes Economic Research Service Bill Crosswhite, Wendell Holmes, Sara Mazie, Steve Milmoe, Ed Overton, Don Seaborg, Jewell Tolliver Agricultural Cooperative Service Celestine Adams Agricultural Research Service Joan C. Courtless Foreign Agricultural Service Donald Washington Forest Service Robert B. Phelps Food and Nutrition Service John Braden Human Nutrition Information Service Gerald Smith Economic Picture of U.S. Agriculture Assistant Secretary of Agriculture for Economics summarizes Urban 25.003 6,131 319 ft 530 706 303 230 Cropland and range account tof 60% of the urban increase. Very little land is converted back to a resource use once urbanized Changes in major land uses in fast-growth counties Percent Residential 50 Transportation 33 High-value crops 20 Mixed urban 19 Forest Cropland □ Range Range 24,965 Wetland - 2.8 -3.9 -4.8 - 6.1 L -10 0 1970-80 ERS data on land use change. 10 20 30 40 50 43 Wetland restoration costs (which are shared between Government and the landowner) stay rela- y , TTVL * 11 , 5 ^ 52 per acre as rs$e ™ solely by the Federal Government) rise from $135-$515per acre as reserve size increases About 65 million acres of existing wetlands remained in private ownership in 1982, much of that on farms 12 - 2million Nacres tbs, between ' Chart 60 Cropland on former wetland soils, 1982 Thousand acres l~ 1 0-99 HHlOO-999 1,000-2,999 ] 3,000-4.999 5,000-9.000 Chart 61 Costs of an agricultural wetland reserve, existing and former wetland $ billion ® r Cropland on former wetland soil 2 - Existing wetland Easement costs Restoration costs Nonfederal rural land. National Resources Inventory. 1990 data, estimated. Chart 62 Remaining wetlands, 1982 Chart 63 Wetland losses, United States: Mid-1950 s to mid-1970 s Agriculture 10.6% Urban 1.0% Other 0.6% Nonfederal rural land. National Resources Inventory. The pie represents 108 million acres of wetland existing in the rrtd-1950's Irrigation Irriqated crops accounted for $26 billion of total crop value. Irrigated acres by region vary annually due to changes in crop and input prices, commodity programs, climatic factors, and techno ogy adoption. The West and South are much more dependent upon irrigation than other parts of the United States. Chart 64 Irrigation and farm production Irrigation 4.8% Land in farms 964 million acres Irrigation 14.8% Harvested cropland 282 million acres Irrigation 37.8% Crop value $68.8 billion 1987 data. Source: Census of Agriculture. Chart 65 Irrigated acres by production region Chart 66 Irrigated harvested cropland as a percent of total harvested cropland Million acres Data tor 1982 and 1987 are from the Census of Agriculture. Data for other years are ERS-USDA estimates. Percent 0-1 1-5 5-20 20-60 60-100 1987 data. Source: Census of Agriculture 45 Irrigation The percentage of harvested cropland irrigated varies by crop. Differences can be due to geographic concentration of production and variations in cultural practices. The Midwest and South rely most heavi/y on groundwater for irrigation. Nearly all declines in irrigated acreage between 1982 and 1987 occurred in the Western States. * Chart 67 Percent of harvested cropland irrigated Chart 68 Percent of irrigation water from groundwater sources Rice Orchards Irish potatoes Vegetables Sugarcane Sugar beets Dry beans Cotton Peanuts Hay Barley Corn tor silage Corn for grain Grain sorghum Wheat Soybeans All cropland harvested 0 20 40 60 80 100 Percent 1985 data. Source: U.S. Geological Survey Chart 69 Change in irrigated acres, 1982-87 Percent More than 25 10 to 25 0 to 10 -1 to -10 M Less than -10 Not applicable Source: Census ol Agriculture. 46 Timber Products U S production of timber products rose in 1988 and exports rose to record levels. Consumption and -Vroorts declined for the first time since 1985. Imports supplied a Me over 23 percent of total consumption in 1988. About 18 percent of total production was exported. Chart 70 Timber products production Billion cubic feet Estimated volume of timber required to produce indicated products. Source: Based on Bureau of the Census data. Chart 71 Timber products consumption Billion cubic feet Estimated volume of timber required to produce indicated products. Source: Based on Bureau of the Census data. Chart 72 Timber products Imports Chart 73 Timber products exports Billion cubic feet 1960 65 70 75 80 85 Estimated volume of timber required to produce indicated products. Source: Based on Bureau of the Census data. Billion cubic feet 5 3 - 1 - 1960 65 Estimated volume of timber required to produce indicated products. Source: Based on Bureau of the Census data. 47 Forests and Timber Forests occupy approximately a third of the Nation's total land area. Two-thirds of the forest area is timberland, or productive forests, owned by the public, forest industries, farmers, and other in¬ dividuals and firms. In total, U. S. timberland contains 830 billion cubic feet of timber 57 percent softwood species and 43 percent hardwoods. Chari 74 Forest land areas of the United States 1987 data. Hawaii and Alaska included in Pacific Coast. Chart 75 Forest land areas of the United States, bv type, 1987 Million acres 250 200 150 100 50 o L North South -Other forest land Reserved timberland Timberland Rocky Mountains Pacific Timberland is forest land capable of producing 20 cubic feet per acre of industrial wood annually and not reserved from harvests. Chart 76 Timberland ownership In the United States, Million acres 200 150 100 50 0 —Other private Forest industry Farmer ublic North South Rocky Pacific Mountains Chart 77 Volume of timber on timberland in the United States, 1987 Billion cubic feet 300 r Hardwood species Softwood species North South Rocky Pacific Mountains Net volume, which is gross volume less deductions for rot. roughness, and poor 48 Conservation US DA and related State and local conservation program expenditures exceeded$3.2 billion in 1989 Over 60 percent of total USD A conservation expenditures are related to the Conservation Reserve Program (CRP). The Corn Belt and Northern Plains account for nearly 60 percent of all conservation-tilled acres. Almost 90 percent of the acres with conservation tillage produced corn, soybeans, or small grain in 1989. Chart 78 ... USDA and related State and local program conservation expenditures $ billion 3.5 3.0 2.5 2.0 1.5 1.0 .5 0 1983 ■ State and local USDA other conservation USDA CRP 86 87 88 89 Chart 79 USDA conservation expenditures, 1989 CRP cover, S546.8 Technical assistance and extension, $436 million Conservation data and research. $226.6 million Other cost-sharing, $218.8 million Project conservation, $197.5 million CRP rental payments. $1,192 million Total expenditures: $2,818 million Chart 80 Tillage practices on acres planted, by region Northeast Southeast Pacific Appalachian Delta States Mountain Southern Plains Lake States 35% | 8.1 15% |; 9.8 13% ’2 13.1 13.4 36% y 11% E 22 %^ 23% HH 19 % 14.9 20.4 28.2 31.3 Conservation tillage No tillage and ridge tillage Mulch tillage Conventional tillage 15-30% residue cover tillage 0-15% residue cover tillage Northern Plains 26% Corn Belt 0 10 20 30 40 50 60 70 80 90 Million acres 1989 data. Source: Conservation Technology Information Center. Percentages show total regional conservation tillage use as a percent of acres planted in the region. Numbers show total acres planted in the region. Chart 81 ... Total acres planted with conservation tillage by crop, 1989 Small grain, fall seeded 19.9% Small grain, spring seeded 11.4% Sorghum 5.1% Other 5.1% Soybeans, double crop 4.8% Conservation Excess production capacity peaked in 1986 and continues to decline. Conservation Reserve Pro¬ gram enrollment is approaching the 40-45 million acre goal. Enrollment is concentrated in the Plains and Mountain States, with lowest participation in the Northeast. Chart 82 Excess production capacity in 12 commodities Percent Chart 83 Total acreage enrolled in the Conservation Reserve Program as of February 1989 One dot equals 1.000 acres. Total enrollment through the eighth sionuo. 30 6 million flnrp o Chart 84 Conservation reserve acreage ps a percentage of eligible cropland Chart 85 Average annual per acre rental payment of conservation reserve lands by State Percent r I Under 10 m 10-24 HI 25-49 Over 49 Dollars [ I Under 40 ! 1 40-49 ^ 50-59 Over 59 1989 data. 1989 data 50 Farmer Cooperatives The decrease in number of U.S. farmer cooperatives in 1987 reflects a continuing trend of merger, consolidation, acquisition, and dissolution. The long-term decline in memberships reflects in part he decreasing number of U.S. farms and farmers. Farmer cooperative total assets increased in 1987 by 4.8 percent. Chart 86 Farmer cooperatives in the United States Thousand cooperatives services. Chart 88 U.S. farms and farmer cooperative memberships Millions Memberships include duplication that cannot be eliminated using current reporting methods. Chart 87 Mergers and dissolution of U.S. farmer cooperatives Number 300 r 1981 82 83 84 85 86 87 Based on a list of all U.S. farmer cooperatives maintained by the Agricultural Cooperative Service. Mergers also Include acquisitions and consolidations. Other includes cooperatives dropped due to inactivity, reassignment and mlscelaneous reasons. Chart 89 Distribution of farmer cooperatives by size of assets % of farmer cooperatives Assets ($ million) 1987 data. 51 Farmer Cooperatives Value of farm products marketed by farmer cooperatives increased 5.9 percent and supplies handled dropped 5.5 percent in 1987. The 24 cooperatives with gross business of $500 million or more accounted for a greater percentage (38.3 percent) of total gross dollar volume than the 4 606 cooperatives with business of less than $15 million (21.5 percent). Chart 90 Distribution of farmer cooperative business by size of cooperative % of business handled Size of cooperative based on gross sales. 25-99.9 100-499.9 $ million 1,000 and over Chart 91 Farm products marketed by farmer cooperatives 1987 data. Total net marketing business = $44.2 billion. Other products include dry edible beans and peas. nuts. rice, tobacco, and miscellaneous products. Chart 92 Farm supplies handled by farmer cooperatives 1987 data. Total net farm supply business = $14.3 billion. Other supplies include containers, meats and groceries, and miscellaneous farm supplies. 52 Population Vef outmigration from nonmetro counties became widespread in the mid-1980 s. Poverty is consis ently higher among nonmetro children, and is especially high in families where the mother is rais¬ ing the children by herself. Chart 93 Nonmetro counties with net outmigration, 1980-83 Chart 94 Nonmetro counties with net outmigration, 1983-88 Chart 95 Children under 18 years living with one parent Percent 1970 1980 1988 Chart 96 Children below poverty level, by family type and residence Percent 53 Personal Income Metro residents consistently have higher income than nonmetro residents. Transfers grew faster than earnings during 1970-87. Social Security, medicare/medicaid, and other retirement/disability programs account for over 80 percent of government transfers to metro and nonmetro residents. Chart 97 Trends in per capita income Chart 98 Personal income by source Thousand 1987 $ Source: Local Area Personal Income . U.S. Department of Commerce. Bureau of Economic Analysis. Percent 100 80 60 40 20 0 Metro Source: Local Area Personal Income . U.S. Department ol Commerce. Bureau of Economic Analysis. Chart 99 Transfer payments as percentage of total personal income Percent Source: Local Area Personal Income . U.S. Department of Commerce. Bureau of Economic Analysis. Chart 100 Government transfer payments to individuals Percent Social Security Medicare/medicaid Other retirement and disability Public assistance Veterans' benefits Unemployment compensation Other transfers 1987 data. Source: Local Area Personal Income. U.S. Department of Commerce. Bureau of Economic Analysis 54 Poverty The poverty rate is higher in nonmetro than metro areas. Farm poverty was very high in the early 1980’s, but has declined since then. Most poor families work for at least part of their income. Elderly blacks and young black children living in nonmetro areas have very high poverty rates. Chart 101 Poverty rates Chart 102 Poor families by sources of income % of residents The farm poverty rate is the proportion of all residents living on farms who are poor. Data for 1984 not available. Source: Current Population Survey. Bureau of the Census. % of poor families Earnings only Earnings and Social Security only Earnings and public assistance only Other combinations with earnings Social Security only Public assistance only Social Security and public assistance only Other combinations without earnings No income reported 2.6 1.8 8.3 9.0 33.0 24.8 5.0 4.9 10.5 19.5 3.2 1.5 14.2 16.9 2.4 1987 data. Source: Current Population Survey. Bureau of the Census. Chart 103 Percentage of the elderly in poverty Chart 104 Percentage of young children in poverty United States Nonmetro Metro central city Metro outside central city Percent United States Nonmetro Metro central city Metro outside central city Percent South Nonmetro Metro central city 21.3 47.4 19.2 34.6 Metro outside 11.1 central city 37 3 1987 data. The elderly are persons 65 and over. Source: Current Population Survey. Bureau of the Census. South Nonmetro Metro central city Metro outside central city 1987 data. Young children are under 6 and live in families. Source: Current Population Survey. Bureau of the Census. 55 Employment Nonmetro economies outside the Northeast had lower employment growth and higher unemploy¬ ment than metro areas between 1982 and 1988. Chart 105 Employment change from previous year Percent Source: Bureau of Labor Statistics. Chart 106 Unemployment rates for metro and nonmetro areas Percent Adjusted unemployment includes those unemployed, those not looking lor work because jobs are unavailable, and half of those working part-trne because full-time jobs are unavailable. Beginning third quarter 1985. CPS metro/nonmetro definition based on 1980 Census. Source Current Population Survey. Chart 107 Employment growth by region, 1982-86 Chart 108 Unemployment by region Percent 25 |PP Metro | Non metro Percent Northeast Midwest Source: Bureau of Labor Statistics. South West 12 9 6 3 0 Metro Non - metro Northeast Metro Non - metro Midwest Metro Non - Metro Non - metro metro South West Source: Bureau of Labor Statistics. 56 Employment Employment grew at a faster rate in nonmetro areas than metro areas in 1988, for the first time since the mid-1970’s. However, teenagers and minorities continue to experience relatively high levels of unemployment. Nonmetro employment growth rates were above 15.4 percent in 13 States between 1982 and 1988, while the nonmetro unemployment rate was over 1.5 times the U.S. average in 11 States. Chari 109 Nonmetro unemployment by race/ethnlcity Percent Beginning third quarter 1985. CPS nonmetro definition based on 1980 Census. Source: Current Population Survey. Chart 110 Nonmetro unemployment by age Percent Chart 111 Nonmetro unemployment by State, 1988 Chart 112 Nonmetro employment growth by State, 1982-88 U.S. average (5.5%) or less 1-1.5 times U.S. average Over 1.5 times U.S. average No nonmetro counties Above the U.S. average No nonmetro counties Source: Bureau of Labor Slatistics. Source: Bureau of Labor Statistics. 57 Food and Fiber System Employment The food and fiber system is a major employer in nonmetro America. In 800 counties (785 non¬ metro and 15 metro) farming and farm-related industries employ a third or more of the local labor force. In 1986, the system accounted for nearly 30 percent, or 5.9 million, of the jobs in nonmetro America. Chart 113 Agribusiness counties, 1986 Chart 114 States with nonmetro areas most dependent on food and fiber sector employment County employment type; Hi Specialized (a third or more in farm production); Diversified (a third or more in farm production, agricultural input, and agricultural processing and marketing industries) Nebraska Iowa Minnesota North Dakota Missouri Tennessee Alabama Wisconsin South Dakota North Carolina Farm production Farm inputs industries Processing and marketing industries Other J 0 10 20 30 40 50 60 70 Percent of nonmetro employment 1986 data. Other category includes wholesale and retail trade ol agricultural products. Chart 115 Location of food and fiber employment Chart 116 Food and fiber employment in nonmetro areas Total Farm production Farm inputs Processing and marketing Wholesale and retail trade of agricultural products 1986 data. 0 20 40 60 80 100 Percent Region United States Northeast Lake States Corn Belt Northern Plains Appalachian Southeast Delta Southern Plains Mountain Pacific EE ee ED wmz. m El m me Farm production Farm inputs industries Processing and marketing industries Other J 0 10 20 30 40 50 60 70 Percent of nonmetro employment 1986 data. Other category includes wholesale and petal trade of agriculuxal products. 58 Nonmetro Versus Metro Employment Nonmetro counties adjacent to metro areas had better growth than more remote counties. Non¬ metro employment was stable or increased in all States except Louisiana. North Dakota, Okla¬ homa, Wyoming, Nebraska, and West Virgnia. Chart 117 Metro and nonmetro employment change since 1982 Chart 118 Average annual growth rates for nonmetro counties, 1982-87 Percenl of 1982 1983 84 Source: Bureau of Economic Analysis. 85 86 87 Urbanized adjacent Urbanized nonadjacent adjacent Less urbanize* nonadjacent Totally rural adjacent Totally rural nonadjacent Percent 2.5 1.8 1.2 Source Bureau ot Economic Analysis Chart 119 Metro and nonmetro employment change by State, 1982-87 Metro Nonmetro 1 1 Percent increase llffl Percent decrease 59 Industry Employment Employment in all industries grew more in metro than in nonmetro areas except lor the natural resources and manufacturing industries. Regionally, the West has the smallest percentage of manufacturing jobs. The agricultural sector is largest in the Midwest. Chart 120 Average annual employment change by industry, 1982-87 Chart 121 Importance of industries in metro and nonmetro areas All Industries Goods-producing Natural resources Manufacturing Construction Service-producing Trade Services Government L_ -4 Metro I ] Nonmetro VAAAAAAAA. ■'AAAAA/ 1 -2 0 2 4 6 Percent Government 15.1% Manufacturing 14.6% Construction 5.4% Agriculture 1.9% — Mining .5%- 5.4% Metro 1987 data. Nonmetro Chart 122 Regional importance of nonmetro industries Government 15.4%- South Midwest 1987 data. 60 Farm Credit Provision for loan losses caused the Farm Credit System’s net income to be at its lowest in 1985- 86. Overall farm debt has declined since then, and the distribution of debt has changed slightly. Agricultural interest rates remained strong in 1989 and continued to be highly differentiated be¬ tween lenders. Chart 123 Comparison of the farm credit system: Net interest income and net income Billion dollars -3.0 L 1984 85 86 87 88 89 Net interest Income minus provisions tor loan losses and other charges, equals net income. Values for 1989 are annuaized, second quarter data. Chart 124 Loans outstanding by major agricultural lenders Billion dollars 150 r 125 100 75 50 25 FmHA real estate Life insurance company real estate — FCS real estate Commercial bank real estate - FmFIA nonreal estate FCS nonreal estate -Commercial bank nonreal estate 1985 86 87 88 89 End ot year data except for 1989. Includes real estate and nonreal estate loans. Source: Agricultural Finance Data book. 1989. Chart 125 Selected agricultural Interest rates Percent 20 16 | 3-Month Treasury bills ^ FCS nonreal estate Commercial bank FmHA nonreal estate FmHA real estate FCS real estate 87 88 Yearly data except for 1989. Source: USDA AF0-32. selected banks of the FCS. and Agricultural Fi nance Data Book, various issues. 89 61 Banking Commercial banks headquartered in rural areas constituted 48 percent of all failures from 1983 to 1988, but only 33 percent of failures in 1988. Commercial banks headquartered in rural areas out¬ number urban-based banks, but hold only 12 percent of all bank deposits. About 31 percent of the savings and loans (S&L’s) are headquartered in rural areas, but hold only about 9 percent of S&L assets. Chart 126 Commercial bank failures by county, 1983-88 Chart 127 Savings and loan failures by county, 1983-88 Metro Nonmetro Metro Nonmetro Chart 128 Nonmetro/metro banks and bank assets by size % of U.S. total Under $25 million $25-$99 million $100 million—$1 billion Over $1 billion 50 25 Metro Nonmetro Bank assets Metro Nonmetro Number of banks Chart 129 Nonmetro/metro savings and loans assets by size % of 100 75 U.S. total Under $25 million $25-$99 million $100 million—$1 billion Over $1 billion Metro Nonmetro Number of savings and loans Metro Nonmetro Savings and loans assets December 31. 1988 data. U.S. commercial banks. December 31. 1988 data. 62 Earnings Total earnings grew at an annual rate of 2 percent for the nonmetropolitan United States during the 1969-86 period. This growth is due to growth in employment rather than any change in earnings per job. Growth was strongest in the South and West, but year-to-year stability of earnings growth was highest in the Northeast. Chart 130 Earnings growth in nonmetropolitan counties, 1969-86 Chart 131 Stability of earnings growth in nonmetropolitan counties, 1969-86 ] Metro Chart 132 Components of earnings growth in nonmetropolitan counties, 1969-86 Percent 4 r -1 All counties High-growth Low-growth High-stability Low-stability 63 Economic Indicators A sluggish manufacturing sector slowed growth in real gross national product in 1989. Chart 133 General economic indicators: Industry % of 1982 % of 1977 % change from previous year Ratio $ billion $ billion 1982 525 500 475 450 425 400 1984 85 86 87 88 89 64 Economic Indicators Inflation increased and interest rates rose, but the unemployment rate remained at decade lows and real consumer income continued to climb in 1989. Chart 134 General economic Indicators: Personal economy % change from previous year $ billion 1982 % of all civilian workers % change from previous year Percent % of disposable personal income 65 Consumer Prices Retail food prices rose nearly 6 percent in 1989, the greatest increase since 1981. The rise was the result of lower supplies of some products due to lingering effects from the drought of 1988 and poor weather conditions in the first half of 1989. Strong consumer demand for meats also played a role in higher prices. Chari 135 Consumer price index for food i p All food I I All items I- ■hi 1 I Annual percentage change 14 - M. I 1979 81 83 85 1989 forecast. Source: Bureau of Labor Statistics. 87 89 - 12 - 10 8 6 4 Food at home 2 0 Food away from home i- t H-| 1 1979 81 83 85 87 89 Chart 136 Retail price, farm value, and price spread for food % of 1982-84 Chart 137 Farm value share of retail food prices Percent Eggs 58 Choice beef 56 Frying chicken 58 Fresh milk 46 Frozen orange juice concentrate Pork 38 39 Average for market basket 30 HP Fresh fruit and vegetable 26 Margarine 26 Canned tomatoes 11 White bread 8 1988 data. Farm value share is the proportion the farmer receives from the dollar the consumer spends. The remainder of the dollar goes to marketing firms. 66 Food Marketing Costs The marketing bill, the largest share of the food dollar, has risen faster than the farm value of raw foodstuffs, reflecting the rising cost of labor, packaging, and other inputs. Labor costs were respon¬ sible for over a third of the food dollar. Farm value, as a percentage of the total food dollar, is lower for the away-from-home market due to added costs of preparing and serving food. Chart 138 Food processing, wholesaling, and retailing costs % of 1982-84 1989 forecast. The marketing cost index measures changes in worker wages, salaries, and supplemental benefits, and prices of purchased inputs such as packaging materials and fuel and power. Chart 140 What a dollar spent on food paid for in 1988 Farm value 25 $ - Marketing bill: Packaging 8 $ - Transportation 4.5 4 — Before-tax profits 3.0 bbyists, trade association employees, and tudents from high school to graduate school re just some of the groups who can benefit, jven farm policy experts will want to order uiltiple copies of this informative report to help how their clients what it’s all about. lasic Mechanisms of U.S. Farm Policy uses asy-to-understand language and diagrams to lescribe farm policy mechanisms. Be sure to irder enough for both your staff and your clients! A sampling of what's in “Basic Mechanisms of U.S. Farm Policy. The complete array of farm policy mechanisms can appear overwhelming to anyone unfamiliar with the history of U.S. agricultural legislation. But each mechanism originated in Congress, reflecting public concerns about food, agriculture, and the needs of farmers. SOME BASIC MECHANISMS OF U.S. FARM POLICY Target Price Loan (Nonrecourse loan) Rate Deficiency Payment Original Deficiency Reduced (Findley) Loan Rate Emergency Compensation Acreage Reduction Program (ARP) Paid Diversion Base Acres Program Yield Program Production Basic Commodities Acreage Conservation Reserve Conservation Use Payment Limitation Projected Deficiency Advance Deficiency Base Acres & Program Yield 0-92 & 50-92 Commodity Certificate Posted County Price (PCP) PIK and Roll Export Enhancement Farmer-Owned Reserve (FOR) Corn (& Wheat) Catalog Reserve Rollover Conservation Reserve Program Disaster Payment Marketing Loan Part one of this report concen¬ trates on the left side of this list, and Part two covers the seven mechanisms at the top right. Part three covers the re¬ maining seven mechanisms on this list. isic Mechanisms of U.S. Farm Policy (MP-1479) Yes! Send me_ copies of Basic Mechanisms of U.S. Farm Policy for $_. Order up to 50 copies and pay $6.50 per copy. Save 25%! Order 50 or more and pay just $4.90 per copy. ayment method Bill me □ Enclosed is $_. Visa □ MasterCard Total charges $_, For fastest service, order toll free, 1-800-999-6779 (8:30-5:00 ET, in the United States and Canada; other areas, please call 301-725-7937) • Use purchase orders, checks drawn on U.S. banks, cashier’s checks or international money orders. edit card number: JJJJJJJJJJJJJJJJ Make payable to ERS-NASS. Add 25% for postage to foreign addresses (includes Canada). □□ Expiration date: Month Year Name Andress City, State, Zip_ Daytime phone () Mail to: ERS-NASS P.O. Box 1608 Rockville, MD 20849-1608 UNITED STATES DEPARTMENT OF AGRICULTURE ECONOMIC RESEARCH SERVICE 1301 NEW YORK AVENUE. NW. WASHINGTON, D. C. 20005-4788 fc»30 ju 3> c^> A. gricuiturai esearch ervice griculture andbook umber 690 liNIVtRSJtY OF ILLINOIS IGRIOULTURE LIBRARY Diagnosis of Honey Bee Diseases UNIVERSITY OF ILLINOIS LAW LIBRARY MAY 7 1991 federal depository ~ 76 > '• Mention of companies or commercial products does not imply recommendation or endorsement by the U.S. Department of Agriculture over others not mentioned. Copies of this publication may be purchased from the National Technical Informa¬ tion Service, 5285 Port Royal Agricultural Research Service has no additional Uopd'eS Cor free distribution. Abstract Shimanuki, Hachiro, and David A. Knox. 1991. Diagnosis of Honey Bee Diseases. U.S. Department of Agriculture. Agriculture Handbook No. AH-690, 53 p. Apiary inspectors and beekeepers must be able to recognize bee diseases and parasites and to differentiate the serious diseases from the less important ones. This handbook describes laboratory techniques used to diagnose diseases and other abnormalities of the honey bee and to identify parasites and pests of the honey bee. Emphasis is placed on the tech¬ niques used by the U.S. Department of Agriculture Bee Research Labora¬ tory. Included are directions for submitting, through APHIS-PPQ or State regulators, samples of suspected Africanized honey bees for identification of subspecies. Also included are directions for sending diseased brood and adult honey bees for diagnosis of bee disease. Keywords: honey bee disease, honey bee disorder, honey bee parasite, honey bee pest. Africanized honey bee AGRICULTURE LIBRARY JUN 1 2 1991 UNIVERSITY OF ILLINOIS Contents Page Introduction . ] Methods of Diagnosing Disease . i Techniques of Microscopic Examination. 1 Microinjection Techniques. 2 Removal of Digestive Tract. 5 Brood Diseases . 5 Bacterial Diseases. 7 Fungal Diseases.17 Viral Disease: Sacbrood.19 Mixed Infections.19 Diseases of Adult Bees. 19 Protozoan Diseases.19 Bacterial Diseases. 23 Viral Diseases.24 Noninfectious Disorders ..25 Neglected Brood. 26 Overheated Bees.27 Genetic Lethality.27 Plant Poisoning. 27 Pesticide Poisoning.30 Parasitic Honey Bee Mites .32 Honey Bee Tracheal Mite (Acarapis woodi) .33 Varroa jacobsoni .37 Tropilaelaps clareae .42 Pests .. Wax Moths.43 Bee-louse (Braula coeca) .43 Melitfiphis alvearius .44 Appendix A. Directions for Sending Diseased Brood and Adult Honey Bees for Diagnosis .45 Appendix B. Identification of Africanized Honey Bee .47 References .49 This publication supersedes "Diagnosis of Honey Bee Issued April 1991 ii Diagnosis of Honey Bee Diseases Hachiro Shimanuki 1 and David A. Knox Introduction Inspection for bee disease is an important part of beekeeping. Apiary inspectors and beekeepers must be able to recognize bee diseases and parasites and to differentiate the serious diseases from the less important ones. The purpose of this publication is to acquaint readers with labora¬ tory techniques used to diagnose diseases and to detect and identify parasites, pests, and other abnormalities of the honey bee. We realize that different laboratory methods are used by others; where possible, those methods are described. However, emphasis is placed on the techniques used by the U.S. Department of Agriculture Bee Research Laboratory. Directions for submitting samples for diagnosis or subspecies identifica¬ tion are included in appendixes A and B. Methods of Diagnosing Disease Techniques of Microscopic Examination Most bee diseases can be diagnosed by observing the associated micro¬ organisms with a light microscope. The following techniques are com¬ monly used to prepare microscope slides for examination: Modified Hanging Drop The modified hanging drop technique (Michael 1957) can be very useful for differentiating diseases of the brood. Residue from a suspected cell is first mixed with water. Then a drop of this suspension (smear) is placed on a cover glass. The suspension used should always be dilute and only slightly turbid. The smear is dried and fixed under a heat lamp, or the smear can be air dried and heat fixed by passing it rapidly through a bunsen burner flame two or three times. The fixed smear is stained with carbol fuchsin 2 or a suitable spore stain for 10 seconds. Enough stain should be placed on the cover glass to cover the entire smear. The excess 'Shimanuki is a microbiologist and Knox an entomologist at Bee Research Laboratory, Agricultural Research Service, U.S. Department of Agriculture, Beltsville, MD 20705. 2 Solution A: 0.3 g basic fuchsin (90% dye content), 10 mL ethyl alcohol (95%); solution B: 5 g phenol, 95 mL distilled water. Mix solutions A and B. 1 stain is then washed off with water. While the smear is still wet. the cover glass is inverted with the smear side down and placed onto a standard microscope slide previously coated with a very thin layer of immersion oil. The slide is gently blotted dry and examined with a microscope using the oil immersion objective. Results: Organisms that are not heat fixed are caught in areas where pockets of water have formed in the oil, and the organisms usually exhibit Brownian movement (see section on American foulbrood). Simple Stain This method relies solely on differentiating the bacteria by morphology. Place a drop of the suspension directly on a microscope slide. Heat fix and stain the smear as described in the previous section. Carbol fuchsin, methylene blue, and safranin are examples of stains that can be used. Air- dry or gently blot the stained smear. Place a drop of immersion oil directly on the smear. No cover glass is necessary. Examine the slide, using the oil immersion objective. Results: Organisms are uniformly stained and easily distinguished. Gram Stain The Gram stain is a standard microbiological method that can be substi¬ tuted for the simple stain. Briefly, the procedure is as follows: A fixed smear is stained with crystal violet, immersed in iodine solution, decolor¬ ized in ethyl alcohol, and counterstained with safranin. Results: Gram¬ positive organisms are blue; Gram-negative organisms are red. Wet Mount The wet mount is especially useful for examining fungi or protozoa. Macerate a portion of the sample in water. Place a drop of the suspension on a microscope slide, and carefully drop the cover glass on it to minimize air pockets. No stain is required. The wet mount is usually examined with the dry objectives of a microscope. Results: Organisms refract light and are therefore visible on the slide. A phase-contrast microscope may be helpful, especially if an oil immersion objective is required. Microinjection Techniques To diagnose some diseases or to determine toxic levels of sample materi¬ als, it may be necessary to feed or inoculate larvae, pupae, or adult honey bees. Michael (1960) developed a technique using a microinjector 2 equipped with a syringe and a 30-gauge needle. The microinjector can be calibrated to repeatedly deliver uniform inoculating volumes as small as 1 pL. The apparatus can be used to introduce material orally into the midgut (ventriculus) of a larva or to feed individual adult honey bees. The microinjection technique can also be adapted to direct injections into the body cavity of larvae, pupae, and adults. Collecting Honey Bee Larvae and Pupae Honey bee larvae 3 to 5 days old are readily obtained by removing a brood frame containing the desired larvae from a colony and placing it horizon¬ tally above a towel-lined tray in an incubator at 34°C. Within a few hours, the larvae crawl from their cells and drop to the tray below. Pupae can be easily obtained by collecting 5-day-old larvae as described above and incubating them in petri dishes until pupation occurs. This method of collecting larvae and pupae in large numbers saves considerable time and labor. It also eliminates the damage that can occur when attempting to remove these immature forms from cells by mechanical means. Oral Introduction Larvae Honey bee larvae as young as 3 days and weighing as little as 25 mg can be force-fed by carefully inserting the needle through the mouthparts and into the esophagus (fig. 1). When the actuating lever is depressed, a Esophagus Microinjector Figure 1. Force-feeding a honey bee larva. 3 predetermined volume ot material is propelled through the esophagus and into the midgut, with no physical damage to the larva. After feeding, the larvae are placed in petri dishes lined with filter paper and incubated at 34°C. Adults Individual adult honey bees can also be fed known volumes using the microinjector. Adult bees are collected and held in a cage for about 4 hours without food. The material to be fed, at the final concentration, should be mixed into a sucrose solution to make it attractive to the bees. The microinjector is first actuated to produce a known volume of liquid at the tip of the needle. Then a bee from the cage is grasped by the wings, held up to the drop ot liquid, and allowed to feed. Cold temperatures can be used to slow the bees and make them easier to handle. Avoid the use of carbon dioxide as an anesthetic to aid in handling bees, because they are reluctant to feed after exposure to carbon dioxide and their longevity is reduced. After feeding, the bees are placed in small cages with a supply of sugar syrup and held in an incubator at 34°C. Direct Injection Care should be taken to insert only the tip of the needle into the hemocoel and to not exceed inoculating volumes of 2 pL per bee. Brood Injections are usually confined to 4- to 5-day larvae. The larva is held gently between the first and second fingers and the thumb, and the larva must be absolutely parallel to the needle. After the integument is punc¬ tured by the needle under gentle pressure, the inoculum is expelled directly into the dorsal blood vessel; the needle is then withdrawn in a slow, steady movement. Any excessive pressure on the larva by the fingers, particularly when withdrawing the needle, must be avoided to prevent bleeding. Pupae can be inoculated dorsally between the third and fourth abdominal seg¬ ments or through the propodeum of the thorax. After injection, the larvae or pupae are placed in petri dishes lined with filter paper and incubated at 34 C. If excessive bleeding has occurred, it can be seen on the filter paper. Pupae can also be left in brood combs and inoculated in the head capsule (Wilson 1970). The pupal head is exposed by removing the cell cap. and the needle is inserted between the ocelli or through the clypeal sclerite. 4 Adults Adult bees can be injected either through the propodeum of the thorax or dorsally through the intersegmental membrane between the third and fourth abdominal segments. Adult bees should be carefully subjected to carbon dioxide anesthetic before and during the process of injection. When bee longevity is a factor in the test, cold temperature can be used as an anesthetic. After injection, the bees are placed in small cages with a supply of sugar syrup and held in an incubator at 34°C. Removal of Digestive Tract Intact digestive tracts that have been removed from adult honey bees are very useful for the detection of protozoan diseases (refer to fig. 9). The digestive tracts can be easily obtained by removing the head of the bee to free the digestive tract, grasping as much of the stinger as possible with a pair of fine tweezers, and then with a steady, gentle pull withdrawing the entire digestive tract. Freshly killed honey bees are required for this procedure. Brood Diseases Brood combs from healthy colonies typically have a solid and compact brood pattern. Almost every cell from the center of the comb outward contains an egg, larva, or pupa. The cappings are uniform in color and are convex (higher in center than at margins). The unfinished cappings of healthy brood may appear to have punctures, but since cells are always capped from the outer edges to the middle, the holes are always centered and have smooth edges. In contrast, brood combs from diseased colonies usually have a spotty brood pattern (pepperbox appearance), and the cappings tend to be darker, concave (sunken), and punctured. Also, the combs may contain the dried remains of larvae or pupae (scales), which are found lying lengthwise on the bottom side of brood cells. Sometimes scales are difficult to locate because of the condition ot the comb. Scale material can be easily located using long-wave ultraviolet or near¬ ultraviolet light. Exposure to wavelengths of 3100-4000 angstroms will cause scale material to fluoresce. Some discretion must be used with this technique because honey and pollen may also fluoresce. Symptoms of various brood diseases are summarized in table 1. Symptoms of a contagious disease are sometimes mimicked because of an unrelated factor. For instance, often brood that is neglected because of a shortage of nurse bees will die from either chilling or starvation. 5 to a> 0 jQ to a> to O 0 to ■o P O O to 3 o o > o to E o a E >* D O a E o O ® P o P O O k__ P O x U P o o p o o co >* a> c o P O O =J o C a CD a o o P O O ,0 D o c o o 0 E < E o a E > CO 73 0 O D O > Pp O OPp § O w vir -3 +- CO D ' >— co — 3< D O ^ M— j : (DCt^D ^ = (1) w ^d)2t(D ° O P O o o — . _ "O O co ■7— P 0 C 0)0 p o o - 5 O o£ Cl£ CD O '5 o - 5 coco > Q. 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3 O 0C ^ 0 -S O) 0 c O>w D O CO O > Si CO Q p8 ,g>® CO P O P O 0O o 9- O O) C ^ o. •o op ^ o J?— c?5 73) P O 0 P 8p ° O p 2 Op Q) O U <§-0 >-p ^ ^ © o O 4z O :S|? 0 0 > |z® CD $ O c ©p E| 2 o ^p 0 Or O) = o) 0 O © = -^ O 532S5 0 O-c 0 D X O °£ 5-^ CD ScP = p - 2 O 0 73)0 C^_ O D t O ®P=o to C ®CD ^ 0 X < o £ © P-Q' o . 5= P 3 = X P to 0 0)0 3 0^=07 0 O O £o 0‘ w .o 73)ir - CO -j— — co — E|2o - ^ Q) 5°£_ C C P 0 3 0< O 0 OP 0>E^ O D:= O D)E p o n 0 O0D 2o® . 0 Q 1 ® llll nil. o 0 t3 © 2 o o O P CO O Symptoms can also be the result of a failing queen, laying workers, toxic chemicals, or poisonous plants. (See section on Noninfectious Diseases.) Bacterial Diseases American Foulbrood Bacillus larvae is the bacterium that causes American foulbrood disease (AFB). Bacillus larvae is a slender rod with slightly rounded ends and a tendency to grow in chains (fig. 2). The rod varies greatly in length, from about 2.5 to 5 microns (pm), and is about 0.5 pm wide. The spore is oval and approximately twice as long as wide, about 0.6 by 1.3 pm. When stained with carbol fuchsin, the spore walls appear reddish purple and quite clear in the center. The spores may form clusters and appear to be stacked. Approxi¬ mately 2.5 billion spores are produced in each infected larva. If the larva has been infected for less than 10 days, the vegetative cells are present, and some newly formed spores may be seen. The modified hanging drop technique can be very useful for differentiating American foulbrood from other brood diseases. In areas of the smear where pockets of water are formed in the oil, the spores of Bacillus larvae exhibit Brownian movement. This is an extremely valuable diagnostic technique because the spores formed by the other Bacillus Figure 2. Bacterium that causes American foulbrood disease (not to scale): Top, Bacillus larvae vegetative cells; middle, Bacillus larvae spore formation; bottom. Bacillus larvae spores. 7 species associated with the known bee diseases usually remain fixed (see table 2). It is important to note that Brownian movement can be affected by slide preparation; also, debris and other bacteria can exhibit this motion. Therefore, Brownian movement must not be used as the sole criterion for diagnosis but must be considered together with the character¬ istic morphology of the spores and the gross larval symptoms. If micro¬ scopic examination is not conclusive, cultural tests can be made using the same suspension. Cultivation of Bacillus larvae Thiamine (vitamin B^ and some amino acids are required for the growth of Bacillus larvae. Routine culture media such as nutrient broth will not support the growth of this organism. Good vegetative growth occurs on Difco brain heart infusion fortified with 0.1 mg thiamine hydrochloride per liter of medium (BHIT) and adjusted to pH 6.6 with HC1, but sporula- tion does not occur. Satisfactory growth and sporulation occur on the yeast extract, soluble starch, and glucose media recommended by Bailey and Lee (1962). The medium can be liquid, semisolid (0.3% agar), or solid (2% agar). For more information on sporulation, see Dingman and Stahly (1983). Bacillus larvae spores also reproduce in the hemolymph of honey bee larvae, pupae, and adults when artificially introduced by injection (Michael 1960, Wilson and Rothenbuhler 1968, Wilson 1970). To culture Bacillus larvae , we prepare spore suspensions by mixing diseased material (scales) with 9 mL sterile water in screw-cap tubes. (We use cotton swab applicators to remove and transfer the scales from Table 2. Differentiation of Bacillus species in honey bees Brownian Species movement' Catalase production Nitrate reduction Growth on nutrient agar Bacillus larvae + — + _ Bacillus alvei + - + Bacillus laterosporus + + + Bacillus pulvifaciens - + + ' In modified hanging drop technique. the comb to the tubes.) The suspension is heat shocked at 80°C for 10 minutes (effective time) to kill nonsporeforming bacteria. A sterile cotton swab is used to evenly spread a portion of the suspension (approximately 0.2 mL) over the surface of BHIT agar plates, which are then incubated for 72 hours at 34°C. Individual colonies are small (1-2 mm) and opaque; however, if large numbers of viable B. larva spores are inoculated, a solid layer of growth will cover the plate. There are no reliable methods for making plate counts of Bacillus larvae because fewer than 10 percent of the spores will produce visible growth on the media presently available (Shimanuki 1963). By calibrating our methods using spore and plate counts, we have determined that a minimum of 100 B. larvae spores are required to produce visible growth on BHIT. Diagnostic Tests for Bacillus larvae Holst Milk Test. The Holst milk test (Holst 1946) is a simple test based on the fact that a high level of proteolytic enzymes is produced by sporulating Bacillus larvae. The test is conducted by suspending a suspect scale or a smear of a diseased larva in a tube containing 3-4 mL of 1% powdered skim milk in water. The tube is then incubated at 37°C. If B. larvae is present, the suspension should clear in 10-20 minutes. It should be noted that this test is not always reliable. Nitrate Reduction. Bacillus larvae reduces nitrate to nitrite (Lochhead 1937). The nitrate reduction test can be performed on a medium such as BHIT, which contains potassium nitrate (1-2 mg/L of medium). After growth has occurred, the addition of a drop of sulfanilic acid-alpha- naphthol reagent produces a red color if nitrate has been reduced to nitrite. Diagnosis should not be based on this test alone but on this test along with larval gross symptoms, bacterial morphology, and growth characteristics of the bacterial colony. Catalase Production. A drop of 3% hydrogen peroxide is placed on an actively growing culture on a solid medium. Most aerobic bacteria break down the peroxide to water and oxygen and produce a bubbly foam, but Bacillus larvae is almost always negative for this reaction (Haynes 1972). Fluorescent Antibody. The fluorescent antibody technique requires the preparation of specific antibodies stained with a fluorescent dye. Rabbits are injected with pure cultures of Bacillus larvae, and the active antiserum is collected and stained with a fluorescent dye. This fluorescent antiserum 9 is mixed with a bacterial smear on a slide and allowed to react. The excess antiserum is washed off the slide, and the slide is then examined with a fluorescence microscope. B. larvae appears as brightly fluorescing bodies on a dark background (Toschkov et al. 1970, Zhavnenko 1971. Otte 1973, Peng and Peng 1979). Viability Test for Bacillus larvae One method of controlling AFB is to destroy the viability of Bacillus larvae spores in contaminated bee equipment. This can be accomplished by gamma or electron beam irradiation or by fumigation with a sterilant gas such as ethylene oxide. Assessment of the efficacy of these methods should be based on the number of spores remaining viable in a test sample of brood comb containing at least 10 AFB scales. A spore suspension is prepared from the sample comb by mixing 10 scales in 10 mL sterile water. Since each scale contains about 2.5 billion spores (Sturtevant 1932), 1 mL of the suspension should contain 2.5 billion spores. A portion of the suspension (0.2 mL = 500 million spores) is spread onto solid BHIT plates as previously described and incubated for 72-96 hours. The results of the viability tests are reported as the approximate number of viable spores on a per-scale basis. If no colonies form on the medium, the results are recorded as <100 viable spores per scale; 1-9 colonies are recorded as <1,000 per scale; 10-99, <10.000 per scale; over 100 colonies per plate, >10,000 viable spores per scale; and when the plate is com¬ pletely overgrown with colonies, the results are reported as no detectable reduction of viable spores. Terramycin (Oxytetracycline) Resistance Tests Isolates ot Bacillus larvae can be screened for sensitivity to oxytetracycline based on the size of inhibition zones on agar plates. A spore suspension ot the B. larvae isolate to be tested is spread on solid BHIT as described previously. A disk (BBL Sensi-Disc) containing 5 gg oxytetracycline is then placed on the plate, and the plate is incubated at 34°C for 72 hours. The zones formed by sensitive strains usually average 50 mm in diameter. Alternatively, oxytetracycline incorporated into liquid BHIT will inhibit the growth of sensitive strains of B. larvae at concentra¬ tions as low as 12 gg/L ot medium (Gochnauer 1953). Care is necessary in making these tests, and adequate control strains should be included. 10 Any substantial reduction of the zone size or an increase in the concentra¬ tion of oxytetracycline required to prevent growth of Bacillus larvae in liquid medium would be evidence of the development of resistant strains (Gochnauer et al. 1975). However, when interpreting the results of the tests, the effects of growth rates should be considered. Because strains often grow at different rates, one may falsely conclude that a strain is resistant. No resistance of B. larvae to oxytetracycline has yet been reported. Detection of Bacillus larvae Spores in Hive Products Honey. Occasionally it may be necessary to examine honey for the presence of Bacillus larvae. Due to the high concentration of carbo¬ hydrate and other natural bacteriostatic substance(s) in honey, the exami¬ nation of honey requires special considerations. The classical method (Sturtevant 1932, 1936) is to dilute the honey 1:9 with water, centrifuge the mixture to concentrate the spores in the sediment, and then examine the sediment microscopically for the presence of spores. We (Shimanuki and Knox 1988) have developed the following technique to detect Bacillus larvae spores in honey. Honey to be examined is heated to 45°C to permit easier handling and to decrease viscosity for more uniform distribution of any spores that may be present. Twenty-five milliliters of honey is placed in a 50-mL beaker and diluted with 10 mL of sterile water. The diluted honey is then transferred into a 1.75-inch (44- mm) dialysis tube. The open end is tied after the tube is tilled, and the tube is submerged in running water for 18 hours or in a water bath with three to four water changes in that period. Following dialysis, the contents of the tube are centrifuged at about 2,000 g for 20 minutes. The super¬ natant is carefully removed with a pipet to leave approximately 1 mL ot residue. This residue is resuspended in 9 mL of water in a screw-cap vial and heat shocked at 80°C for 10 minutes to kill nonsporeforming bacteria. Next, 0.5 mL of the suspension is spread onto a plate of BHIT agar. The plate is incubated at 37°C for 72 hours and examined tor colonies ot B. larvae. Difficulties can sometimes occur when honey samples contain other sporeforming bacteria that may completely cover the surface ot the plate. Since approximately 100 Bacillus larvae spores are required to produce visible growth on BHIT, this technique can demonstrate the presence ot B. larvae spores in samples that contain a minimum of 80 spores per mL ot undiluted honey (25 mL honey X 80 spores/mL = 2,000 spores/dialysis = 2,000 spores in 10 mL or 200 spores/mL; 0.5 mL = a 100-spore inocu- 11 y lum). Lower spore levels can possibly be detected by the use of larger honey samples or a second centrifugation to further concentrate the spores. Pollen. Bacillus larvae spores can also be recovered from bee-collected pollen pellets by physically removing bits of AFB scale. A series of sieves ol different sizes is helpful. It scales are not detected, one may pass a water-pollen suspension through No. 2 filter paper, centrifuge the filtrate, and culture the pellet as described above (Gochnauer and Comer 1974). Beeswax. We have had some success in recovering spores morphologically similar to those of Bacillus larvae by melting beeswax in boiling water, removing the beeswax cake after cooling, and centrifuging the water at 2,000 g for 20 minutes. The sediment is then examined microscopically for the presence of spores. Spores have also been recovered from contami¬ nated beeswax by chloroform extraction (Kostecki 1969). However, in both cases, positive identification of the spores is not possible because the recovery techniques render the spores nonviable. European Foulbrood Melissococcus pluton (= Streptococcus pluton) is the bacterium that causes European foulbrood disease (EFB). Melissococcus pluton has been reclassified into the new genus Melissococcus (Bailey and Collins 1982a and b). However, Melissococcus pluton has still not been adequately described to be accepted for the current editions of “Bergey’s Manual of Determinative Bacteriology.” Melissococcus pluton is generally observed early in the infection cycle before the appearance of the varied microflora associated with this disease. The M. pluton cell is short, nonsporeforming, and lancet shaped. The cell measures 0.5-0.7 by 1.0 pm and occurs singly, in pairs, or in chains (fig. 3). Cultivation of Melissococcus pluton Melissococcus pluton can be isolated on a medium developed by Bailey (1959). The medium consists of 1 g yeast extract (Difco), 1 g glucose, 1.35 g potassium dihydrogen phosphate (KH,P0 4 ), 1 g soluble starch, 2 g agar, and distilled water to make 100 mL; the pH is adjusted to 6.6 with potas¬ sium hydroxide (KOH) and the mixture is autoclaved at 10 lb per square inch (116 C) for 20 minutes. It has been found that the addition of cysteine (0.1 g per 100 mL) improves the multiplication of M. pluton (Bailey and Collins 1982b). 12 §4 t <•!&& i t .fe f * ^ ; # :» I Figure 3. Melissococcus pluton and Bacillus alvei ( X 1200). « *.< tr f'» S i % It is difficult to isolate Melissococcus pluton on artificial media because of its growth requirements and the competition from other bacteria. Also, once isolated, identification of M. pluton is difficult due to its pleomorphic nature in culture (fig. 4). Melissococcus pluton is best isolated when few if any other organisms are present. According to Bailey (1959), it is best to dry smears of diseased larval midguts on a slide. A water suspension of this material or a suspension prepared from larvae (apparently healthy, infected, or dead), cappings, etc., can be streaked on freshly prepared Bailey’s agar medium. Or decimal dilutions of these suspensions can be inoculated into molten Bailey’s agar medium (45 C) and poured into plates (Bailey 1981). The plates are incubated anaerobically at 34°C. The “Gas Pak” (BBL) Anaerobic System, including a disposable hydrogen and carbon dioxide generator, is used to obtain anaerobic conditions. Small white colonies of M. pluton should appear after 4 days. 13 us: X Figure 4. Melissococcus pluton as it appears in culture (X 1200). Serodiagnosis Pinnock and Featherstone (1984) developed an enzyme-linked immunosorbent assay (ELISA) for detecting Melissococcus pluton. Using this technique, they were able to demonstrate the presence of M. pluton even in apparently healthy honey bee colonies. Organisms Associated With European Foulbrood Some organisms do not cause European foulbrood. but they influence the odor and consistency of the dead brood and can be helpful in diagnosis. These secondary invaders include the following: Bacillus alvei. The bacterium Bacillus alvei is frequently present in cases of European foulbrood disease (EFB). It is a rod 0.5-0.8 pm wide by 2.0- 5.0 pm long (fig. 5). Spores measure 0.8 by 1.8-2.2 pm. Like Bacillus lai vae , the spores may be clumped and appear stacked. The sporangium may be observed attached to the spore. Typical strains of B. alvei spread V Figure 5. Bacteria associated with European foulbrood disease (not to scale): Top, Bacillus alvei; middle, Bacillus laterosporus; bottom, Entero¬ coccus faecalis. vigorously on nutrient agar and may show “motile colonies”; free spores may lie side by side in long rows on the agar. The growth of this bacterium produces an unpleasant odor. Bacillus laterosporus (= Bacillus orpheus). Rods of Bacillus laterosporus measure 0.5-0.8 pm by 2.0-5.0 pm, and the spores 1.0- 1.3 by 1.2-1.5 pm (fig. 5). An important diagnostic feature is the production of a canoe-shaped parasporal body that stains very heavily along one side and the two ends, and remains firmly adherent to the spore after lysis of the sporan¬ gium. The clear portion with the finely outlined wall is the spore. B. laterosporus grows moderately on nutrient agar, becoming dull and opaque, and spreads actively if the agar surface is moist. Growth on nutrient agar with 1% glucose added (glucose agar) is thicker and may become wrinkled. Enterococcus faecalis (= Strepto¬ coccus faecalis = Streptococcus apis = Streptococcus liquefaciens). Ovoid cells (elongated in the direction of chain) are 0.5-1.0 pm in diameter and are usually in pairs or short chains (fig. 5). This organism resembles Melissococcus pluton and may exhibit Brownian movement when the modified hanging drop technique is used. Growth occurs on nutrient agar usually within 1 day. Colonies are generally smaller than 2 mm; they are smooth and 15 convex, with a well-defined border. When magnified, the colonies appear light brown and granular. Bacterium eurydice (= Achromobacter eurydice ). There is no standard description of Bacterium eurydice. White (1912) described this organism as a small, slender Gram-negative rod with slightly rounded ends, occur¬ ring singly or in pairs and measuring 0.5-1.4 pm long by 0.4-0.7 pm wide. According to White (1920), Bacterium eurydice is best isolated by plating the midgut contents of infected larvae on glucose agar and incubating at room temperature. Growth is slow and never luxuriant, and colonies are convex, smooth, and glistening. However, later researchers, who were unable to isolate B. eurydice as described by White, used the name Bacterium eurydice for a Gram-positive bacterium isolated from diseased larvae. Therefore, mention of the name in the literature causes confusion. This organism is not included in the current editions of Bergey’s manual. Bacillus apiarius. The bacterium Bacillus apiarius is rarely encountered and may or may not be legitimately associated with EFB. Rods are 0.6-0.8 pm in diameter and often less at the poles. Special diagnostic features include the ridged, thick, rectangular spore coat and the stainable remnants of the sporangium, which remain attached for a considerable time. Growth can occur on Sabouraud dextrose medium. Powdery Scale Bacillus pulvifaciens is the bacterium that causes powdery scale disease. Powdery scale disease is seldom reported, perhaps because the average beekeeper is unable to identify it. A useful diagnostic characteristic is the scale that results from the dead larva. The scale is light brown to yellow and extends from the base to the top of the cell. The scale is powdery; when handled, it crumbles into a dust. Bacillus pulvifaciens vegetative cells measure 0.3-0.6 pm by 1.5-3.0 pm. The spores are 1.0 by 1.3-1.5 pm. The bacterium can be isolated on nutrient agar, but growth is heavier on glucose agar. When first isolated, the organism produces a reddish-brown pigment that can be lost by subculturing. Bacillus pulvifaciens closely resembles Bacillus larvae, but the spores do not exhibit Brownian movement in the modified hanging drop technique. Also, B. pulvifaciens is distinguished by its ability to grow at 20°C and by its growth on nutrient agar. 16 Fungal Diseases Chalkbrood Ascosphaera apis is the fungus that causes chalkbrood disease. Ascosphaera apis is a heterothallic organism and develops a character¬ istic spore cyst when opposite thallic strains (+ and -) fuse. Spore cysts measure 47-140 pm in diameter (fig. 6). Spore balls enclosed within the cyst are 9-19 pm in diameter, and individual spores are 3.0-4.0 pm by 1.4-2.0 pm. Chalkbrood disease can be easily identified by its gross symptoms. An affected larva becomes over¬ grown by fluffy cottonlike mycelia and swells to the size of the cell. It only one strain (+ or -) of mycelium is present, the larva dries into a hard, shrunken, white chalklike mummy—thus the name chalkbrood. When the + and - mycelia are present in a diseased larva, spore cysts can form, and the resulting mummies appear either mottled (black on white) or com¬ pletely black. In heavily infected hives, mummies can be found at the hive entrances or on the bottom boards. Mummies can sometimes be detected in brood cells by tapping the comb against a solid surface. This easy removal of larval remains also differentiates chalkbrood from other brood diseases. Ascosphaera apis grows luxuriantly on potato dextrose agar fortified with 4 g yeast extract/L. Growth and sporulation also occur on malt agar but less profusely and with no aerial hyphae; this facilitates subculturing and microscopic examination. Cultures have a characteristic fruity odor similar to that of fermenting peaches. The optimum temperature tor growth is 30°C. Ascosphaera apis can be easily isolated from newly infected larvae or fresh mummies. These can be placed directly on the medium and incu¬ bated. New mycelial growth is usually visible within 24 hours. Small blocks of agar containing mycelia can be transferred to new plates to obtain pure cultures and isolates of the + and - strains. A. apis can be Figure 6. Spore cyst of Ascosphaera apis containing spore balls, which in turn contain spores. 17 v isolated from old mummies by placing them on water agar (agar with no added nutrients), incubating them, and transferring the new mycelial growth to a nutrient medium. Difficulties sometimes occur because A. apis may fail to grow or may be overgrown by other fungi, which can contaminate old mummies. If only one strain (+ or -) is isolated, a fluffy cottonlike growth will eventually cover the plate. When both the + and - thalli are isolated, spore cysts form throughout the culture. The + and - thalli are morphologically identical. They can be distinguished by inoculating isolates on opposing sides of a plate. When opposite thalli grow together, a line of spore cysts forms at the juncture. Stonebrood Stonebrood is usually caused by Aspergillus flavus , occasionally A. fumigants, and sometimes other Aspergillus species. These fungi are common soil inhabitants that are pathogenic to adult bees, other insects, mammals, and birds. The disease is difficult to identify in its early stages of infection. The fungus grows rapidly and forms a characteristic whitish- yellow collarlike ring near the head end of the infected larva. A wet mount prepared from the larva shows mycelia penetrating throughout the insect. After death, the infected larva becomes hardened and quite difficult to crush—hence the name stonebrood. Eventually, the fungus erupts from the integument of the insect and forms a false skin. At this stage, the larva may be covered with green powdery fungal spores. The spores of Aspergillus flavus are yellow green, and A.fumigatus spores are gray green. These spores can become so numerous that they fill the comb cells that contain the affected larvae. Stonebrood can usually be diag¬ nosed from gross symptoms, but positive identification of the fungus requires its cultivation in the laboratory and subsequent examina¬ tion of its conidial heads (fig. 7). Aspergillus spp. can be grown on potato dextrose or Sabouraud Figure 7 Conidiol heods of Asp&rgillus dextrose astars flavus. 18 Viral Disease: Sacbrood Morator aetatulas is the virus that causes sacbrood disease. It is the only common brood disease that is caused by a virus. Since sacbrood-diseased larvae are relatively free from bacteria, laboratory verification is usually based on gross symptoms and the absence ot bacteria. Positive diagnosis requires the use of a special antiserum. Affected larvae change from pearly white to gray and finally black. Death occurs when the larvae are upright, just before pupation. Consequently, affected larvae are usually found in capped cells. Head development of diseased larvae is typically retarded. The head region is usually darker than the rest of the body and may lean toward the center of the cell. When affected larvae are carefully removed from their cells, they appear to be a sac filled with water. Typically the scales are brittle but easy to remove. Sacbrood-diseased larvae have no characteristic odor. Mixed Infections Bacillus larvae produces a potent antibiotic that eliminates competition from other bacteria typically associated with honey bee larvae. For this reason, American foulbrood and European foulbrood are rarely found in the same colony, except in cases where AFB is just becoming established in colonies that already have EFB. It is not unusual to find chalkbrood and sacbrood on the same comb or on a comb with larvae infected with AFB. However, no single larva has been found to be infected with more than one disease. This is an important point to remember when selecting a sample for disease diagnosis. Diseases of Adult Bees Most diseases of adult bees are difficult to diagnose because the gross symptoms are not unique. For instance, inability to fly, unhooked wings, and dysentery are general symptoms associated with many disorders. In most cases, microscopic examination is required for proper diagnosis. Protozoan Diseases Nosema Nosema apis is the protozoan that causes nosema disease. Nosema apis spores are large, oval bodies, 4-6 pm long by 2-4 pm wide (fig. 8). The spores develop exclusively within the epithelial cells of the ventriculus of 19 the adult honey bee. Nosema disease usually manifests itself in bees that are confined; therefore, the heaviest infections are found in winter bees, package bees, bees used for pollination in green¬ houses, etc. No single symptom typifies nosema disease. Differences between healthy bees and heavily infected bees can be seen by removing the digestive tract and examining the ventriculus. The ventriculus of a healthy bee is straw brown, and the individual circular constrictions are clearly seen (fig. 9). In a heavily infected bee, the ventriculus is white, soft, and swollen, obscuring the constrictions - (White 1918). However, positive diagnosis can be made only by micro¬ scopic examination of suspect bees or their fecal material for the presence of Nosema apis spores. Samples of bees to be examined can be dried or preserved in alcohol. If the sample is partially decomposed, the presence ot yeasts and molds resembling N. apis may make an accurate diagnosis difficult. For quick, routine examinations, the abdomens from 10 or more bees are removed, placed in a dish w ith 1.0 mL water per bee abdomen, and ground with a pestle or the rounded end of a clean test tube. A cleaner preparation can be obtained by grinding free digestive tracts. A wet Figure 9. Top. Digestive tract from a healthy honey bee. Note the individual circular constrictions on the ventriculus. Bottom. Digestive tract of a honey bee with nosema disease. Figure 8. Nosema spores as they appear in a wet mount (X 400). 20 mount is prepared from the resulting suspension and examined under the high dry objective of a compound microscope. Alternatively, individual bees can be examined to obtain an approximate percentage of infected bees in a colony. Also, a quantitative measure of levels of Nosema infection can be determined using a hemocytometer as described by Cantwell (1970). Nosema can also be detected without sacrificing workers or queens, by examining their fecal material. A colony can be sampled by collecting feces of worker bees on glass plates near the hive entrance, scraping off a deposit, mixing it with water, and preparing a wet mount from the result¬ ing suspension (Wilson and Ellis 1966). Suspect queens can be held in small petri dishes or in glass tubes and allowed to walk freely. They usually defecate within 1 hour. Queen feces appear as drops of clear, colorless liquid, which are then transferred to a microscope slide with a pipet or capillary tube. A cover glass is placed over the feces before examination with a high dry objective (L'Arrivee and Hrytsak 1964). Amoeba Malpighamoeba mellificae is the organism that causes amoeba disease. Since this protozoan is found in the Malpighian tubules of adult bees, diagnosis can be made only by the removal and micro¬ scopic examination of the tubules for the presence of amoeba cysts. The cysts measure 5-8 pm in diameter and can be seen in the infected Malpighian tubules (fig. 10). The Malpighian tubules are long, threadlike projections originating at the junction of the midgut and the hindgut. The tubules can be teased away from the digestive tract with a pair of fine tweezers and then Figure 10, Cross sections of Malpighian tubules. Top, Healthy tubule; bottom, tubule containing cysts of Malpighamoeba mellificae. 21 placed in a drop of water on a microscope slide. A cover glass is posi¬ tioned over the tubules while applying uniform pressure to obtain a flat surface for microscopic examination. Malpighamoeba mellificae can be discerned using a high dry objective and then changing to the oil immer¬ sion objective for more detail. Gregarines Four gregarines (protozoans of the order Gregarinida) are associated with honey bees. Monoica apis, Apigregarina starnmeri , Acuta rousseaui, and Leidyana apis. The immature stages, or cephalonts, average about 16 by 44 pm. Cephalonts are oval and consist of two distinct body segments: the posterior segment is larger. The mature stages, or sporonts, average about 35 by 85 pm and have a reduced anterior segment (fig. 11). Gregarines are found attached to the epithelium of the midgut of adult honey bees. Gently remove the midgut from the digestive tract of a suspect bee and place it on a microscope slide in a drop of water. The - midgut can be separated from the digestive tract at the point of attachment with the proventriculus (honey stomach) and hindgut using fine tweezers and a scalpel. Gently break open the midgut with fine tweezers and a probe, and place a cover glass over the resulting suspension. Gregarines can be seen using the low-power objective of a compound microscope. Figure 11. A gregarine sporont (mature stage). 22 Flagellates Crithidia (= Leptomonas ) species are the flagellates associated with honey bees. Flagellates have been found either free in the lumen or attached to the epithelium of the hindgut and rectum of adult honey bees (Fyg 1954). Flagellates vary in size from 5 to 30 pm. Some appear as pearlike bodies with flagella; others are long threadlike forms or are round without flagella (Lotmar 1946). To look for flagellates, remove the digestive tract of a suspect bee and place it on a microscope slide in a drop of water. Then, using fine tweezers and a scalpel, separate the hindgut and rectum at the point of attachment with the midgut. Macerate the hindgut and rectum, using a fine pair of tweezers and a probe. Place a cover glass on the resulting suspension and observe under the high dry objective of the microscope. Bacterial Diseases Septicemia Pseudomonas aeruginosa (= Pseudomonas apiseptica) is the bacteiium that causes septicemia in honey bees. This disease results in the destruc¬ tion of connective tissues of the thorax, legs, wings, and antennae. Conse¬ quently, the affected bees fall apart when handled. Dead or dying bees may also have a putrid odor. Pseudomonas aeruginosa rods measure 0.5-0.8 by 1.5-3.0 pm. They are Gram-negative and occur singly, in pairs, or in short chains. A bacterial smear and Gram stain can be easily prepared after removing a wing from the thorax and dipping the wing base in a drop of water on a microscope slide. To isolate this organism, streak the base of a wing across Difco Pseudomonas isolation agar or Pseudomonas Agar F. The optimum temperature for growth is 37°C. Pseudomonas aeruginosa in culture is characterized by the excretion ot diffusible yellow-green pigments that fluoresce in ultraviolet light (wavelength below 260 nm). Septicemia disease can also be diagnosed by reproducing the disease symptoms in healthy caged bees. This is accomplished by preparing a water extract (macerate the equivalent of one suspect bee per mL of water) and inoculating healthy bees through the thorax (see Methods) or dipping them in the water extract. Bees with septicemia die within 24 hours and exhibit the typical odor and the “break apart” symptom after approxi¬ mately 48 hours. 23 Spiroplasmosis Spiroplasma species is the bacterium that causes spiroplasmosis. Spiroplasma is a helical, motile, cell-wall-free prokaryote that is found in the hemolymph of infected adult honey bees. The organism is a tiny, coiled, and sometimes branched filament 0.7-1.2 pm in diameter (fig. 12). Its length increases with age and ranges from 2 to >10 pm (Clark 1977 1978a). Spiroplasma can be seen in the hemolymph using the oil immersion objective of a phase-contrast microscope. Hemolymph can be taken from adult bees by puncturing the intersegmental membrane directly behind the first coxae with a fine capillary tube made from the tip of a Pasteur pipet. This organism can be cultured in standard mycoplasma broth medium (GIBCO) and in Singh’s mosquito tissue culture medium with 20% fetal calf serum. Viral Diseases Chronic Bee Paralysis Bees affected by chronic bee paralysis are usually found on the top bars of the combs. They appear to tremble uncontrollably and are unable to fly. Figure 12. Spiroplasma species (X 21,840) 24 In severe cases, large numbers of bees can be found crawling out the hive entrance. Individual bees are frequently black, hairless, and shiny. However, in some cases, paralysislike symptoms can be caused by toxic chemicals. Ideally, the diagnosis of paralysis disease is made using serological techniques. Since this is beyond the capability of most laboratories, diagnosis is usually made by observing symptoms in individual bees and, when possible, colony behavior. Paralysis disease can be diagnosed by reproducing the disease symptoms in caged bees. This can be done by spraying, feeding, or injecting a water extract made from the suspect bees. The extract is prepared by macerating the equivalent of one suspect bee in 1 mL of water. It is then centrituged to eliminate large suspended matter and passed through a 0.45-pm filter to remove bacteria. To feed up to 20 caged bees, mix 2 mL ot the extract with an equal volume of sugar syrup. For inoculation, each bee receives 1 pL of the extract through a dorsal abdominal intersegmental membrane (see Methods). The symptoms of paralysis should be visible after 6 days. Appropriate control bees should be treated with extracts made from healthy bees. Filamentous Virus Filamentous virus is also known as F-virus and bee rickettsiosis. This disease, previously thought to be of rickettsial origin, can be diagnosed by examining the hemolymph of infected bees using phase-contrast micros¬ copy. The hemolymph of honey bees infected with this virus is milky white and contains many spherical to rod-shaped viral particles of a size close to the limit of resolution for light microscopy. The viral particles consist of a folded nucleocapsid within a viral envelope (fig. 13) and are 0.4 by 0.1 pm (Clark 1978b). Noninfectious Disorders Noninfectious disorders can be the result of neglect, lethal genes, pollen or nectar from poisonous plants, toxic chemicals (pesticides), etc. Most often, dead or discolored pupae result from a noninfectious condition. For a good review of noninfectious diseases, see Tucker (1978). 25 Figure 13. Filamentous virus (X 67.250). Each viral particle consists of a nucleocapsid enclosed in a viral envelope Neglected Brood Normally nurse bees feed and protect the brood. However, if there is a sudden shortage of adult bees, the larvae and pupae suffer and may die of chilling, overheating, or starvation. Chilled Brood Chilling usually occurs in early spring when brood nests expand rapidly and there is a shortage of adult bees to cover all the brood. Consequently, chilled brood is found most often on the fringes of the brood area and healthy brood at the center. However, chilling can also happen during cold weather following any sudden reduction of the worker bee population. Chilled larvae and pupae are often yellowish, tinged with black on segmen¬ tal margins. They may also be brownish or black, crumbly, pasty, or watery. In extreme cases, brood cells are punctured and uncapped, and pupae are decapitated by the adult bees. It should be remembered that decapitation can also result from the larvae of the lesser wax moth. Overheated Brood The overheating of brood develops when there is a sudden loss of worker bees available to cool the colony during hot weather. Larvae that died 26 from overheating become brownish or black and are watery, pupae have a black, greasy appearance. Newly emerged adult bees may be wingless. Cappings of brood cells can appear melted, darkened, sunken, and punctured. Starved Brood Normally when there is shortage of food in a colony, larvae are removed and/or consumed by the adult bees. However, when there is a sudden shortage of adult bees available to feed the larvae, the larvae starve. Affected larvae are not restricted to the periphery of brood combs. The most striking feature of starved brood is the larvae crawling from the brood cells in search of food. Starved brood is almost always restricted to the larval stage. However, emerging bees may starve if they were stressed as pupae by chilling or overheating and if there are too few nurse bees to feed them soon after they have chewed through their cappings. These bees usually die with only their heads out of the cells and with their tongues extended. Overheated Bees Overheating in worker bees can occur when bees are confined in their hives during hot weather without proper ventilation or access to water. Bees dying from overheating crawl about rapidly while fanning their wings. They are often wet, and their wings appear hazy. In some cases, an abnormally large accumulation of dead bees may be seen at the hive entrance. Genetic Lethality Bees can also die from genetic faults during all stages of development, usually without exhibiting symptoms of known diseases. However, drone brood from laying workers and drone-laying queens often die with symptoms resembling EFB but in the absence of known pathogenic agents. Genetic lethality is the suspected cause of this condition. Plant Poisoning Poisonous plants can be a problem under certain conditions in limited areas. If a plant's nectar is poisonous, the symptoms of plant poisoning are limited to the blooming period of the plant. However, if the poison is in the pollen, the symptoms may linger as long as the pollen remains in the combs. There is no clear-cut method for differentiating between plant poisoning and pesticide poisoning. The effects of plant poisoning are 27 usually more gradual and last longer than the effects of pesticide poison¬ ing. Plant poisoning usually occurs in the same geographical area at the same time each year, whereas pesticide poisoning is indiscriminate. For a good review of poisonous plants, see Barker (1978). Some examples of plant poisoning are listed below and in table 3. Purple Brood Purple brood occurs when adult bees collect and use the pollen and nectar from Cyrilla racemiflora (titi, southern leatherwood). This “disease” is characterized by the blue or purple color of the affected larvae. Paralysis Aesculus californica (California buckeye) is probably the best known of the poisonous plants in the United States. Field bees exhibit symptoms similar to those of chronic bee paralysis; i.e., the bees are black and shiny from loss ot hair and they tremble. Also, either the eggs do not hatch or the larvae die soon after hatching. Milkweed Pollinia The pollen of milkweed (Asclepias species) is produced in pollinia (coherent pollen grains) that are attached in pairs by a slender fdament. When removed from a flower, the pollinia resemble a wishbone with pollen masses hanging from the ends. Honey bees become ensnared in the thin pollinia attachment and free themselves by pulling the pollinia from the flower (fig. 14). Honey bees often become seriously encumbered and unable to effectively fly or crawl because of the structures that remain attached to their body parts. Figure 14. Milkweed pollinia attached to honey bee. 28 30 [) o : O j O >_ Q) © >. C T) C « Pi fils >'5D£ CD ° -Q ^ 13 o o c 2 0) O "O © © © o "o § © ^ D 13 © o > D E © © Q_ © D © C 00 O' >-T3 © © >- © © 5 = > lx _c © o O) ^ © co -P 5 c o © ts >• D E co -5 ® >. 13 C O. E o o|o Q_ 13 O c .0 o Z> CL o CL 13 • ,?o © © I? S'g - 1 - LX -C o D) © 00 E ■© o o 5 c o t5 © t T3 c ^ = §■ ~0 -Q *° 0 © © E o © O © 5 -£ £XJ ! 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Pesticide Poisoning The most apparent indication of serious pesticide poisoning is the sudden loss of adult bees. This loss is characterized by the appearance of many dead and dying adult bees and sometimes pupae at the colony entrances. However, in many instances, the bees are lost in the field before returning to the colony. If the pesticide is brought back to the hive by the foragers, the nurse bees are killed when they feed on contaminated honey or pollen and the brood will exhibit symptoms of neglect or poisoning. The symptoms ot poisoned honey bees often depend on the class of pesticide involved (table 4). A residue of a pesticide may be present as the original pesticide, or as an identifiable degradation product, or both. Frequently, the amount of residue is extremely small. Pesticide analysis consists primarily of the following (from Wilson et al. 1980): (1) Blending and extraction of the biological material (such as bees or pollen) with a suitable solvent system to maximize the recovery of suspected pesticides and their metabolites. This eliminates the bulk of the biological substrate. (2) A series ot liquid-liquid extractions and column chromatographs to further separate the residues from other materials of biological origin. A very fast method for cleanup is gel permeation chromatography. Large molecules of biological origin emerge from the column first, thereby directly trimming the crude extract down to a much cleaner sample. (3) Detection of residues at the highest possible sensitivity to avoid interference from substances not previously removed. The most popular detection system is gas-liquid chromatography, wherein the residue- containing sample is volatilized and chromatographed as a vapor. Thin- layer and paper chromatography are also useful for establishing the identity ot pesticide residues. Visualization for detection usually involves spraying the chromatograms with a chemical that reacts with the pesticide or metabolite to produce a characteristic color. Other methods used for residue analysis include mass spectroscopy and fluorometric techniques. The residue must be capable of absorbing visible light or ultraviolet light. The intensity of the reemitted light is measured at some suitable wave¬ length and compared with standards. 30 05 c 'c o o a 4— o CO £ o a E > oo c . o > © 05 o P o x © c o o c .© o ~ m « CO © © 0)15 -IS +2 Q) E >E © © 2 O E D 4= r> 5 3 o O c o o a 4— o © D) O c © © £ © •© © 13 © © P P C C © O-g oP a) a - n 10 ^ © © 05-2 > © © © O' CL io « © >-P 2 > P 0 n a ° -p E © --f © o 13 O > D) O C B 5 © EP © a x u> x © X P co 0 8>o o 05 T3 =§ ® P P © © >- P)C O O o o o © p © p E © © o D ■C © -. « .X Q) 0)0 © .Q 0 15 5£p -O O) © C o E © c . 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O CO © a E o x © o o X © E p c o X o o o c «o O o 9- X p p> c w © E ay o 5 o EP o P p c o c p 9 © p pp c X X.E 8 "o Oo POx x © E c o X p X C p- PE| o P-8 E 8 © ~ .co PC® O © •!= N ^ >- a CL - CO HO 0^-0 X © o o 2x2 a a o O'® o a E ®9 - © c c P X © p a o X (X o a Z q © . c © o O P o c p = I o' o o C P T3 O o < X o C 2 O o 32 El §8 SI 0 O co . ^ /~N Zo o p -2 2 o © UE © o p ' L_ o o P x c © -E E E 05 II Is Q ci c c o E 5E o © X E © p £ O o © c o X o X CO u~> O Z Q a x P n o C o O c z O Q x . © % E p£ c D I X •E®1 £ ■ ■ p (11 10 = P C P o •= ill ifs ir © O x 9 p © .E o 8 ^ ■F ° c P O P g c o > co r. o c -1C < E Jr © °P ^ O PZ D o 61 8 9 05 O Oa c P o © p o b c o 52 U x © o £ o p b O > c © X p o o o g c o o CD 10 gm B C Acarapis woodi: Apodeme (A) 2/3 propodosoma (P); coxal plate (B) shallow indentation; tarsal joints (C) <10 gm. Figure 15. Morphological characters separating Acarapis species. 32 Honey Bee Tracheal Mite (Acarapis woodi) The female Acarapis woodi , or honey bee tracheal mite, is 143-174 urn long and the male 125-136 pm. The body is oval; widest between the second and third pair of legs; and whitish or pearly white with shining, smooth cuticle. A few long hairs are present on the body and legs. This mite has an elongate, beaklike gnathosoma with long, bladelike styles (mouthparts) for feeding on the host. The population of Acarapis woodi may vary seasonally. During the period of maximum bee population, the number of bees with mites is reduced. The likelihood of detecting tracheal mites is highest in the fall. In sampling for this mite, one should try to collect either moribund bees that may be crawling near the hive entrance or bees at the entrance as they are leaving or returning to the hive. These bees should be placed in 70% ethyl or methyl alcohol as soon as they are collected. One should not collect bees that have been dead for an unknown period because they are less than ideal for the diagnosis of tracheal mites. No one symptom characterizes this disease. An affected bee could have disjointed wings and be unable to fly, or have a distended abdomen, or both. Absence of these symptoms does not necessarily indicate freedom from mites. Positive diagnosis can be made only by microscopic examina¬ tion of the tracheae; since only Acarapis woodi is found in the bee tracheae, this is an important diagnostic feature. A healthy trachea appears cream color or white. The trachea of a severely infested bee has brown or black blotches with crustlike lesions and is obstructed by many mites in different stages of development (fig. 16). The trachea must be examined carefully for the presence of mites. The trachea may not always be discolored when mites are present, and a cloudy or discolored trachea does not always contain mites. Methods for diagnosing Acarapis woodi are listed below. Each of these methods has its advantages and disadvantages. Method 1 Pin the bee on its back and remove the head and first pair of legs by pushing them off with a scalpel or razor blade in a downward and forward motion (fig. 17). Using a dissecting microscope, remove the first ring of the thorax (tergite of prothorax) with forceps. This exposes the tracheal 33 ¥ Figure 16, Trachea containing mites (X 40). trunks in the mesothorax (fig. 18). When the infestation is light, it is necessary to remove the trachea. Place the trachea in a drop of lactic acid on a glass slide for clearing, and cover with a cover glass for examination at X 40-100 on a compound microscope. Method 2 Grasp the bee between your thumb and forefinger and remove the head and first pair of legs. Then with a scalpel, razor blade, or fine pair of scissors, cut a thin transverse section from the anterior face of the thorax in such a way as to obtain a disk. Place the disk on a microscope slide and add a few drops of lactic acid. This makes the material more transparent and also helps to separate the muscle. With the aid of a dissecting microscope, carefully separate the muscles, remove the trachea, and examine the preparations as in method 1. This procedure is recommended for quick examination of a few bees. Figure 17, Positioning a bee for dissection. 34 Figure 18. Location of Esophaaus _ Trachea Salivary the trachea in glands the thorax. Wing muscles Tergite of Point of prothorax “collar" Method 3 Cut a few thoracic disks as described in method 2, place them on a slide, and add a few drops of 10% potassium hydroxide (KOH). Heat the slide gently for 1-2 minutes (do not boil), cover with a cover glass, crush the disks lightly, and examine microscopically. This procedure is advanta¬ geous when the bees have been dead for some time. Method 4 Prepare transverse-section disks from the thoraces of 50 honey bees as described in method 2, place them in 5% KOH, and incubate at 37°C for 16-24 hours. The KOH dissolves the muscle and fat tissue, leaving the trachea exposed. Then examine the disk-trachea suspension under a dissecting microscope. Remove suspicious tracheae from the disks and examine the tracheae microscopically (X 40-100). This procedure is recommended for large samples of bees. Method 5 Remove the heads, abdomens, wings, and legs from 20-200 thoraces and place them in a homogenizing jar with 25 mL of water. Homogenize three times for several seconds at 10,000 rpm, using just enough water to rinse the inside of the jar. Then strain the suspension through a 0.8-mesh sieve and rinse with water. The final volume of the filtrate should be about 50 mL. Centrifuge the filtrate at about 1,500 g for 5 minutes and discard the supernatant. Then add a few drops of lactic acid to the preparation, and allow it to stand for 10 minutes. Finally, place the sediment on a slide for examination. In this method, a microscope with an oil immersion objec¬ tive is required to correctly identify Acarapis woodi because other mites associated with honey bees are morphologically similar. This technique is described by Colin et al. (1979). Method 6 In the flotation method (Camazine 1985), bees cannot be stored or killed in alcohol. For cleanest preparations, remove the head, wings, legs, and abdomen (saving only the thoraxes) of recently killed bees. This removal is easily done using one’s fingers when the bees are frozen. Place 25-100 bees in a household blender with enough water added to cover the blades. Blend the preparation for no more than 15 seconds, just until the thoraces are broken apart. (Blending for longer periods will pulverize the tracheae.) Pour the resulting mixture into a series of test tubes (2-3 cm in diameter). Most of the denser muscle fibers and cuticular fragments fall to the bottom while the tracheae and air sacs float, forming a thin whitish layer on the surface of the water. Suction off this layer with a pipette, place on one or more slides, and cover with cover slips. Examine the slides under a compound microscope at X 100-250. Examine the slide in a systematic manner for darkened, blotchy, and discolored tracheae and for undamaged tracheae that may also contain mites and eggs. Method 7 In the modified methylene blue staining technique (Peng and Nasr 1985). prepare transverse-section disks from the thoraces of 50 bees as described in method 2. Place the disks in a beaker of 8% KOH solution, and heat to boiling with continuous gentle stirring of the disks. Remove the solution from the heat and continue stirring until the soft tissues inside the disks are dissolved and cleared (about 10 minutes). Excessive stirring and heating will damage the specimens and subsequently reduce the color intensity of the mites. Recover the disks from the KOH by filtration through a perforated Tissue-Tek processing capsule. After filtration, cover the processing capsule with a lid. place in a beaker, and wash with tap water to remove the remaining KOH. After washing, transfer the processing capsule to a modified methylene blue staining solution (prepared by first dissolving 1% aqueous methylene blue and then addins sodium chloride to make a 0.85% sodium chloride solution). Immerse the capsule in that solution for 5 minutes and then in distilled water for 2-5 minutes; finally, rinse the capsule with 70% ethyl alcohol. Examine the disks for stained mites within the tracheae under a dissecting microscope at X 10-25. Method 8 Differentiation of live mites from dead mites (Eischen et al. 1986) is the method of choice for evaluating chemicals used to control tracheal mites. 36 Anesthetize live bees with carbon dioxide and remove the abdomens with a scalpel to prevent being stung during examination. Remove the head and first pair of legs of each bee by holding the bee on its back and gently pushing this section oft with a downward and forward motion. Place each bee, held in this position, under a dissecting microscope, and remove the first ring of the thorax with fine forceps. This exposes the tracheal attachment to the thoracic wall, which is often the only location of mites in a light infestation. Remove tracheae that appear abnormal with tweezers and transfer to a glass slide containing a thin film of glycerol. Then dissect the tracheae using a pair of fine needle probes. Mites are consid¬ ered dead if they do not move; also, dead mites often appear discolored and desiccated. Living mites have a translucent gray or pearl color and move within a few seconds after dissection. Method 9 For serodiagnosis, Ragsdale and Furgala (1987) produced an antiserum against extracts of Acarapis woodi-infested tracheae to be used as the primary antibody in a direct enzyme-linked immunosorbent assay (ELISA). Ragsdale and Kjer (1989) improved the ELISA technique, making it as reliable as dissection for the detection of A. woodi. Their ELISA is accurate, sensitive, reproducible, cost effective, rapid, and easy to use. Varroa jacobsoni The mite Varroa jacobsoni can be found on adult bees, on the brood, and in hive debris. The most severe parasitism occurs on the older larvae and pupae, with drone brood being preferred to worker brood (Ritter and Ruttner 1980). In heavy infestations, pupae may not develop into adult bees. The adults that do emerge may have shortened abdomens, mis¬ shapen wings, and deformed legs and may weigh less than healthy bees (De Jong et al. 1982b). The adult female mite is oval and flat, about 1.1 mm long and 1.5 mm wide, and pale to reddish brown; it can easily be seen with the unaided eye. The mites attach to the adult bee between the abdominal segments or between body regions (head, thorax, abdomen) and are therefore difficult to detect. However, they can be easily recognized against the white surface of pupae. Male mites are considerably smaller and are pale to light tan (Delfinado-Baker 1984). The life cycle of V. jacobsoni is summarized in figure 19. 37 10 Mites transfer through close contact between bees. 1 Adult bee. with Varroa feeding on hemolymph 1 1 6 1 -6 eggs developing N from egg to larva to protonymph to deutonymph adult female egg | / 9 Adult females leave cell with emerging bee. Male and immature stages stay in cell. 8 Mating within cell 7-8 days adult female Figure 19. Life cycle of Varroa jacobsoni. (Courtesy of Roger A. Morse.) 38 It is important to note that the bee-louse, Braula coeca, resembles Varroa jacobsoni in size and color. However, Braula , being an insect, has six legs that extend to the side (fig. 20). Varroa , an arachnid, has eight legs that extend forward (fig. 21). When sampling, remember that the number and location of mites in a colony vary according to time of year. The number of mites is lowest in spring, increases during summer, and is highest in fall. During spring and summer, most mites are found on the brood (especially drone brood). In late fall and winter, most mites are attached to adult worker bees. Methods of Examining Adult Honey Bees For a sample of adult honey bees, 500 to 1000 bees should be collected. This can be done by brushing honey bees off the comb through a large-mouthed funnel (of paper or cardboard, etc.) into a container or by using a modified portable car vacuum cleaner. Individual honey bees can be examined with or without the aid of a hand lens or a dissecting microscope. When the mites are moving about on a bee, they are fairly easy to detect; but once they attach themselves between segments, they are difficult to find. Mites can be detected and collected by three methods, as follows: Shaking Method Varroa jacobsoni can be dislodged by shaking the bees in liquids such as hot water, alcohol, detergent solution, hexane. 39 Figure 20. Braula coeca, dorsal view. Figure 21. Varroa jacobsoni, ventral view. gasoline, or diesel fuel. We recommend 70% alcohol (ethyl or isopropyl) because some of the other materials are dangerous or difficult to use. The alcohol kills and preserves the bees for other purposes, such as examina¬ tion tor Acarapis woodi. De Jong et al. (1982a) found that hand-shaking bees in alcohol for 1 minute dislodged about 90% of the mites and that mechanical shaking on a rotary shaker for 30 minutes removed 100% of the mites. The mites are collected by passing the bees and alcohol through a wire screen (8- to 12-mesh) to remove the bees and then sieving the alcohol through a 50-mesh screen or cotton cloth. The screen or cloth is then examined for mites. 40 Ether Method This technique is a rapid and efficient detection method in the field and avoids the handling, shipping, and time-consuming procedures associated with shaking adult bees in alcohol or other solvents. The bees (500-1,000) are collected in ajar and anesthetized with ether delivered from an aerosol can (this aerosol product is sold in auto-parts stores as an aid to start engines). A 1- to 2-second burst of material is adequate. The bees are then rotated in the jar for about 10 seconds. The majority of mites will have dislodged from their hosts and should be adhering to the inside wall of the jar. To complete the process, the bee sample is deposited on a white surface and spread around. This should cause any remaining mites to fall onto the white substrate. The bees should be examined immediately after the application of ether because the mites tend to stick to the bees if left in the jar for more than a few minutes. Alternatively, the bees can be left in the jar to which alcohol is added for laboratory shaking and preservation. Heating Method Live adult honey bees can be shaken into a wire-based cage and placed in an oven over white paper. The bees are heated for 10-15 minutes at 46°- 47°C. Then Varroa jacobsoni, if present, can be observed on the white paper (Crane 1978). Methods of Examining Brood To look for mites on brood, the pupae (preferably drone) are examined. Varroa jacobsoni can be easily seen against the white surface of worker or drone pupae after they are removed from their cells. It is suggested that a minimum of 100 drone pupae per colony be examined. The pupae can be collected by one of the following methods: • The classic method of pupal collection is to uncap each cell and then remove the pupae with forceps or a hive tool. • Groups of pupae can be quickly and easily removed from their cells by inserting a capping scratcher at an angle through the cappings and lifting the brood and cappings upward (Szabo 1989). • With a long-bladed knife, the caps are sliced off an area of brood 4-6 square inches. The comb (frame) is then sharply jarred on a hard, flat, white surface such as a hive top. The brood will fall onto the white surface, and the mites can be easily observed. 41 • The brood comb can be incubated at 37°C, followed by examination of all the emerged bees and remaining brood. Methods of Inspecting Hive Debris Debris in a hive (such as wax particles, pollen, dead bees and brood, and mites) normally falls to the hive floor and is removed by house-cleaning bees during warm weather. This material can be collected and examined for the presence of Varroa jacobsoni as follows: • The collection of hive debris can be facilitated by white construction paper on the hive floor. The paper is stapled under a wood (1/4-inch) and wire (8- to 12-mesh) frame, which protects the paper and debris from the bees. The paper is examined for mites, which can be easily seen against the white background. A magnifying glass or dissecting microscope can be helpful in locating the mites in the debris. Sticky boards or shelf paper (with the adhesive surface exposed) instead of construction paper will help hold the debris. • The acaricides used to treat mite infestations can also be applied to bee colonies in combination with the paper method to detect Varroa jacobsoni. Apistan is currently approved and available for this purpose. After treatment, the mites drop to the paper and can be easily detected. It is important that the paper have a sticky surface (see previous paragraph) to hold any recovering mites. • A flotation method can be used to examine debris for Varroa jacobsoni. Hive debris is placed in ajar or pan and covered with 98% alcohol. The mites float to the surface while the heavier debris sinks (Ritter and Ruttner 1980). • Mites can sometimes be collected in dead bee or pollen traps attached to the colonies. Tropilaelaps clareae The distribution of Tropilaelaps clareae (fig. 22) is still restricted to Southeast Asia. These mites also parasitize adult bees and brood, and they have been reported to infest colonies infested with Varroa jacobsoni (Deltinado-Baker and Aggarwal 1987). Female mites are about 1 mm long and 0.6 mm wide; the male is slightly smaller. These mites are difficult to detect because of their small size and their brownish color, which blends perfectly with brood cappings and comb. Tropilaelaps 42 Figure 22, Tropilaelaps clareae, ventral view. clareae can be found by observing under a magnifying glass or a dissect¬ ing microscope a brood comb suspected of being infested. In the field, when the comb is hit on a light-colored surface, dislodged mites may be seen moving on that surface. The mites can be picked up with a fine brush moistened with alcohol. Pests Wax Moths The greater wax moth. Galleria mellonella (fig. 23), is the most serious pest of honeycombs. Comb damage can also be caused by the lesser wax moth, Achroia grisella (fig. 23), and the Mediterranean flour moth, Anagasta kuehniella. These moths are an especially serious problem in tropical and subtropical climates, where warm temperatures favor the rapid development of the moths. Female greater wax moths lay their eggs in a cluster, usually in the cracks or between the wooden parts of the hive. The larvae of the moths are the destructive stage. They actually obtain nutrients from honey, castoff pupal skins, pollen, and other impurities found in the beeswax, but not the beeswax itself. Consequently, older combs are more likely to be infested than new combs or foundation. Bee-louse (Braulo coeca ) Braula coeca , or bee-louse (fig. 20), is actually not a louse but a wingless fly that feeds on honey. No detrimental effect on adult bees has been attributed to the bee-louse, but its larvae can damage the appearance of 43 Figure 23. Greater wax moth. Galleria mellonella (top), and lesser wax moth, Achroia grisella (bottom) (not to scale). These species are easily distinguished from each other by shape of the wings and comparative sizes. Greater wax moth is about two times larger than lesser. comb honey. Adult bee-lice can be found on adult workers and queens. It is important to note that Braula coeca resembles Varroa jacobsoni in size and color. However. Braula, being an insect, has six legs that extend to the side. Varroa , an arachnid, has eight legs that extend forward. Meliffiphis alvearius Melittiphis alvearius is a little-known mite that is associated with adult honey bees but is not considered to be a pest. It is unlikely that M. alvearius would be confused with other mites found in honey bee colo¬ nies. The adult female mite is ovate, flattened dorsoventrally, 0.79 mm long and 0.68 mm wide, brown, and well sclerotized with numerous stout and spinelike setae. It is included here because it was found in California during a survey tor Varroa and because of increasing reports on its distribution (Delfinado-Baker 1988). 44 "V Appendix A. Directions for Sending Diseased Brood and Adult Honey Bees for Diagnosis The accuracy of the diagnosis of any bee disease depends on the condition of the sample. Mail the sample in a wooden or heavy cardboard box. The sample can be loosely wrapped in a paper bag, paper towel, newspaper, etc. Avoid wrappings such as plastic bags, aluminum foil, waxed paper, tin, or glass because they allow fungi to grow on the samples. Samples are accepted from anyone and in most cases are processed within one working day. How to Address and Package Samples • Send all samples to Bee Disease Diagnosis USDA, ARS, Bee Research Laboratory Building 476, BARC-East 10300 Baltimore Avenue Beltsville, MD 20705-2350. • A short description of the problem along with your name and address should accompany the sample. Also, your name and address should be plainly written on the top ot the box. Samples of Brood • The sample of comb should be about 4 square inches. • The sample should contain as much of the dead or discolored brood as possible. • NO HONEY SHOULD BE PRESENT IN THE SAMPLE. • If a comb cannot be sent, the probe used to conduct a ropiness test may contain enough material for tests. The probe can be wrapped in paper and sent to the laboratory in an envelope. However, this method is unsatisfac¬ tory for verification of sacbrood disease, which requires special antisera. We rely on gross symptoms for this diagnosis. 45 Samples of Adult Honey Bees • Send at least 100 bees in a sample. • If possible, select bees that do not appear normal or that died recently. Decayed bees are not satisfactory for examination. • Bees submitted for the identification of mites should be placed in 70% ethyl or methyl alcohol as soon as possible after collection. These samples should be in leak-proof containers. 46 appendix B. Identification of Africanized Honey Bee 3RL Protocol for Identification of Africanized Honey Bee -ABIS, as described in the “Fast Africanized Bee Identification System FABIS) Manual” (Sylvester and Rinderer 1987), will be used as a screening method to identify European bees. The full 26-character -norphometric analysis will be conducted on all samples of honey bees not identified as European by FABIS. Therefore, when the Bee Research Laboratory (BRL) declares a sample Africanized, the identification is the result of the full 26-character morphometric analysis and is definitive. Submission of Honey Bees to BRL for Identification of Subspecies It is assumed that most honey bee samples submitted for identification of subspecies will be from Animal and Plant Health Inspection Service, Plant Protection and Quarantine (APHIS-PPQ). Samples will also be accepted from State regulatory personnel by prior arrangement. A sample of 100 bees should be submitted. Anyone submitting suspected Africanized honey bees should understand our priorities for identification and sample submission. They are as follows: Priority No. I : Swarms (colonies) anywhere in the United States involved in severe stinging incidents. Priority No. 2 : Any swarms (colonies) in the United States that have been pretested and identified as non-European by FABIS. 47 Honey bees submitted for identification should be placed in 70% ethyl or methyl alcohol as soon as possible after collection. Each sample should consist of at least 100 workers in leak-proof containers and be sent directly to Bee Disease Diagnosis USDA, ARS, Bee Research Laboratory Building 476, BARC-East 10300 Baltimore Avenue Beltsville, MD 20705-2350. A short history of the problem along with the sender’s name, address, and telephone number should accompany the sample. If results are urgently required, please advise the Laboratory by telephone (301-344-2205) of the shipment and send the sample via Express Mail. References Atkins, E.L. 1975. Injury to honey bees by poisoning. In Dadant and ions, eds., The Hive and the Honey Bee, pp. 663-696. Dadant and Sons, Jamilton, Illinois. iailey, L. 1959. An improved method for the isolation of Streptococcus iluton and observations on its distribution and ecology. Journal of Insect 3 athology 1:80-85. Bailey, L. 1981. Honey bee pathology. 124 pp. Academic Press, Inc., ^ondon. Bailey, L., and M.D. Collins. 1982a. Taxonomic studies on Strepto- :occus platan. Journal of Applied Bacteriology 53:209-213. Bailey, L„ and M.D. Collins. 1982b. Reclassification of Streptococcus oluton (White) in a new genus Melissococcus pluton. Journal of Applied Bacteriology 53:215-217. Bailey, L., and D.C. Lee. 1962. Bacillus larvae: Its cultivation in vitro and its growth in vivo. Journal of General Microbiology 29:711-717. Barker, R.J. 1978. Poisoning by plants. In R.A. Morse, ed„ Honey Bee Pests, Predators, and Diseases, pp. 275-296. Cornell University Press, Ithaca and London. Burnside, C.E., and G.H. Vansell. 1936. Plant poisoning of bees. U.S. Department of Agriculture, Bureau of Entomology and Plant Quarantine. E-398, December 1936. Camazine. S. 1985. Tracheal flotation: A rapid method for detection of honey bee acarine disease. American Bee Journal 125:104-105. Cantwell, G.E. 1970. Standard methods for counting nosema spores. American Bee Journal 110:222-223. Clark, T.B. 1977. Spiroplasma sp„ a new pathogen in honey bees. Journal of Invertebrate Pathology 29:112-113. Clark, T.B. 1978a. Honey bee spiroplasmosis, a new problem for beekeepers. American Bee Journal 118:18-19,23. 49 Clark, T.B. 1978b. A filamentous virus of the honey bee. Journal of Invertebrate Pathology 32:332-340. Colin, M.E., J.P. Faucon, A. Giauffret, and C. Sarrazin. 1979. Anew technique for the diagnosis of acarine infestation in honeybees. Journal of Apicultural Research 18:222-224. Crane, E. 1978. The Varroa mite. Bee World 59:164-167. De Jong, D., D. De Andrea Roma, and L.S. Goncalves. 1982a. A comparative analysis of shaking solutions for the detection of Varroa jacobsoni on adult honeybees. Apidologie 13:297-306. De Jong, D„ P.H. De Jong, and L.S. Goncalves. 1982b. Weight loss and other damage to developing worker honeybees from infestation with Varroa jacobsoni. Journal of Apicultural Research 21:165-167. Delfinado-Baker, M. 1984. The nymphal stages and male of Varroa jacobsoni Oudemans - a parasite of honey bees. International Journal of Acarology 10:75-80. Delfinado-Baker, M. 1988. Incidence of Melittiphis alvearius (Berlese), a little known mite of beehives, in the United States. American Bee Journal 128:214. Deltinado-Baker, M., and K. Aggarwal. 1987. Infestation of Tropilaelaps clareae and Varroa jacobsoni in Apis mellifera ligustica colonies in Papua New Guinea. American Bee Journal 127:443. Deltinado-Baker, M., and E.W. Baker. 1982. Notes on honey bee mites of the genus Acarapis Hirst (Acari: Tarsonemidae). International Journal of Acarology 8:211-226. Dingman, D.W.. and D.P. Stahly. 1983. Medium promoting sporulation of Bacillus larvae and metabolism of medium components. Applied and Environmental Microbiology 46:860-869. Eischen, F.A., J.S. Pettis, and A. Dietz. 1986. Prevention of Acarapis woocii infestation in queen honey bees with amitraz. American Bee Journal 126:498-500. 50 'yg, W. 1954. Uber das Vorkommen von Flagellaten in Rectum der lonigbiene (Apis mellifera L.). Mitteilungen der Schweizerischen ■ntomologischen Gesellschaft 27:423-428. Jochnauer, T.A. 1953. American foulbrood and chemical controls. Report of the Iowa State Apiarist, pp. 10-14. Jochnauer, T.A., and J. Corner. 1974. Detection and identification of iacillus larvae in a commercial pollen sample. Journal of Apicultural Research 13:264-267. jochnauer, T.A., B. Furgala, and H. Shimanuki. 1975. Diseases and ;nemies of the honey bee. In Dadant and Sons, eds.. The Hive and the Honey Bee, pp. 615-662. Dadant and Sons, Hamilton, Illinois. daynes, W.C. 1972. The catalase test: An aid in the identification of Bacillus larvae. American Bee Journal 112:130-131. Holst, E.C. 1946. A single field test for American foulbrood. American Bee Journal 86:14, 34. Kostecki, R. 1969. Studies on improvement of control of American foulbrood of the honey bee. (From Polish.) Pszczelnicze Zeszyty Naukowe 13:97-135. L'Arrivee, J.C.M., and R. Hrytsak. 1964. Coprological examination for nosematosis in queen bees. Journal of Insect Pathology 6:126-127. Lochhead, A.G. 1937. The nitrate reduction test and its significance in the detection of Bacillus larvae. Canadian Journal of Research 15:79-86. Lotmar, R. 1946. Uber Flagellaten und Bakteria im Dunndarm der Honigbiene. Schweizerische Bienen-Zeitung, Beih. 2:49-76. Michael, A.S. 1957. Droplet method for observation of living unstained bacteria. Journal of Bacteriology 74:831-832. Michael, A.S. 1960. A new technique for studying bee diseases. XVII International Beekeeping Congress, Bologna-Roma, 1958, Official Report, Second Volume, pp. 73-77. Imola Tipografia Galeati, 1960. 51 ¥ Otte, E. 1973. A contribution of the laboratory diagnosis of American toulbrood of the honey bee with a particular reference to the immuno¬ fluorescence method. Apidologie 4:331-339. Peng, Y-S., and M.E. Nasr. 1985. Detection of honey bee tracheal mites (Acarapis woodi) by simple staining techniques. Journal of Invertebrate Pathology 46:325-331. Peng, Y-S., and K-Y. Peng. 1979. A siudy on the possible utilization of immunodiffusion and immunofluorescence techniques as the diagnostic for American foulbrood of honeybees (Apis mellifera). Journal of Invertebrate Pathology 33:284-289. Pinnock, D.E., and N.E. Featherstone. 1984. Detection and quantification of Melissococcus pluton infection in honeybee colonies by means of enzyme-linked immunosorbent assay. Journal of Apicultural Research 23:168-170. Ragsdale, D.W., and B. Furgala. 1987. A serological approach to the detection of Acarapis woodi parasitism in honey bees using an enzyme- linked immunosorbent assay. Apidologie 18:1-9. Ragsdale, D.W., and K.M. Kjer. 1989. Diagnosis of tracheal mite (Acarapis woodi Rennie) parasitism of honey bees using a monoclonal based enzyme-linked immunosorbent assay. American Bee Journal 129:550-553. Ritter, W., and F. Ruttner. 1980. Diagnoseverfahren (Varroa). Allgemeine Deutsche Imkerzeitung 5:134-138. Shimanuki, H. 1963. In vitro and in vivo studies of Bacillus larvae. Ph D. dissertation, Iowa State University of Science and Technology. 150 pp., Ames, Iowa. Shimanuki H., and D.A. Knox. 1988. Improved method for the detection of Bacillus larvae spores in honey. American Bee Journal 128:353-354. Sturtevant, A.P. 1932. Relation of commercial honey to the spread of American foulbrood. Journal of Agricultural Research (Washington DC) 45:257-285. Sturtevant, A.P. 1936. Quantitative demonstration of the presence of spores of Bacillus larvae in honey contaminated by contact with American 52 Dulbrood. Journal of Agricultural Research (Washington DC) 52: 97-704. lylvester, H.A.. and T.E. Rinderer. 1987. Fast Africanized bee identifica- ion system (FABIS) manual. American Bee Journal 127:511-516. ; Z abo, T.I. 1989. The capping scratcher: A tool for detection and ontrol of Varroa jacobsoni. American Bee Journal 129:402-403. loschkov, A.. T. Vallerianov, and A. Tomov. 1970. Die mmunofluoreszenzmethode und die Schnelle und Spezifische Diagnotik ier Amerikanischen Faulbrut bei der Bienenbrut. Bulletin Apicole de Documentation et dTnformation 13:13-18. rucker, K.W. 1978. Abnormalities and noninfectious diseases. //; R.A. Vlorse, ed.. Honey Bee Pests. Predators, and Diseases, pp. 257-274. Cornell University Press, Ithaca and Fondon. White, G.F. 1912. The cause of European foulbrood. U.S. Department of Agriculture, Bureau of Entomology Circular 157, 15 pp. White, G.F. 1918. Nosema disease. U.S. Department of Agriculture Bulletin 780, 59 pp. White, G.F. 1920. European foulbrood. U.S. Department of Agriculture Bulletin 810, 39 pp. Wilson, C.A., and L.L. Ellis. 1966. A new technique for the detection of nosema in apiaries. American Bee Journal 106:131. Wilson, W.T. 1970. Inoculation of the pupal honeybee with spores of Bacillus larvae. Journal of Apicultural Research 9:33-37. Wilson, W.T., and W.C. Rothenbuhler. 1968. Resistance to American foulbrood in honey bees. VIII. Effects of injecting Bacillus larvae spores into adults. Journal of Invertebrate Pathology 12:418-424. Wilson, W.T., P.E. Sonnet, and A. Stoner. 1980. Pesticides and honey bee mortality. In Beekeeping in the United States, pp. 129-140. U.S. Department of Agriculture Handbook 355. Zhavnenko, V.M. 1971. Indirect method of immunofluorescence in the diagnosis of foulbrood (American and European) (in Russian). Veterinariya (Kiev) 8:109-111. United States Department of Agriculture Forest Service Agriculture Handbook No. 691 UNIVERSITY OF ILLINOIS LAW LIBRARY MAY 2 0 1b j FEDERAL DEPOSITORY A/. 76 : V Cl) ■ t UNIVERSITY OF ILLINOIS tSBlCULTURE LIBRARY ROOT Di s easE by Charles G. Shaw III Rocky Mountain Forest and Range Experiment Station Fort Collins, Colorado and Glen A. Kile Commonwealth Scientific and Industrial Research Organization Division of Forestry and Forest Products Hobart, Tasmania, Australia Agriculture Handbook No. 691 Forest Service United States Department of Agriculture Washington, D.C. March 1991 AGRICULTURE LIIRARY JUN 1 2 1991 UNIVERSITY OF ILUHOff Preface Armillaria root disease has been the object of intensive basic and applied study by pathologists, physiologists, taxonomists, and others since Robert Hartig published his classical work in 1874. Even with this immense collective effort, persistent confusion has obscured the real significance of Armillaria as a pathogen. Only re¬ cently have pathologists accepted that Armillaria com¬ prises numerous species with distinct distributions and pathogenicities. This treatment resolves many contra¬ dictory claims and observations made about Armillaria species and the often serious root diseases they cause. Armillaria is, however, more than just a serious patho¬ gen. Economic importance aside, Armillaria possesses many fascinating biological attributes that are broached in this volume. These include bioluminescence; antibi¬ otic and alcohol production; multiple morphological forms including rhizomorphs; in vitro fructification; peculiar mycorrhizal associations with the roots and tubers of some achlorophyllous plants; an unusual nuclear cycle; and others. In our view, the amplitude of this variability makes species of Armillaria well suited as experimental organisms for studying fungal devel¬ opment, physiology, genetics, and speciation. Through this volume we strive to synthesize the avail¬ able information on the taxonomy, physiology, and life history of Armillaria spp. This material is further devel¬ oped to clarify the impacts, dynamics, management, and control of the root diseases caused by various spe¬ cies of Armillaria in diverse natural and exotic forests, orchards, and amenity plantings throughout the world. The book begins with a discussion of the taxonomy and nomenclature of Armillaria species. Through this treat¬ ment, we not only learn how to correctly refer to these organisms but also discover why so much confusion has surrounded their taxonomy and nomenclature. This leads into chapter 2 wherein the concept and sig¬ nificance of biological species are explored as are the sexual patterns and life cycle of the fungus. The nutri¬ tional, biochemical, and physiological requirements of the fungus and the biochemical basis for its interactions with hosts are considered in chapter 3. Attributes of inoculum and the infection process are discussed in chapter 4. Disease symptoms and diagnosis, both on individual trees and in stands, are treated in chapter 5. Pathogenicity and various ways of assessing it are dis¬ cussed in chapter 6. The next three chapters consider the role of stress factors in promoting disease and ad¬ dress disease development in natural forests and manmade plantations. Chapter 10 introduces math¬ ematical modeling as a means to quantify disease de¬ velopment and to predict the consequences of various management actions. Chapter 11 presents management and control methods, including recent information on antagonistic organisms. This book was conceived through discussions on Armillaria held among members of the International Union of Forestry Research Organizations' Working Party on Root and Butt Rots of Forest Trees. This is one of the largest, oldest, and most active IUFRO groups. Many members of that group have authored chapters for this book; many others provided ideas, advice, and encouragement. The volume stands as a tribute to the spirit of international cooperation in forestry research that is fostered by IUFRO. The worldwide interest in, and importance of, Armillaria root disease is reflected by the contributions to this volume: 24 authors from 9 nations. Managing not only the vast amount of manuscript provided by these authors but also their often contrasting ideas, opinions, and personal reflections into a single volume with some meaningful composition and structure be¬ came our unique challenge. iii Our ambition has been and remains the presentation of accurate information about Armillaria. Clarity of ex¬ pression became the driving objective we used as a final arbiter for many difficult decisions. We wanted to remove as many potential disruptions to smooth read¬ ing as possible yet preserve an international character. Thus, we retained words and expressions unique to certain countries or cultures, but we imposed uniform spelling and punctuation standards throughout all chapters. We also sidestepped standard botanical no¬ menclature. For general discussion in the text, we chose where pos¬ sible to use common names as established in standard references. Coping with genus, specific epithet, au¬ thorities and multiple revisions, plus abbreviations, parentheses, and brackets proved extraordinarily te¬ dious during manuscript preparation and revision. Ultimately, we judged the nomenclature system to be too clumsy to meet our objective of clear expression. We met the obligation for scientific accuracy by adding an Appendix which cites in alphabetical order both Latin and common names with the appropriate stan¬ dard references. To overcome nomenclatural problems with reference to various Armillaria species, we used specific epithets only where investigators have identi¬ fied their isolates. We used the generic term " Armillaria " where identity is uncertain. The timing of this work seems particularly important as our knowledge of these organisms and the diseases they cause has increased markedly in recent years. We hoped that by compiling the information at this time we could stimulate and help focus further research while also providing a basis for wise and informed management of Armillaria diseases. In addition to an analysis, synthesis, and consolidation of the vast literature that has accumulated, as well as the advancement of concepts and insights to assist future research on Armillaria, this volume celebrates the many achievements of the past. We believe this Hand¬ book on Armillaria root disease will be of interest and value to graduate students, mycologists, pathologists, and forest managers for many years. Charles G. Shaw III USDA Forest Service Rocky Mt. Forest & Range Expt. Stn. Fort Collins, CO, USA Glen A. Kile CSIRO Division of Forestry & Forest Products, Hobart, Tasmania, Australia w Acknowledgments We are pleased to thank the various internal and exter¬ nal reviewers whose suggestions, thoughts, and sage advice have helped to improve the text and Kevin Cook for his editorial efficiency. We are deeply in¬ debted to the CSIRO Division of Forestry and Forest Products, Australia, for the support provided to the co¬ editors. We are especially indebted to Jean Richmond (Hobart, Tasmania) and Sheila Ames (Fort Collins, Colorado) for their patient contributions to typing and constant reorganization of the manuscript, as well as their diligent attention to the Fax machines in their respective locations. The last minute heroics of Mike Larsen, Nora Trowbridge, Sara Stockover, Frances Barney, and Bob Hamre are also appreciated. Dedication Robert Hartig (1839-1901) The 'Father of Forest Pathology/ who concluded that wood decay was caused by microorganisms and pro¬ vided convincing evidence for the pathogenicity of several fungi attacking trees. His monographic treat¬ ment of Agaricus melleus in Wichtige Krankheiten der Waldbaume (1874) has had an enduring influence on the perceptions of pathogenic behavior and study of Armillaria. A detailed account of Hartig's remarkable contributions to forest pathology is found in the American Phytopathological Society, English transla¬ tion of this work (Phytopathological Classics No. 12, 1975). vi Characteristics of Armillaria root disease. A: Infection of a seedling by rhizomorphs from an inoculum segment colonized by Armillaria: B: Mycelial fan in the cambial region at the base of a recently killed tree. Such fans can be diagnostic of tree death by Armillaria: C: Armillaria infection center in pole-sized ponderosa pine showing disease progression through the stand; D: Signature on an aerial photograph of an Armillaria root disease infection center. (C.G. Shaw III, R. Williams) vii Contents Preface . Hi CHAPTER 1 Nomenclature, Taxonomy, and Identification.1 Roy Watling, Glen A. Kile, and Harold H. Burdsall, ]r. Armillaria (FR.:FR.) Staude — Nomenclature and Typification.2 Generic Characteristics.3 Relationships With Other Agarics.4 Relationships Within Armillaria .4 Present and Excluded Species of Armillaria .4 Taxonomic Characters and Identification.6 Conclusions.9 CHAPTER 2 Life Cycle, Interfertility, and Biological Species.10 Jean-Jacques Guillaumin, James B. Anderson, and Kari Korhonen The Sexual System.11 Mating Reactions Among Haploids.11 Mating Between Diploids and Haploids.12 The Caryological Cycle.12 Vegetative Diploidy.12 Somatic Haploidization.13 A Possible Haploid Stage in Basidiomes.14 Nuclear Behavior in the Hymenium.15 Identification and Occurrence of Biological Species ...15 Identification.15 European Species.16 North American Species.17 Australasian Species.17 Other Regions.17 Variation Within Biological Species.18 The Identification of Genotypes.18 Non-Heterothallic Armillaria Species.19 isions.19 CHAPTER 3 Ontogeny and Physiology.21 Michael O. Garraway, Aloys Hiitterman, and Philip M. War go Structure and Morphogenesis.21 Development of Basidiomes.21 Production of Basidiomes in Culture.22 Pseudosclerotial Plates and Zone Lines.23 Rhizomorphs.24 Cytology of Rhizomorphs.25 Organization of the Differentiated Rhizomorph.27 Uptake and Transport of Nutrients and Water ..27 Concluding Comments on Rhizomorph Structure .28 Nutrition and Physiology.28 Factors Affecting Growth and Development.28 Nutritional Factors.28 Environmental Factors.33 Genetic Factors..-.34 Biochemical Changes Associated with Growth and Development.34 Cell-Wall Polysaccharides and Other Macromolecules.34 Enzymes.35 Nature of Phenoloxidizing Enzymes Produced by Armillaria .36 Conclusions. 37 Miscellaneous Themes in the Physiology of Armillaria .37 Protease.37 Antibiotics and Other Metabolites.37 Bioluminescence.38 Effect of Environment, Nutrition, and Growth Factors.38 Mechanism of Fungal Bioluminescence.38 Physiology of Host-Pathogen Interactions.39 Genetic Control.39 Metabolic Control.39 Pathogen Factors.39 Host Factors.41 Predisposition Effects.43 Stress.43 via Nutritional Changes Phenol Degradation Host-Induced Lysis. Conclusions. 44 45 46 46 CHAPTER 4 Inoculum and Infection.48 Derek B. Redfern and Gregory M. Filip Inoculum.48 Source of Inoculum.48 Substrate Quality—Conifers Versus Hardwoods ...49 Substrate Specialization.51 Longevity of Inoculum and Persistence of the Fungus.52 Factors Affecting Growth of Rhizomorphs from Inoculum.53 Variation Among Species.53 The Effect of Soil on Rhizomorph Growth.53 Moisture.54 Temperature.54 pH.55 Inhibitory Substances.55 Organic Matter and Soil Nutrient Status.55 The Distribution of Rhizomorphs in Soil.56 Inoculum Potential and Infection.56 Inoculum Potential.57 Infection.59 Conclusions.60 CHAPTER 5 Infection, Disease Development, Diagnosis, and Detection.62 Duncan J. Morrison, Ralph E. Williams, and Roy D. Whitney The Infection Process and Disease Development.62 The Infection Process.62 Host Response to Infection.63 Post-Infection Development.65 Effects on the Host.66 Physiology of Symptom Development and Host Killing.67 Disease Diagnosis.68 Above-Ground Symptoms on Individual Plants ....68 Reduction of Shoot Growth. Changes in Foliage Characteristics. Crown Dieback. Stress-Induced Reproduction. Basal Stem Indicators. Symptom Development in Relation to Extent of Colonization. Confirmation of Armillaria Occurrence. Mycelial Fans. Rhizomorphs. Basidiomes. Decay. Isolation Technique and Appearance in Culture. Biotic and Abiotic Conditions Causing Similar Symptoms. Disease Detection. Conclusions. 68 69 69 69 69 70 71 ,71 ,71 .73 .73 .73 .74 .74 .75 CHAPTER 6 Pathogenicity and Virulence.76 Steve C. Gregory, John Rishbeth, and Charles G. Shaw III Pathogenicity, Virulence, and Disease.76 Saprophytic and Parasitic Behavior.77 Conditions for Disease.77 Decay and Disease.78 Host Specialization.78 Assessing Pathogenicity and Virulence.78 Choice of Host for Inoculation Trials.78 Choice of Inoculum.81 Rhizomorphs and Measurement of Disease in Inoculation Trials.82 Field Observation.83 Indirect Methods of Assessing Virulence.83 ix Differences in Pathogenicity and Virulence.84 European and North American Species.84 Australasian Species.86 Non-Australasian Tropical and Subtropical Species.87 Conclusions.87 CHAPTER 7 Host Stress and Susceptibility.88 Philip M. War go and Thomas C. Harrington Stress Concepts and Host-Pathogen Interaction.88 Variation Among Armillaria Species, Host, and Site.88 Host Vigor and Predisposition.89 Stresses and Resistance to Armillaria .90 Barriers and Energy Reserves.90 Pathogen Nutrition.92 Stress Agents and Armillaria Root Disease.93 General.93 Abiotic Stress Factors.93 Light.93 Temperature.94 Moisture.94 Nutrients and Other Soil Factors.95 Pollutants.96 Disturbance from Partial Cutting.97 Biotic Stress Agents.98 Insect Defoliation.98 Other Insects.99 Other Diseases.99 Managing Stress.101 Conclusions.101 CHAPTER 8 Ecology and Disease in Natural Forests.102 Glen A. Kile, Geral I. McDonald, and James W. Byler Geographical Distribution of Species.102 Host Range.103 Modes of Behavior in Natural Forests.103 Decomposer.105 Mycoparasite.105 Mycotrophic (Mycorrhizal) Associations.105 Necrotrophic Plant Pathogen.106 Armillaria Species as Primary Pathogens.106 Non-Lethal Primary Disease: Root Lesions, Cankers, Butt Rot.106 Lethal Primary Disease.110 Root Rot in Boreal Forests and Western North American Coastal Coniferous Forests.110 Ring Disease of Mountain Pine.: 110 Armillaria Root Disease in Mixed Coniferous Forests of Western North America.Ill Armillaria Root Disease in Dry Sclerophyll Eucalypt Forests.114 Armillaria Species as Secondary Pathogens.114 Dispersal and Distribution.115 Basidiospores.116 Rhizomorphs and Root Contacts.117 Spatial Distributions.117 Pathogenicity, Environment, Host Resistance, and Primary Disease Expression.118 Forest Management and Disease.119 Conclusions.121 CHAPTER 9 Armillaria in Planted Hosts.122 Ian A. Hood, Derek B. Redfern, and Glen A. Kile Distribution and Importance.122 Europe and the Soviet Union.123 North America .125 Central and South America..127 Africa.128 Asia and the Pacific.129 Australasia.130 Disease Development and Impact.132 The Significance of Plantations.132 Inoculum Potential.132 Induced Host Stress.132 Choice of Species.132 Monocultures.132 Disease Dynamics.132 Disease Establishment.132 Disease Distribution Pattern.134 Secondary Disease Spread.134 Subsequent Disease Development.135 Stress and Predisposition.135 Disease Loss.136 Loss and Crop Age.136 Evaluation of Disease Impact.136 Plantation Management and Disease.137 Planting.137 Cultivation and Weed Control.138 Thinning and Pruning.138 Fertilization.139 Control of Other Pests or Diseases.139 Conclusions. CHAPTER 10 Modeling the Dynamics, Behavior, and Impact of Armillaria Root Disease.150 Charles G. Shaw III, Albert R. Stage, and Peter McNamee Carryover to Regeneration After Harvest.155 Representation of Management Actions.155 Inoculum Removal.155 Silvicultural Treatments.155 What Data are Required? .156 Conclusions.156 CHAPTER 11 Avoiding and Reducing Losses from Armillaria Root Disease.157 Susan K. Hagle and Charles G. Shaw III Needs Assessment.158 Control Options.158 Silvicultural Considerations for Natural Forests ..158 Avoiding Hazardous Sites.160 Resistance.161 Other Cultural Considerations.162 Direct Reduction of Inoculum.163 Chemical Protectants, Eradicants, and Curatives .166 Biological Control and Its Integration With Other Methods.168 Conclusions.172 Literature Cited.174 Scientific and Common Names of Plants .221 About the Authors.227 Index.231 History and Structure of the Western Root Disease Model.151 Critical Model Relationships and Associated Assumptions and Hypotheses.152 Spatial Resolution.152 Center Dynamics.152 Inside Established Centers.152 Expansion of Centers.154 xi CHAPTER 1 Nomenclature, Taxonomy, and Identification Roy Watling, Glen A. Kile, and Harold H. Burdsall, Jr. C onfusion has surrounded the nomenclature and taxonomy of the genus Armillaria (Fr.:Fr.) Staude for over a century. Until re¬ cently, taxonomists have consistently dis¬ agreed on the exact description of the genus and its correct name according to the International Code of Bo¬ tanical Nomenclature. This confusion has created un¬ certainty for taxonomists and plant pathologists, and has hindered the study of this widely distributed and economically important genus of fungi. Based on the analyses of Watling and others (1982), we consider the genus to be a natural grouping, and that Armillaria is the appropriate generic name. This conclusion has been widely accepted since that publication (Antonin 1986, Berube and Dessureault 1988, Guillaumin and others 1985, Intini 1988, Marziano and others 1987, Rishbeth 1983, Roll-Hansen 1985, Romagnesi and Marxmuller 1983, Termorshuizen and Arnolds 1987). The first record of an Armillaria species was probably either in 1729 (Micheli) or 1755 (Battarra). However, not until the later classical authors began to describe the larger fungi could several taxa now assigned to Armillaria in its restricted sense ( Armillaria sensu stricto ) be unequivocally recognized. From the pathologists' viewpoint, confusion has arisen from the assumption of many authors that Armillaria mellea (Vahl: Fr.) Kummer is a single variable or polymorphic species (Singer 1956) that occurs in both temperate and tropical regions. Although this contention is supported by maps purporting to show worldwide distribution (Dis¬ tribution of Plant Diseases 143, 3rd ed. 1969) and by host lists on an international or local basis (Laemmlen and Bega 1974, Pegler and Gibson 1972, Raabe 1962a), classical European authors such as Bolton (1788-91) re¬ alized that several taxa were involved. European interest in morphological studies of Armillaria was renewed in the 1970's (Romagnesi 1970, 1973,1978; Singer 1970a,b; Singer and Clemencon 1972). The demonstration of a bifactorial sexual incom¬ patibility system in an Armillaria species (Hintikka 1973) led to studies that showed several intersterile groups, termed "biological species", could be recog¬ nized in Europe (Korhonen 1978,1980) although, as such, "biological species" had no standing within the International Code of Botanical Nomenclature. Ander¬ son and Ullrich (1979) expanded this approach using North American isolates. Morphological and genetic data have subsequently been combined to link many biological species to morphological species and vice versa (see chapter 2). Many laboratories now consis¬ tently test interfertility to identify unknown vegetative isolates. Armillaria probably contains about 40 species, of which several may have restricted geographical distributions or vegetation associations. The movement of phanerog¬ ams or their products from one area of the world to an¬ other may, however, have extended distributions of some species. Species of Armillaria are necrotrophic pathogens of plants, and in one case of another agaric, and mycotrophic associates of achlorophyllous plants (see chapter 8). Some ecological niches recorded for mem¬ bers of the genus are undoubtedly exploited by several species, but the formerly broad concept of A. mellea ap¬ plied by many authors has confounded recognition of the species involved. Retaining voucher specimens of basidiomes 1 and vegetative isolates from phytopatho- logical studies is thus extremely important. Although the ability to identify species of Armillaria has ad¬ vanced rapidly only in recent years, we have accumu¬ lated a wealth of observational and experimental information on various aspects of Armillaria biology which makes it one of our better-known genera of Agaricales. Nomenclatural and taxonomic aspects of Armillaria in general and the European species in particular have been amply described in recent years (Antonin 1986, ‘The term basidiome is used in this publication in preference to less specific terms such as basidiocarp, carpophore, fructification, fruit body, fruiting body, sporocarp, sporophore (Maas Geesteranus 1971). Taxonomy and Identification 1 Guillaumin and others 1985, Herink 1973, Marxmuller 1987, Roll-Hansen 1985, Romagnesi and Marxmuller 1983, Termorshuizen and Arnolds 1987, Watling 1987, Watling and others 1982). This chapter provides an in¬ troductory survey of the major issues in the nomencla¬ ture and taxonomy of the genus. Armillaria (Fr.:Fr.) Staude— Nomenclature and Typification In Fries' Systema Mycologicum (1821), 12 species, includ¬ ing Agaricus melleus, were accepted in the tribe Armillaria, which he had established 2 years earlier (Fries 1819). The tribes Armillaria and Lepiota were later combined (Fries 1825) with the latter name used for the enlarged group. However, Fries (1838) reverted to Armillaria for some species. By this time, the number of species in the tribe had doubled, but its scope remained unchanged in his later Monographia Armillariarum Suecicae (Fries 1854). Staude (1857) was the first to raise Fries' tribe to ge¬ neric rank. Singer (1951b, 1955a,b, 1986) has disputed whether Staude's entry meets all the requirements for valid publication, but Staude is now generally accepted as the validating author of the genus (Donk 1949, 1962; Watling and others 1982). Singer (1951b, 1955a,b) pro¬ posed Kummer (1871) as the correct author for Armillaria, and has recently reiterated that belief (Singer 1986), a conclusion we do not accept. Thus, Singer (1986) has argued that the publication of Staude (1857) is inadmissible according to the International Code of Botanical Nomenclature, but nothing has changed since Donk (1949, 1962) clearly discussed the status of Staude's account. Watling and others (1982) found no reason to disagree with Donk's findings. Both Staude and Kummer (1871) include within their ge¬ neric concept Agaricus melleus, and as far as anyone can decide from the available information, it agrees with that outlined within Fries' (1821) tribe Armillaria. Fries (1821; p. 26) includes a reference to Battara (1755) un¬ der synonymy of tribe Armillaria but nowhere dis¬ cusses this entry further. We think that this one mention can hardly support Singer's statement "de¬ fines the basic scope of the tribus." Nothing in Fries (1821) or in Battara (1755) necessitates further explora¬ tion, and this re-emphasizes the importance of Systema Mycologicum (Fries 1821) in forming a clear base line. Clements and Shear (1931) subsequently selected it as type species for the genus in their comprehensive sur- i he nomenclature of the genera of fungi. ing Staude's authority for Armillaria, the the genus follows in a straightforward nde (1857) included four species: Ag. us, Ag , ; c Ag. aurantius, and Ag. robustus. The last two are now considered species of Tricholoma (Fr.) Staude, and Ag. mucidus is placed in Oudemansiella Spegazzini (or Mucidula Pat.). Adopting either Ag. aurantius or Ag. robustus as the type could lead to Armillaria becoming a synonym of Tricholoma. Kuhner (1988) suggested Ag. mucidus as the type, but this was never recommended by any earlier author. This choice would be unfortunate as Ag. mucidus has little in com¬ mon with Ag. melleus. The selection of Ag. melleus as type by Clements and Shear (1931), Dennis and others (1954), and Donk (1949,1962) was supported by Watling and others (1982). Agaricus melleus VahkFr. is based on leones plantarum, Flora Danica (1792), vol. 6(17): 9, plate 1013 (1790), M. Vahl (fig. 1.1) [= Armil¬ laria mellea (VahkFr.) Kummer in Der Fuhrer in die Pilzkunde (1871)]. As no herbarium specimen was avail¬ able to support this plate, neotypic material was desig¬ nated (Watling and others 1982). The generic name Armillariella (Karsten 1881) typified by Ag. melleus has been used in many publications; if Armillaria is based on a species other than Ag. melleus, Armillariella would become available. Karsten's genus is logical if Armillaria is typified by Ag. luteovirens Alb. FIGURE 1.1 — Agaricus melleus, as illustrated by Martin Vahl in Flora Danica (1790 - 1792). Marxmuller and Printz (1982) considered this figure could also represent Armillaria borealis, although Marxmuller (1987) accepted it as Agaricus melleus. 2 Taxonomy and Identification & Schw.:Fr., as supported by Singer (1951a). However, this species was not originally in Fries' tribe, a prereq¬ uisite for consideration. Armillariella is therefore an ob¬ ligate synonym of Armillaria. Floccularia Pouzar is the correct genus for Ag. luteovirens and its allies. Incorporating Armillaria into Clitocybe (Fr.) Staude has sometimes blurred the identity of what we believe to be a natural genus. While first proposed by Ricken (1915), French mycologists have most frequently fol¬ lowed this approach, for example Kuhner and Romagnesi (1953) and Heim (1950,1963). The latter in¬ cluded tropical species of Armillaria in his concept of Clitocybe. This proposal does not interfere with typifica- tion, as Armillaria would simply become a synonym of Clitocybe. However, clear differences exist in basidiome development between Armillaria and Clitocybe (Watling and others 1982). Additionally, Bennell and others (1985) showed radical differences in basidiospore wall morphology between A. tnellea and Clitocybe nebularis (Batsch:Fr.) Kummer. Clitocybe tabescens (Scop.:Fr.) Bres. is the species usually cited as a link between the two genera. It is similar to A. tnellea in basidiome devel¬ opment, basidiospore wall structure, and its bifactorial heterothallism (Anderson 1982). This species is thus best placed in Armillaria, probably as A. socialis (DC:Fr.) Herink [synonym A. tabescens (Scop.:Fr.) Emel.j. Singer (1951a) and Herink (1973) suggested subcatego¬ ries of the genus. Singer divided Armillaria (as Armillariella) into two sections distinguished by the presence or absence of a veil (annulate and exannulate species), a subdivision he later maintained (Singer 1986). Herink (1973) followed Singer and recognized Armillaria as an annulate subgenus and Desarmillaria as an exannulate subgenus. He placed Armillaria tnellea in the first and A. socialis in the second. His ideas agree with our own concepts, although we believe it will eventually be possible to subdivide the subgenus Armillaria into related subgroups. Generic Characteristics Various morphological, cultural, and other features help distinguish Armillaria from other genera of Agaricales. Collectively, these characters define the ge¬ nus, and variations among them define species. The following are the salient characteristics of Armillaria: Habit — clitocyboid with slightly sinuate, adnexed, subdecurrent or decurrent gills; bivelangiocarpic or metavelangiocarpic development in annulate spe¬ cies, apparently monovelangiocarpic development in exannulate species; solitary, gregarious, or cae- spitose. Pileus — fleshy, thinning towards margin. expallant, hygrophanous or not; color variable yel¬ low-brown, yellow-olivaceous, ochraceous, rusty- tawny, umber, cigar brown, less commonly buff or clay pink, sometimes ivory, pallid, or even mouse- gray; surface glabrous, scurfy, squamulose, squamules darker than ground color, sometimes re¬ stricted to disc; glabrescent as scales are lost; dry or becoming viscid to distinctly viscid, in some species almost glutinous. Stipe — central, fibrous-fleshy, not characteristically cartilaginous; often becoming hollow and the outer¬ most layers splitting and curling back to expose flesh; more or less annulate with floccose-membra- nous to arachnoid veil; often arising from sheets of white mycelia or from well-differentiated black rhizomorphs, and/or, associated with plaques of thin, black, tough tissue. Lamellae — close to subdistant; moderately thick; nearly white, ivory, or cream-color at first but fre¬ quently becoming spotted with cinnamon-buff, rusty-tawny, or sometimes, particularly with age, with a tinge of purple or distinctly pink; sinuate; adnexed to deeply decurrent. Flesh — of pileus pale and of stipe white at first, be¬ coming as dark as umber or Vandyke brown down¬ wards and sometimes tinted red or bluish at base where colonized by pigment-producing bacteria or nectriaceous fungi. Spore-print — white to cream-color darkening slightly on drying, and in herbarium material. Basidia — 4-spored, sometimes 2-spored; thin- walled; with or without a basal clamp-connection; hyaline; smooth-walled in aqueous alkali solutions or if thick-walled [= crassobasidia (Chandra and Watling 1983)] then appearing silvery or glassy, and/or, becoming ochraceous or fulvous. Basidiospores — ellipsoid; inamyloid; hyaline, yel¬ lowish cream-color or ochraceous in aqueous alkali solutions; weakly cyanophilic; thin to moderately thick-walled; smooth or slightly verruculose or rugulose with broad, blunt usually prominent apiculus; lacking germ-pore or apical differentiation (thinning or thickening). Cheilocystidia — present or absent, often incon¬ spicuous; variable in shape sometimes catenulate- septate; thin-walled or becoming slightly thick-walled with age sometimes with apical prolon¬ gation and with or without basal clamp-connection; smooth; hyaline to honey-colored in aqueous alkali solutions. Pleurocystidia — absent or, if present, thin-walled; poorly differentiated and rarely visible above the level of the basidia. Pileipellis — an irregular, disrupted trichodermium consisting of (i) an irregular, easily destroyed suprapellis composed of groups of fulvous or cinna¬ mon, subparallel, ascendant, loosely to strongly Taxonomy and Identification 3 adhering hyphae intermixed with broad, frequently encrusted hyphae (which form the scales), often with clamp-connections; ascendant hyphae becom¬ ing repent to form a rather amorphous adnate layer; (ii) mediopellis - of parallel to subparallel hyphae forming a cutis that may or may not gelatinize but sooner or later becomes the outermost layer; and (iii) subpellis - a compact hyphal layer. Stipitipellis — parallel hyphae overlain by more or less strongly developed, irregular, filamentous velar remnants; in parts of stipe free from velar material showing development of cylindric to elongate clav- ate or lageniform caulocystidia. Pileus and stipe trama — monomitic; hyphae inamyloid, generally lacking clamp connections. Hymenophoral trama — bilateral at first and re¬ maining so or becoming regular with age although always demonstrating some divergent arrangement; constitutive hyphae generally lacking clamp-connec¬ tions; inamyloid. Vegetative growth — variable on agar media but typically reddish-brown crustose surface mycelium; usually slow growing; with or without tufts of cin¬ namon aerial mycelium; with or without reddish- brown rhizomorphs or with white to cream-color rhizomorphs embedded in the medium with emergent reddish-brown tips; rhizomorphs branch monopodially, dichotomously, or irregularly; veg¬ etative mycelium often bioluminescent; cells uni- or multinucleate; nuclei apparently diploid. Rhizomorphs — mycelial aggregations with a mela- nized outer layer and pale, apical growing tip; pro¬ duced in culture and from infected lignicolous material. Single basidiospore isolates — from heterothallic species typically slow growing; producing white, fluffy to cottony mycelium, sometimes with areas of brown or reddish; with or without sparse rhizomorph development; nuclei haploid. Compatibility system — bifactorial; heterothallic with multiple alleles at the incompatibility loci; some species possibly homothallic. Relationships With Other Agarics Modern classifications of the Agaricales link Armillaria s.s. with the Tricholomataceae (Jiilich 1981; Kiihner 1980; Singer 1951a, 1986). However, even in the tem¬ perate northern hemisphere where the agarics have been most intensively studied, only Jiilich (1981) indi¬ cated a strong relationship between Armillaria and an¬ other genus in the Tricholomataceae, Tricholomopsis Smger. Possible relationships to the Cystodermataceae ' Romagnesi 1980), the Entolomateaceae (Bennell and 1 985), and the Amanitaceae (Heifer and Watling 1^89) also have been discussed. The many distinctive morphological characteristics of the genus, the production of characteristic rhizo¬ morphs, both parasitic and saprophytic capabilities, and the apparently diploid nuclei in the vegetative my¬ celium (see chapter 2), lead us to believe that it stands quite distantly from other agaricoid genera. Thus, Jiilich's (1981) introduction of the family Armil- lariaceae to accommodate the genus has great merit. Relationships Within Armillaria Apart from the subgeneric distinction between devel¬ opmental patterns in annulate and exannulate species and its inference of relatedness, no systematic attempt has been made to assess the phylogeny of species based on differences in morphology, physiology, biochemis¬ try, ecology, pathology, or sexual compatibility system. Computer-aided comparative studies of such attributes could assist research into species relatedness. Divergent nucleic acid composition has probable utility in ascertaining species relatedness. Anderson and oth¬ ers (1987) concluded that some particular DNA se¬ quences may be appropriately variable for phylogenetic studies. Subsequently, Anderson and oth¬ ers (1989) showed that some European Armillaria spe¬ cies and the equivalent or unidentified North American Biological Species, or NABS, (Anderson and Ullrich 1979; Berube and Dessureault 1988,1989) could be placed in distinct classes based on restriction maps of ribosomal DNA. These are: rDNA class 1, A. ostoyae (= NABS I); class 2, A. gemina (= NABS II); class 3, A. bo¬ realis; class 4, A. sinapina (= NABS V); NABS IX, X; class 5, A. calvescens (= NABS III), A. gallica (= NABS VII), A. cepistipes (= NABS XI?); class 6, A. mellea (= NABS VI). The classes are believed, with the possible exception of rDNA class 4, to represent natural groupings. In addi¬ tion, classes 1, 2, and 3 were considered to be closely related with rDNA classes 2 and 3 derived from the more widely distributed DNA class 1. Greater resolu¬ tion through detailed mapping of particular regions of the genome will assist phylogeny development. As Anderson and others (1989) have suggested, reconsid¬ ering ecological, morphological, and distributional data for taxa on the basis of restriction polymorphisms would be informative. Present and Excluded Species of Armillaria Singer (1978) prepared a key to the world taxa (as Armillariella) he considered distinct. This key needs to be updated in light of the new taxa recognized and concepts developed since that time. Table 1.1 lists 36 taxa which we believe have been documented suffi- 4 Taxonomy and Identification TABLE 1.1 — The current nomenclature and geographical occurrence of 36 Armillaria species (some as Armillariella). The citation for the original description of each species is given. Italic numbers indicate those identified as both morphological and biological species. 1. A. mellea (Vahl:Fr.) Kummer (= Korhonen D., Anderson and Ullrich NABS VI). Europe, North America, North Asia, Japan, Africa? (type species)+. 2. Armillariella affinis Singer. Central America. In Fieldiana (Bot.).21 12 (1989). 3. A. borealis Marxmuller & Korhonen (= Korhonen A.). Northern Europe, Russia. In Bull. Soc. Mycol. Fr. 98:122 (1982). 4. A. calvescens Berube & Dessureault (= Anderson and Ullrich NABS III). North America. In Mycologia. 81:220 (1989). 5. A. cepistipes Velenovsky (= Korhonen B., Anderson and Ullrich (Morrison) NABS XI?). Europe, North America?, Japan. In Ceske Houby. 1:283 (1920). 6. A. fellea (Hongo) Kile & Watling. New Guinea. In Rep. Tottori Mycol, Inst. 14:97 (1976). 7. A. fuscipes Petch (= A. heimii Pegler and A. elegans Heim). East and West Africa, Sri Lanka, Madagascar. In Ann. Roy. Bot. Gdn., Peradeniya. 4:299 (1909). t 8. A. gallica Marxmuller & Romagnesi (= A. lutea Gillet sensu Arnolds and Temorshuizen, and Watling; A. bulbosa (Barla) Kile and Watling; Korhonen E., Anderson and Ullrich NABS VII). Europe, North America, Japan. In Bull. Soc. Mycol. Fr. 103:152 (1987).# 9. A. gemina Berube & Dessureault (= Anderson and Ullrich NABS II). North America. In Mycologia. 81:217 (1989). 10. Armillariella griseomellea Singer. South America. In Beih. Nova Hedw. 29:40 (1969). 11. A. hinnulea Kile & Watling. South-eastern Australia. In Trans. Brit. Mycol. Soc. 81:131 (1983). 12. A. limonea (Stevenson) Boesewinkel. New Zealand. In Kew. Bull. 19:13 (1964). 13. A. luteobubalina Watling & Kile. Australia. In Trans. Brit. Mycol. Soc. 71:79 (1978). 14. A. mellea var. camurenensis Henning. West Africa. In Engl. Bot. Jahrb. 22:107 (1895). 15. A. melleorubens (Berkeley & Curtis) Saccardo. Caribbean. In J. Linn. Soc. 10:283 (1869). 16. A. macrospora Peck. North America. In Bull. Torrey Bot. Club. 27: 610(1900). 17. A. montagnel (Singer) Herink. South America. In Lloydia. 19:182 (1956). 18. A. nigritula Orton. Great Britain. In Notes Roy. Bot. Gdn., Edinb. 38:316 (1980). 19. A. novae-zelandiae (Stevenson) Herink. New Zealand, Eastern Australia, New Guinea, South America? In Kew Bull. 19:14(1964). 20. A. olivacea (Rick.) Herink. South America. In Lloydia. 19:180 (1956). 21. A. omnituens (Berkeley) Saccardo. India. In Hooker's J. Bot. 2:46 (1850). 22. A. ostoyae (Romagnesi) Herink (= A. polymyces (Seer.) Sing. & Clem; A. obscura Schaeff.:Fr., A. montagnei \ia r umbrinolutea Singer, = Korhonen C; Anderson and Ullrich NABS I). Europe, North America, Japan. In Bull. Soc. Mycol. Fr. 86:265 (1970). 23. A. pallidula Kile & Watling. Queensland. In Trans. Brit. Mycol. Soc. 91:307 (1988). 24. A. praecox Velenovsky. Central Europe. In Ceske Houby. 1:282 (1920). 25. A. procera Speggazim. South America. In Bol. Acad. Nac. Cienc.Cordoba. 11:385(1889). 26. A. puiggarii Speggazim. South America. In Bol. Acad. Nac. Cienc. Cordoba. 11:384 (1889). 27. A. saviezii (Singer) Herink. Byelorussia. In Nat. Syst. Sect. Crypt. Inst. Bot. Acad. Sci. URSS. 4(10-12):6 (1938). 28. A. smapina Berube & Dessureault (= Anderson and Ullrich NABS V). North America. In Can. J. Bot. 66:2030 (1988).** 29. A. solidipes Peck. North America. In Bull. Torrey Bot. Club. 27:611 (1900). 30. A. sparrei Singer (Herink). South America. In Lloydia. 19:183 (1956). 31. Armillariella tigrensis (Singer) Raith. South America. In Flora Neotropica Monogr. 3:8 (1970). 32. A. yungensis (Singer) Herink. South America. In Flora Neotropica Monogr. 3:12 (1970). Subgenus Desarmillaria 33. A. ectypa (Fr.) Lamoure. Europe. In Syst. Mycol. 1:108 (1821). 34. A. mgropunctata (Secretan) Herink. Europe. In Mycogr. Suisse. 2: 1046 (1833). 35. A. socialis (DC.:Fr.) Herink. (= A. tabescens (Scop.:Fr.) Emel.). Europe, USA? In Syst. Mycol. 1:251 (1821).* 36. Armillariella watsonii (Murrill) Singer. North America. In Proc. FL. Acad. Sci. 7:111 (1944) + For species 1,3,4,5,8,9,22, and 28, the secondary designations given are those used for the equivalent biological species by Korhonen (1978), Anderson and Ullrich (1979), and Morrison and others (1985). t Synonymy proposed by Kile and Watling (1988) on morphological criteria, although interfertility studies are required for confirmation ** A. sinapina (NABS V) may be synonymous with A. cepistipes (Anderson and others 1980, Guillaumin and others 1989a) but comparisons of basidiome morphology and further interfertility studies between European and North American material are necessary to resolve this question. # The binomial A. gallica is preferred as its identity is unequivocal, being supported by a type specimen, a culture, a full description, and a plate. * While the name A. (Clitocybe) tabescens has been frequently used for a taxon common as a pathogen in southeastern USA, it is probably a different species than that found in Europe (Guillaumin and others 1989a). Taxonomy and Identification 5 ciently to be considered species, although a few addi¬ tional taxa will probably be delineated eventually. It in¬ cludes all species known to be significant to plant pathologists and ecologists. Nomenclatural adjustment of some of Singer's Armillariella species is required. Fourteen of the species have been recognized as both morphological and biological species (see chapter 2), and future interfertility-morphological studies may re¬ sult in changes to the status of other species listed in table 1.1. Since Fries (1821), many species have been placed in Armillaria by virtue of possessing a white to cream- color spore-print and an annulus, which make it very heterogenous. With a more restricted generic concept for Armillaria, knowing where some of these taxa for¬ merly placed within Armillaria are now assigned is use¬ ful. Table 1.2 shows the concordance of Fries (1821) species with modern concepts. Fries (1838,1854,1874) included an additional 34 species in the Armillaria group, only one of which was possibly an Armillaria species s.s. (A. laricinus = A. ostoyae?). Many velate spe¬ cies of Tricholoma have been placed in Armillaria, and T. caligata (Viv.) Rick, and its allies have been traditionally placed by North Americans in the genus (Hotson 1941, Mitchel and Smith 1976, Smith 1979, Thiers and Sundberg 1976). This is erroneous and confusing be¬ cause the species are morphologically, ecologically, and biologically quite distinct from Armillaria species s.s. Romagnesi (1970, 1973), Termorshuizen and Arnolds (1987), Watling (1987), and Watling and others (1982) discussed the identity of Armillaria species illustrated in the classical literature. TABLE 1.2 — Concordance of Fries' Systema Mycologicum (1821) species in Agaricus Tribe III Armillaria with modern concepts. Species Family 1. A. robustus = Tricholoma Tricholomataceae 2. A. persoonii* 3. A. guttatus - Limacella Amanitaceae 4. A. bulbiger = Leucocortinarius Cortinariaceae 5. A. constrictus = Calocybe Tricholomataceae 6. A. subcavus - Limacella Amanitaceae 7. A. mucidus = Oudemansiella Tricholomataceae 8. A. vagans* 9. A. griseofuscus* 10. A. denigratus = Agrocybe erebia Bolbitiaceae 1 1 . A rhagodiosus = Lent in us lepideus Pleurotaceae A. melleus pagans, and A. griseofuscus cannot be equated with d are best considered nomen dubium. Recent major contributions to the description of mor¬ phological variation and the delineation of Armillaria taxa include those of Romagnesi (1970,1973,1978); Marxmuller (1982,1987); Marxmuller and Printz (1982); Romagnesi and Marxmuller (1983); Watling (1987) for Europe; Singer (1956,1969) for South America; Stevenson (1964) and Kile and Watling (1981,1983, 1988) for Australasia; and Berube and Dessureault (1988,1989) for North America. Although Chandra and Watling (1982) redescribed several Indian species, fresh collections are required to complement their herbarium studies. Mohammed and others (1989) and Mwangi and others (1989) reported cultural, genetic, and isozyme studies of African species which will help to resolve their identity. Further research is necessary for other areas such as Siberia, China, and parts of South¬ east Asia. Taxonomic Characters and Identification As with other macromycetes, species of Armillaria are delimited primarily by basidiome morphology (fig. 1.2). While vegetative isolates may be identified or grouped by various methods, basidiomes are essential for the complete description and naming of species. Basidiome macromorphology, pileipellis structure and ornamentation, ring characteristics, stipe ornamenta¬ tion, presence or absence of subhymenial or basidial clamps, location of pigments in cell walls or vacuoles, and basidiospore size and ornamentation are among characters of value for species differentiation. Separa¬ tion of some species by morphological criteria alone is difficult but no more so than in many other agaric gen¬ era. Identification may require using numerous macro- and micromorphological features combined with bio¬ chemical, cultural, and ecological information. A thor¬ ough appreciation of the most useful taxonomic characters will only be derived from careful analyses of all these features (Watling and others 1982). Analysis of European, and to a lesser extent Australasian species (Kile and Watling 1983, Shaw and others 1981), showed that it is possible to identify some species by morphological and physiological attributes of their vegetative mycelia and rhizomorphs as well as by basidiome morphology (table 1.3). Additional simple tests such as the response of the mycelium to light may also differentiate some species (Benjamin 1983; see also Hood and Sandberg 1987). Serological differences among several Armillaria species were demonstrated by Lung-Escarmant and others (1978,1985b) and Lung-Escarmant and Dunez (1979, 1980); serological techniques may, in the future, have a substantial impact on the delimitation of Armillaria 6 Taxonomy and Identification FIGURE 1.2 — Basidiomes of 12 Armillaria species from various regions of the world, demonstrating variation in the macromorphology of basidiomes. A: A, ostoyae; B: A. limonea ; C: A. novae-zelandiae ; D: A. pallidula; E: A. mellea ; F: A. fumosa ; G: A. calvescens ; H: A. luteobubalina ; I: A. gallica ; J: A. sinapina ; K: A. tabescens ; L: A. ostoyae produced in vitro. (G.A. Kile, H. Burdsall, A. Lynch, J. Worrall, P. Wargo, C.G. Shaw III, T. Harrington) Taxonomy and Identification 7 TABLE 1.3 — Morphological, physiological, and serological differences among Armillaria species common in Europe (A. mellea, A. borealis, A. cepistlpes, A. gallica, A. socialis, and A. ostoyae).* Differences between species References 1. Morphology of basidiomes in nature 2. Morphology of basidiomes in vitro 3. Morphology of the mycelium in pure culture 4. Morphology of subterranean rhizomorphs in nature 5. Morphology of subterranean rhizomorphs in a mist box 6. Response to temperature 7. Reaction to phenolic acids and terpenes 8. Polyclonal antibodies All species different Difficult distinction between A. gallica and A. cepistlpes Useful for A. ostoyae, A. borealis and A. cepistlpes All species different except A. gallica and A. cepistipes A. ostoyae, A. mellea and A. gallica different All species different except A. gallica and A. cepistipes Different temperature optima. Poor growth of A, mellea but good growth of A. socialis at 30 degrees C Specific reaction of A. gallica, others quite variable Separate A. mellea, A. gallica, A. ostoyae, A. socialis Romagnesi 1970, 1973 Marxmuller 1982, 1987 Romagnesi and Marxmuller 1983 Roll-Hansen 1985 Motta and Korhonen 1986 Watling 1987 Intim 1988 Guillaumin 1986a Korhonen 1978 Guillaumin and Berthelay 1981 Rishbeth 1986 Mohammed 1987 Intim and Gabucci 1987 Guillaumin and others 1989a Morrison 1982 Mohammed 1985, 1987 Guillaumin and others 1989a Rishbeth 1986 Mohammed 1987 Shaw 1985 Rishbeth 1986 Mohammed 1987 Guillaumin and others 1989a Lung-Escarmant and Dunez 1979, 1980 Lung-Escarmant and others 1978,1985 ‘Modified from Guillaumin species. Fox and Hahne (1989) used monoclonal anti¬ bodies, but the results to date are not as impressive as those obtained by studies using polyclonal antibodies. Refinement of the techniques by developing greater an¬ tibody specificity to overcome problems of cross reac¬ tivity between closely related species may allow accurate identification in the near future, including the 1 - 'ssibilitv of diagnostic kits for rapidly identifying field material. ‘ 1 . ; a lysis supports current species concepts J offers a powerful diagnostic tool. 1986) reported quantitative differ- ; A content between A. mellea and A. gallica. Jahnke and others (1987) and Anderson and oth¬ ers (1987,1989) showed that mitochondrial (mt) DNA was highly conserved within species but divergent be¬ tween them, and that restriction fragment patterns were therefore diagnostic for species. Smith and Anderson (1989) correctly identified 23 North Ameri¬ can isolates using DNA restriction fragment length polymorphisms. Isoenzyme and protein profiles of some northern hemi¬ sphere taxa also differ sufficiently to offer further methods of species separation (Lin and others 1989, Lung-Escarmant and others 1985b, Morrison and oth¬ ers 1984). 8 Taxonomy and Identification The biological species concept has been applied to the genus using single basidiospore isolates to delineate re- productively isolated groups as discussed in chapter 2. Using this particular approach has greatly assisted tax¬ onomists in defining species in genera with restricted interspecific but high intraspecific morphological varia¬ tion. Reproductively isolated groups have been linked to existing taxa (Marxmiiller 1982, Romagnesi and Marxmiiller 1983), led to the description of new taxa (Berube and Dessureault 1988,1989; Marxmiiller and Korhonen in Marxmiiller 1982; Marxmiiller 1987), and established intra- (Anderson and Ullrich 1979, Kile and others 1983, Korhonen 1978) and inter-continental dis¬ tributions (Anderson and others 1980, Guillaumin and others 1989a, Morrison and others 1985a). Conversely, species initially described on conventional criteria were later shown to be biological species (Guillaumin 1986a, Kile and Watling 1988). Cumulative experience suggests that reconciling mor¬ phological (taxonomic) and biological species concepts for most Armillariae will be possible. Although such studies will take time to complete, they should result in robust characterization of species. In cases for which detailed morphological examination supports a single species but interfertility studies indicate otherwise, Watling and others (1982) suggested adopting the macro-microspecies concept in which a macrospecies would consist of morphologically indistinguishable biological species. We support this suggestion. Conclusions Major studies of Armillaria taxonomy have been com¬ pleted in recent years. Linking morphological, cul¬ tural, physiological, and genetic data has often enhanced their individual values; the frequent concor¬ dance of information from a variety of sources has more clearly defined many taxa. Additional collections and application of various techniques to assess phenotypic and genotypic variation within the Armillaria flora in re¬ gions where it is incompletely known remain necessary to enhance our taxonomic understanding of the genus on a worldwide basis. Analysis of collections on which some early names are based will further assist the quest for nomenclatural stability within the genus. The genetic approach to species differentiation, initiated for Armillaria by Korhonen (1978), allowed the identifi¬ cation of species from vegetative isolates. Subsequent work has shown that vegetative isolates also may be distinguished by other cultural or physiological charac¬ teristics. The ability to identify vegetative isolates is highly useful for organisms in which the vegetative phase may often be the only one encountered. Newer techniques such as DNA analysis and production of monoclonal antibodies have the potential to further en¬ hance rapid and reliable identification of vegetative iso¬ lates. The morphological and biological species concepts ap¬ pear largely reconcilable for Armillaria, at least on the basis of our knowledge of temperate species. This per¬ haps fortuitous situation will continue to have a marked impact in clarifying the taxonomy of the genus. A stable nomenclature, well-defined species, and a vari¬ ety of identification techniques are invaluable to pa¬ thologists and ecologists in their attempts to understand the behavior and natural relationships of Armillaria spe¬ cies, clarify their natural relationships, and develop dis¬ ease-control strategies. Progress has been significant in the former areas in recent years. Taxonomy and Identification 9 CHAPTER 2 Life Cycle, Interfertility, and Biological Species Jean-Jacques Guillaumin, James B. Anderson , and Kari Korhonen S pecies are traditionally identified by their morphological characteristics. Within the last few decades, however, the "biological spe¬ cies" concept has assumed an increasingly important role in mycology. A biological species is a group of "individuals" sharing a common gene pool. In the field, there is little or no genetic exchange between biological species (Esser and Hoffman 1977). Although the biological species is a rather limited concept depen¬ dent only on the criterion of interbreeding, it has al¬ ready had a major impact on formal taxonomy. Among basidiomycetes especially, interfertility tests very often conclusively indicate species identity (Boidin 1977, Boidin and Lanquetin 1984). Of course, interfertility tests can only be conducted with sufficient knowledge of the sexual incompatibility systems and life cycles of the fungal group under investigation. In the genus Armillnria, interfertility tests became possible only when the riddle of sexuality was solved, beginning with Hintikka in 1973. In the Basidiomycetes, single basidiospores germinate to produce a mycelium usually consisting of haploid, monokaryotic (uninucleate) cells. In heterothallic spe¬ cies, haploid monokaryons anastomose with one an¬ other upon contact; if they are sexually compatible, a fertile mycelium usually consisting of dikaryotic (bi- nucleate) cells results. In many, but not all, species, the synchronous division of the paired nuclei in a dikaryon accompanies the formation of clamp connec¬ tions, the presence or absence of which is the most widely used criterion for judging whether a pairing of haploid monokaryons is sexually compatible or incom¬ patible. The dikaryon predominates in the vegetative phase of i basidiomycetes. During vegetative growth, the >mp< nent nuclei remain paired but do not fuse. -' idia does nuclear fusion (karyogamy) ^mediately before meiosis and the for- ■ ; 'diospores (fig. 2.1). Most basidiomycetes are heterothallic. The haploid monokaryon is self-sterile, and a dikaryon appears only when two haploid monokaryons carrying differ¬ ent alleles at the mating-type locus or loci contact one another and mate. "Unifactorial" species have one mat- ing-type locus, and the monospore isolates from a single basidiome segregate as two classes or "mating types" ("bipolar pattern of sexuality"). "Bifactorial" species have two mating-type loci, and the monospore isolates from a basidiome segregate as four mating- types ("tetrapolar pattern of sexuality"). A few basidiomycetes are homothallic. The haploid monokaryon is self-fertile, and becomes dikaryotic and fertile even without mating with another strain. "Pseudohomothallic" species have a uni- or bifactorial sexual incompatibility system, but individual basidios¬ pores may receive two postmeiotic nuclei carrying FIGURE 2.1 — Caryological cycles: 1) a typical hymenomycete with dikaryotic secondary stage; 2) a heterothallic Armillaria with dikaryotic subhymenium; 3) a heterothallic Armillaria with diploid subhymenium; 4) a homothallic Armillaria (A. ectypa, there are also homothallic Armillaria species with monokaryotic subhymenium). Open circles are haploid nuclei, dark circles diploid. The cycles of Armillaria are somewhat hypothetical. 10 Life Cycle compatible mating types. The resulting monospore isolates of these species are self-fertile but for a differ¬ ent reason than in true homothallic species. Some early researchers (Kniep 1911, Kiihner 1946) ob¬ served that Armillaria did not fit the general concept of the higher basidiomycete life cycle. They noted the hyphal cells of Armillaria are monokaryotic, irrespec¬ tive of whether the culture originates from a single basidiospore, basidiome tissue, or vegetative material from the field. One plausible explanation was that Armillaria is homothallic. An observation inconsistent with homothallism and inbreeding, however, was that monospore mycelia originating from a single basidiome vary considerably, suggesting meiosis and recombination in a heterozygous parent (Raabe 1953, Snider 1957). The state of knowledge of the Armillaria life cycle was aptly summarized by Raper (1966): "All criteria point to an asexual or homothallic pattern of development, save one: the variability among the monosporous progeny of single fruiting bodies." The Sexual System Mating Reactions Among Haploids Hintikka (1973) made the first and most important contribution to solving the problem of sexual reproduc¬ tion in Armillaria. He observed a macromorphological difference between monospore and tissue cultures of Armillaria. The monospore isolates usually produce a white or light-brown aerial mycelium which gives the colony a fluffy appearance. Cultures from basidiome tissues, however, are flat, crustose, and lack aerial my¬ celia. Based on this morphological distinction, Hintikka showed that Armillaria had a bifactorial sexual incom¬ patibility system. When sibling monospore isolates were confronted in culture, the colony morphology of certain pairwise combinations changed from the fluffy to the flat and crustose appearance. Also, because the cells both of unmated monospore isolates and of basidiome tissues are monokaryotic, he suspected that the nuclei in crustose mycelia were diploid. Diploidization in matings was proved later by several different lines of investigation. According to the bifactorial sexual incompatibility system, each haploid mycelium of Armillaria contains two mating-type alleles. Ax and Bx. After two haploid mycelia (belonging to the same species) contact one another and anastomose, one of four possible events may occur (fig. 2.2): (1) Incompatible mating [A ] B ] xA ] B 1 ]: The haploid partners grow side by side without intermingling. and without any substantial changes in macro- or micromorphology. (2) Compatible mating [A^xA^,]: The partners intermingle eventually to form a homogeneous colony while the morphology changes from the fluffy to the flat, crustose type. Partially disinte¬ grated septa are visible in some hyphae, indicating nuclear migration. Most species also have some dikaryotic hyphae with clamp connections. Nuclear migration and diploidization proceed rather slowly in matings of Armillaria, only about 2-3 times faster than the growth of hyphae (Korhonen 1983). (3) and (4) Hemicompatible common-A and com- mon-B matings [Afi^xA^B^ and A l B 1 xA 2 B l \\ One of these combinations is similar to an incompatible mating, but in the other combination, a broad "bar¬ rage" zone usually develops between the partners. Aerial mycelium is sparse or lacking, and sometimes the crustose mycelial type is also seen in this zone. Some ambiguity persists about the assignment of A and B factors, however. According to one interpreta¬ tion, the latter hemicompatible interaction is com- mon-A because signs of nuclear migration (disintegrated septa) can be found in some hyphae of the barrage zone, suggesting the presence of dif- ferent-B alleles (Korhonen 1978). According to the other interpretation, the crustose mycelium on the barrage zone is a common-B diploid (Guillaumin and others 1983). FIGURE 2.2 — Appearance of different incompatibility factor combinations in matings of A. ostoyae: incompatible, two hemicompatible, and compatible matings (from upper left to lower right). Age of cultures: 6 weeks. (J. Anderson) Life Cycle 11 When single-spore isolates from one basidiome are paired with each other, these four mating factor combinations appear at about equal frequencies. The great majority of pairings within a large population are compatible because the number of different alleles in the population is large, and because in any given pair¬ ing of nonsiblings collected from different localities their alleles are unlikely to be identical. No reliable estimates gauge the total number of different mating- factor alleles in the species of Armillaria. As judged on the basis of some large mating tests, the number must be several dozen at least. In this respect, Armillaria is similar to other bifactorially heterothallic basidio- mycetes. The existence of the same bifactorial sexual incompat¬ ibility system has now been shown in all temperate Armillaria species (Guillaumin 1986a, Guillaumin and others 1983, Kile 1983b, Kile and Watling 1988, Korhonen 1978, Ullrich and Anderson 1978) that have been investigated, except for the very rare Eurasian species Armillaria ectypa (Korhonen unpubl., Guillaumin unpubl.). Matings Between Diploids and Haploids A process analogous to the Buller phenomenon exists in Armillaria (Anderson and Ullrich 1982a; Korhonen 1978,1983). When a fluffy haploid mycelium is paired with a crustose diploid isolate of the same species, in many cases the morphology of the former progres¬ sively changes to crustose, indicating diploidization. The Buller phenomenon in its original sense (Raper 1966) is a mating between a monokaryon and a dikaryon: the dikaryon donates compatible haploid nuclei to the monokaryon, which is "dikaryotized." In Armillaria, the donor mycelium is diploid; the exact mechanisms of diploid-haploid mating are not known. In most cases, the diploid nuclei apparently replace the haploid nuclei in the opposing mycelium; occasionally, however, recombinant diploids appear, indicating that haploidization has taken place in the original diploids (Guillaumin 1986a). The Caryological Cycle Vegetative Diploidy In a typical basidiomycete, the final result of compat¬ ible plating is a heterokaryotic mycelium with two or h ploid nuclei in each cell. In the genus !u result is a diploid mycelium with uni- p i!though the cells in older parts of the omorphs, and in basidiomes, are commonly multinucleate. When two haploid, monokaryotic cells mate, they first unite to form a dikaryotic stage with binucleate cells and clamp connections (fig. 2.1). This stage is only tran¬ sient in Armillaria. Within a few days, the isolated dikaryotic hyphae become monokaryotic. This change is caused by somatic nuclear fusion and diploidization in the tip cells. After nuclear fusion, the cell undergoes mitotic division. This peculiar cell division produces two monokaryotic diploid cells from one dikaryotic cell (fig. 2.3). The diploid tip of the hypha continues to grow and dikaryotic cells are no longer apparent (Anderson 1982, Korhonen 1983, Korhonen and Hintikka 1974). Despite the instability of the dikaryotic hyphae, they can be cultivated by transferring dikaryotic tips repeatedly to a new medium (Korhonen and Hintikka 1974). This mating process has been observed in several spe¬ cies of Armillaria including A. borealis, A. gallica, A. cepistipes, A. ostoyae, and A. tabescens. All of these spe¬ cies have a transient, but distinct, dikaryotic stage in compatible matings (Anderson 1982, Guillaumin 1986a, Korhonen 1978). The mating process in A. mellea seems to be somewhat different. A dikaryotic stage has never been found (figs. 2.1-2.3), and the diploidization mechanism in this species is unclear (Guillaumin 1986a). Several additional lines of evidence show that the veg¬ etative stage of Armillaria is diploid. In A. ostoyae, aux¬ otrophic mutants with various nutritional deficiencies have been recovered from haploid, single-spore iso¬ lates and used as markers to investigate the mating -Y- o O _ 3 _ T ♦3* o J — --> _£Tv_-- • o =• • J -*? ~3 0—0 °»— o J • o o » 3 FIGURE 2.3 — Normal conjugate mitosis (left) and diploidization with subsequent mitosis (right) in a dikaryotic tip cell of A. cepistipes. Dark area is nucleoplasm (chromatine), open circle nucleolus (Korhonen and Hintikka 1974). 12 Life Cycle process (Ullrich and Anderson 1978). In compatible pairings of haploid strains carrying complementary auxotrophic mutations, prototrophic hyphae are recov¬ ered at a high frequency from the periphery of the mated colony. The prototrophic tips invariably consist of uninucleate cells. The observed prototrophy is due to complementarity between the haploid, auxotrophic mates within a diploid nucleus. Diploids are occasion¬ ally formed also in sexually incompatible matings, but only at low frequency and only when strong selection is applied (Anderson and Ullrich 1982a). Diploidy has also been shown by direct measurement of individual nuclear DNA contents using fluorescence photometry of material stained with the DNA binding fluorochromes mithramycin (Franklin and others 1983) and DAPI (Peabody and Peabody 1985) as well as of Feulgen-stained material (Peabody and Peabody 1984). In these kinds of studies, the fluorescence values of individual nuclei vary greatly because the vegetative hyphae are unsynchronized with respect to cell cycle, and because the technique inherently suffers consider¬ able measurement error. Therefore, the most meaning¬ ful tests compare the average fluorescence values of similar cell types of distinctly different ploidy levels. In mithramycin-stained material, purified diploids from matings have on average twice the mean nuclear DNA content of their component haploid strains (Franklin and others 1983). Nuclei with DNA content consistent with diploidy are also found in mated single-spore isolates (Peabody and Peabody 1985). The most convincing evidence for diploidy involved A. ostoyae and sexual reproduction. A single, uninucleate, putatively diploid cell was isolated from a mating of single-spore isolates. The resulting culture formed basidiomes, and all four segregant mating types were identified among the meiotic progeny (Guillaumin 1986a, Korhonen 1980). Besides Armillaria, no other hymenomycete with a dip¬ loid vegetative stage is known to occur in nature. Ex¬ ceptional diploid strains of Schizophyllum (Koltin and Raper 1968) and Coprinus (Casselton 1965) have been produced in the laboratory. Somatic Haploidization The diploid vegetative stage of Armillaria has proved to be remarkably stable. For example, of 1,224 hyphal tips isolated from 17 diploids resulting from both compat¬ ible and incompatible matings of auxotrophic strains, only two expressed segregant, auxotrophic phenotypes (Anderson and Ullrich 1982a). One segregant was from an A* B^ diploid. It retained heterozygosity at both mating-type loci and expressed one of the two auxotro¬ phic markers. The other segregant was from an A^ B= diploid, and was no longer heterozygous at the A locus and expressed both auxotrophic markers. The mecha¬ nism of low-frequency “spontaneous" segregation is not known. Another means of obtaining somatic segregants of Armillaria diploids was to use various agents known to cause somatic segregation in diploids of other, higher fungi. Of benomyl, ultraviolet light, formaldehyde, and para-fluorophenylalanine, only benomyl was effective in increased somatic segregation in Armillaria diploids (Anderson 1983). Two different kinds of selection can be used (Anderson and Yacoob 1984). When the parent diploid is crustose, colonies arising from fragments of benomyl-treated mycelium can be scanned for the fluffy morphology. Alternatively, when the parent diploid is prototrophic and heterozygous for auxotrophic alleles, colonies can be screened for auxotrophy. The first method involves less labor because it is a vi¬ sual screen. The second method involves individual testing of colonies by transfer to minimal medium. With these methods, a range of segregants can be obtained from diploids carrying various combinations of aux¬ otrophic and mating-type markers. Some segregants retain heterozygosity at mating-type loci while some do not, and a variety of auxotrophic requirements are ex¬ pressed in the segregants. Furthermore, the segregants have a variety of mean, nuclear DNA contents ranging from near haploid to near diploid levels (Anderson and others 1985). Because many of the segregants are no longer heterozygous at mating-type loci and have near- haploid DNA contents, the genetic segregation can be assumed to occur by haploidization during which one of each homologous chromosome is lost. Overall, the parasexual system is a workable alternative to sexual reproduction for genetic analysis. This is espe¬ cially so in Armillaria because some species/isolates of this genus do not fruit easily in pure culture. Benomyl- induced haploidization can also be used to obtain fluffy segregants from wild-collected diploid isolates (Ander¬ son and Yacoob 1984). Haploidization may be useful, for instance, in cases when the species identification of diploid isolates in diploid-haploid matings proves diffi¬ cult (Proffer and others 1987). Benomyl's genetic effect on Armillaria diploids raises the possibility that the benomyl in isolation media Maloy 1974) might alter the Armillaria cultures recov¬ ered. Since the concentrations of benomyl used to in¬ hibit common contaminant ascomycetes (Edgington and others 1971) are much lower than that required to destabilize diploids of Armillaria (Anderson 1983), how¬ ever, we believe that low concentrations of benomyl can be safely included in media used to isolate Armillaria. Life Cycle 13 A Possible Haploid Stage in Basidiomes A perhaps even more curious phenomenon than veg¬ etative diploidy is the reappearance of the haploid stage in the basidiomes of most Armillaria species. As had already been shown by Romagnesi (1970), the subhymenial cells and the basidia of these species are clamped. Korhonen (1980) confirmed that these clamped cells are dikaryotic, and the cytophotometric studies of Peabody and Peabody (1985) showed that these paired nuclei have DNA contents consistent with haploidy. Korhonen and Hintikka (1974) obtained pure cultures of dikaryotic hyphae from young macerated gills. The dikaryotic cultures are unstable and rapidly change into monokaryotic diploid hyphae, just as do the dikaryotic hyphae from compatible matings. This characteristic differs among the Armillaria species. Among the European species, A. borealis, A. cepistipes, A. ostoyae, A. gallica, and A. tabescens, all have clamped dikaryotic basidia, whereas the basidia of A. mellea develop from diploid cells and are clampless (Guillaumin 1986a). As stated above, the dikaryotic stage is also not found in the compatible matings of A. mellea. Concerning the non-European Armillaria species, Motta and Korhonen (1986) showed that the basidiomes of NABS VI are clampless (as are those of the correspond¬ ing European species A. mellea) while the basidiomes of NABS VII have clamped basidia, like A. gallica. Accord¬ ing to Berube and Dessureault (1988,1989), the Ameri¬ can species A. sinapina (NABS V), A. gemina (NABS II), and A. calvescens (NABS III) all possess clamped ba¬ sidia. In contrast, the five Australasian species A. luteobubalina, A. novae-zelandiae, A. hinnulea, A.fumosa, and A. pallidula have clampless basidia (Kile and Watling 1983, Podger and others 1978). As A. ostoyae produces basidiomes easily in vitro, the hymenium cytology of the basidiomes obtained in pure culture could be observed by Korhonen (1980) and Guillaumin (1986a). Korhonen noticed that the basidia of A. ostoyae in pure culture were clampless and uni¬ nucleate (like the basidia of A. mellea in nature). Guillaumin (1986a) found that while a majority of basidiomes of A. ostoyae produced in vitro had clampless basidia, some did not. Even the same isolate sometimes yielded basidiomes with either clamped or clampless basidia, suggesting that the determining factor is environmental rather than genetic. The specific conditions determining the occurrence of clamped or clampless basidia, however, have not yet been identi¬ fied. h of dikaryotic elements in the basidiomes of in -i.'lo i i as vet unclear. Tommerup and Broadbent (1975) observed that while the stipe cells are monokaryotic, dikaryotic hyphae arise from multi- nucleate cells near the developing gill folds of basidiome primordia. These authors also observed that the size of individual nuclei in the monokaryotic cells at the basidiome is about twice that in dikaryotic cells. These observations suggest that monokaryotic stipe cells, are diploid and that a nonmeiotic haploidization occurs in the basidiome trama which gives rise to hap¬ loid nuclei in the multinucleate cells and dikaryons of the gills. More recently, Peabody and Peabody (1985, 1987) reported that the monokaryotic cells of the stipe have a mean nuclear DNA content consistent with haploidy. The possible haploidization may thus occur at a stage earlier than proposed by Tommerup and Broadbent (1975). While the nonmeiotic chromosome reduction presents an intriguing possibility, no precedent exists in other, higher fungi for such a regular, nonmeiotic reduction division occurring either within the basidiome or be¬ fore basidiome initiation. Furthermore, because of the problems inherent in comparing the nuclear DNA con¬ tents of very different cell types, alternative explana¬ tions for the results of Peabody and Peabody (1985, 1987) are possible. First, one cannot assume that each individual cell contains a full DNA complement and that no DNA degradation occurred if the stipe cells are not known to be viable. Second, and perhaps less likely, the degree of DNA staining or of fluorescence quenching may depend on the specific cell type. These and other factors might produce a lower than expected average fluorescent yield for stipe cells as compared with other stages. Whether the possible nonmeiotic haploidization occurs in the trama of the basidiome or at a stage preceding the basidiome formation, it would be expected to pro¬ duce a mosaic of haploid strains including all four mat¬ ing types from any diploid strain. If the stipe consists of a mixture of haploids, then, why do cultures isolated from the stipe invariably appear as typically crustose diploids? Arguably, mating may occur among haploid components of the basidiome isolated on artificial me¬ dium, but it should be possible to recover the haploid components by maceration or micromanipulation. To our knowledge, this has not been reported. An alternative explanation for the origin of the sub¬ hymenial dikaryon is that no "extra" nonmeiotic hap¬ loidization occurs in the life cycle of Armillaria species, but that vegetative haploids may exist in the field along with diploids and may participate in the basidiome formation. Even if vegetative "germ-line" haploids do occur in the field, something must explain why cul¬ tures from vegetative material in the field usually ap¬ pear crustose and diploid. Here, too, it could be argued that mating occurs among the haploid components 14 Life Cycle when the material is isolated into pure culture. If this is the case, then it should be possible to recover the veg¬ etative haploids by maceration or micromanipulation. Nuclear Behavior in the Hymenium The behavior of basidium nuclei in Armillaria species has recently been investigated by Chahsavan-Behboudi (1974), Peabody and Motta (1979), Nguyen (1980), and Guillaumin (1986a). Two haploid nuclei enter the ba¬ sidium of those species having a dikaryotic subhymenium, and one diploid nucleus enters the basidium of those species with a monokaryotic subhymenium. From this point, the overall pattern of meiosis and basidiospore formation appears to be simi¬ lar to other hymenomycetes. The four nuclei resulting from meiosis migrate to four spores formed on the basidium. Various anomalies are frequently observed, however. Additional mitotic divisions may occur in the basidium, resulting in more than four nuclei. Only four nuclei, however, move to the top of the basidia and enter the developing basidiospores; the other nuclei degenerate. Also, the number of sterigmata can be two, three, or five instead of the usual four. A small number of basidiospores (l%-5%) are binucleate (Guillaumin 1986a). Observations of the basidia of A. gallica, A. mellea, and A. ostoyae suggest that the haploid chromo¬ some number (n) in these species is four (Guillaumin 1986a, Nguyen 1980). Identification and Occurrence of Biological Species Identification Since Korhonen (1978) and Anderson and Ullrich (1979), interfertility tests have become a common method for routine identification of species and for differentiation of unknown isolates into groups. Mat¬ ing tests are performed using haploid tester strains (monospore isolates) that represent each species to which the isolate could possibly belong. The unknown isolate is paired with all the tester strains, and the mat¬ ing reactions scored according to the appearance of the mycelium. The unmated haploid cultures are generally fluffy, and diploid cultures crustose. However, consid¬ erable variation may occur in colony morphology de¬ pending on the species, isolate, and culture conditions. Haploid cultures are sometimes rather crustose (espe¬ cially in A. gallica and A. cepistipes ); conversely, diploid cultures may be relatively fluffy, (especially in A. mellea). Furthermore, a diploid culture of some species often grows submerged in the agar medium without crustose mycelium (and aerial hyphae). In some species (A. gallica and A. cepistipes), the submerged mycelium discolors malt extract agar medium intensely brown; in others (A. ostoyae), it does not. On the other hand, the haploid isolates have a strong tendency for degenera¬ tion. Their surfaces become flat and wet, and they lose their ability for mating. Distinguishing haploid and diploid cultures by appear¬ ance alone is not always possible. However, the distinc¬ tion is usually clear-cut when the amount of aerial mycelium can be compared between pairings and unmated strains. The single best rule is that compatible matings show a reduction in the amount of aerial my¬ celium relative to the unmated strains, and incompat¬ ible matings show little or no reduction in aerial mycelium. The safest identification in mating tests is obtained when single-spore isolates from the unknown speci¬ men are used in the test (fig. 2.4). Because of the possi¬ bility that the tester and the unknown haploid culture may be conspecific but incompatible due to identical mating alleles, at least two different testers must be used for each species. The pairings are usually done on malt extract agar (l%-2%) in petri dishes. Because the diploidization process in Armillaria is rather slow, the FIGURE 2.4 — Species Identification of haploid isolates in a mating test. Each dish contains two pairings; in each pairing, the upper inoculum is a haploid tester strain, and the lower inoculum is the isolate to be identified. On vertical rows, there are two testers from A. borealis (sp. A), A. cepistipes (sp. B), and A. ostoyae (sp. C), respectively. On horizontal rows, three unknown haploid isolates have been paired with all six testers. The uppermost isolate proves to belong to A. borealis, the middle to A. cepistipes, and the lowest to A. ostoyae Age of cultures: 20 days. (J. Anderson) Life Cycle 15 FIGURE 2.5 — Species identification of diploid isolates in a mating test. The arrangement is the same as in fig. 2.4, but the unknown isolates are diploid. Tester reactions like those of A. ostoyae (lowest right) are not uncommon; the testers show only slight inhibition in growth. (J. Anderson) distance between the two inocula in each pairing should not exceed 3 mm. The results usually can be assessed after 3 weeks at room temperature, or earlier if the inocula are put closer to each other. Diploid cultures can be identified in similar tests (fig. 2.5), which are analogous to the Buller phenomenon. Apparently because of diploidy, the testers' reactions in conspecific diploid-haploid pairings are usually much slower than in haploid-haploid pairings; some¬ times the tester may fail to react at all. This sometimes makes the interpretation of diploid-haploid pairings difficult, and some patience is necessary for good re¬ sults. According to our experience, a vast majority of diploid isolates can be safely identified in diploid-hap¬ loid pairings if six specific procedures are followed: (1) This identification method should be used only in geographic areas where the species composition has first been investigated in haploid-haploid pair¬ ings. The unknown isolate must belong to one of the tester species. (2) Use at least four haploid testers from each sus¬ pected species. The testers should be relatively fresh and not degenerate in colony morphology. (3) Large tests containing material from several spe¬ cies are better than small ones. Always include un¬ paired "control" cultures of the testers and unknowns. It is also desirable to include known diploid cultures of different species in the test series for comparison. (4) Read the results first after about 3 weeks (or ear¬ lier) and again after another 3 weeks or more. Spread many dishes on the table and compare the behavior of the testers in different pairings, espe¬ cially in pairings with known diploids. Relatively small changes in the appearance and growth of the testers may be important. Do not necessarily expect a drastic change from fluffy to crustose. (5) In haploid-haploid matings as well as in haploid- diploid matings, "black lines" are formed in the agar between the cultures if they do not belong to the same species. These lines can often help consider¬ ably in diagnosis. They should not be confused, however, with the margin of the pseudosclerotia consisting of aggregated ("bladder-like") cells (Mallett and Hiratsuka 1986). (6) When the identification is unsuccessful in the first test, make a second attempt using a larger selec¬ tion of testers from the suspected species. Additional criteria may also help in identifying un¬ known diploid isolates, especially from European spe¬ cies. The morphology of mycelial mats in standardized pure cultures (i.e., on malt agar in petri dishes) suffi¬ ciently characterizes the species to assist identification (Guillaumin 1986a, Guillaumin and Berthelay 1981, Intini and Gabucci 1987, Mohammed 1987, Rishbeth 1986). The main drawback of the method is that it can¬ not distinguish A. gallica from A. cepistipes. Although the criterion of culture morphology is less helpful for identification of haploid cultures, mating tests alone are usually sufficient in this case. Guillaumin and oth¬ ers (1989a) have shown that the ability to reproduce in standard culture and the morphology and pattern of subterranean rhizomorph branching obtained in a mist box can also be used for identification (see table 1.3). European Species For the European Armillaria species, a complete synthe¬ sis between the concepts of biological and taxonomic species has been made. This means that the "biological species," which can also be regarded as taxonomic species, differ by many characteristics. Seven species of Armillaria have been found in Europe, five annulate and two exannulate. The fertility within each species and sterility between different species seem to be com¬ plete. Armillaria mellea, A. gallica, and A. ostoyae have a circumboreal distribution. Outside Europe, they have been found in North America and Japan. Interfertility seems to be almost complete between European and American populations of A. mellea (NABS VI) and A. gallica (NABS VII), respectively, but is only partial be- 16 Life Cycle tween these populations of A. ostoyae (NABS I) (Ander¬ son and others 1980, Guillaumin 1986a). Armillaria tabescens may also exist in Europe, North America, and the Far East but recent matings between the European and American forms (Guillaumin unpubl.) indicate that they are intersterile. The situation is even more complex for A. cepistipes, a European species that ap¬ pears to be partially interfertile with two different North American biological species, NABS V and NABS X (Anderson 1986, Anderson and others 1980), plus fully interfertile with NABS XI (=group F, Morrison and others 1985a). NABS XI will likely prove to be con- specific with A. cepistipes. NABS V, however, suffi¬ ciently differs from A. cepistipes to be described as a separate species ( A. sinapina, Berube and Dessureault 1988). Because a complete correspondence between the bio¬ logical species and the morphological species of Eu¬ rope has been established, many other kinds of data can complement or verify the results yielded by the mating tests (see chapter 1). Among these, the morpho¬ logical criteria generally play the most important role, although physiological, morphogenetic, and biochemi¬ cal characteristics may also be used (see chapter 1). North American Species In North America, identifying Armillaria species cur¬ rently consists of placing unknown isolates in one of nine known (annulate) biological species. All but NABS IX an NABS X are now either formally equated to Euro¬ pean species or are described as new species (Berube and Dessureault 1988,1989). Since at least three North American groups, NABS VII, VI, and I, are probably conspecific with the European species A. gallica, A. mellea, and A. ostoyae, respectively, many properties of the three European species are likely to be found in their American counterparts. Nevertheless, such an extrapolation requires caution until more information on North American material is available. For example, Mohammed and Guillaumin (unpubl.) have observed differences between the European species and their American counterparts in such characteristics as cul¬ ture morphology or the conditions needed for sexual reproduction in vitro. Moreover, within NABS I the isolates of eastern and western origin seem to differ in their ability to form basidiomes in vitro and in their level of interfertility with European A. ostoyae (Mohammed and Guillaumin unpubl.). Also, Mexican isolates of NABS I (A. ostoyae) have formed basidiomes in culture (Shaw 1989a). At present, we have no reason to believe that each Armillaria species is panmictic over its entire range. Even though each species is unique overall, genetic differences probably exist among geo¬ graphically separated populations. In addition to the species mentioned above, NABS II, III, IX, and X have been reported in North America (Anderson 1986, Morrison and others 1985a, Shaw and Loopstra 1988). Berube and Dessureault (1989) have formally described NABS II as A. gemina and NABS III as A. calvescens. NABS IX and X await further study. The original testers for all the North American biologi¬ cal species were from Anderson and Ullrich (1979; see also Anderson 1986). Several authors have used these testers to identify North American isolates by haploid- haploid pairings (Berube and Dessureault 1988,1989; Dumas 1988; Mallett and Hiratsuka 1988; Morrison and others 1985a,b; Motta and Korhonen 1986; Proffer and others 1987; Shaw and Loopstra 1988). A large number of testers from these studies are now available. Morrison and others (1985a) discovered a new biologi¬ cal species, NABS XI. As some American groups are entirely or partially compatible with some European species, Motta and Korhonen (1986) and Guillaumin and others (1989a) could also identify some American isolates through matings with European testers. Wargo (1989) and Guillaumin and others (1989a) mated dip¬ loid isolates with the haploid testers (diploid-haploid matings) with less satisfactory results. In spite of recent progress, more information is needed before the breed¬ ing relationships of all Armillaria species in the North¬ ern Elemisphere are known. Australasian Species Five Armillaria species have been found in temperate and subtropical Australasia. The situation is very simi¬ lar to that of Europe after the studies of Kile and Watling (1981,1983,1988). The identification of Australasian Armillaria species is based on the mor¬ phology of the basidiomes. Mating tests have also been extensively used by Kile, who selected a range of hap¬ loid testers for A. luteobubalina, A. hinnulea, A. novae- zelandiae, and A. fumosa. The vegetative morphology of these species is somewhat different and can be helpful for identification. Four species form basidiomes in pure culture (Guillaumin 1986a; Kile and Watling 1981, 1983), which can also aid identification either through observation of basidiome morphology or by obtaining haploid mycelia. Other Regions Morphological species have been described from Af¬ rica, India, Central and South America, and the Carib¬ bean (see table 1.1), but little is known about their status as biological species. Mohammed and others (1989) found genetic criteria of limited value in separat¬ ing African isolates. Little is known about the situation in Africa, China, and southeast Asia. Life Cycle 17 Variation Within Biological Species Because the present species concepts in Armillaria are relatively new, the variation within individual species is poorly understood. Relevant knowledge is accumu¬ lating rapidly, however. Casual observations suggest that intraspecific variation occurs in rhizomorph branching pattern, basidiome and vegetative morphol¬ ogy, pathogenicity, and physiological and biochemical characteristics. Given the variation with these param¬ eters, it is not surprising that polymorphism in isoen¬ zyme profiles (Lin and others 1989, Morrison and others 1985b) and restriction fragment patterns in nuclear (Anderson and others 1987, Anderson and others 1989, Anderson and Smith 1989) and mitochon¬ drial DNA (Jahnke and others 1987, Anderson and Smith 1989) exist in Armillaria species as they do in other species of plants, animals, and fungi that have been investigated (see also chapter 1). Perhaps the most intriguing polymorphisms occur at the mating-type loci. Although the total number of mating-type alleles has not been estimated for any Armillaria species, the numbers of alleles in small samples of strains from local environs in North America (Ullrich and Anderson 1978, Anderson and others 1979), Finland (Korhonen 1978), France (Berthelay and Guillaumin 1985), and Australia (Kile 1983b) have been determined. In all cases the number of alleles was on the order of 10. Considerably more alleles likely exist within each respective species over its entire range. The Identification of Genotypes The identification of fungal individuals (genotypes, clones) and the investigation of their spread in natural substrates may reveal valuable information about the ecology of the fungus in general and about its infection biology in particular. Three methods of genotype iden¬ tification have been used in Armillaria studies. First, the identification can be done on the basis of cultural char¬ acteristics of the isolates (Rishbeth 1978b). Second, genotypes can be identified by "somatic incompatibil¬ ity," the formation of demarcation lines in confronta¬ tions. In wood, for instance, the demarcation lines border the territories of different fungal individuals (Rayner and others 1984). Somatic incompatibility has been applied for the identification of Armillaria geno¬ types in several studies (e.g., Adams 1974; Anderson and others 1979; Hood and Morrison 1984; Hood and Sandberg 1987; Kile 1983b, 1986; Korhonen 1978; Mallet and Hiratska 1985; Shaw and Roth 1976; Siepmann 1985 Thompson 1984). The method is simple: two dip¬ loid isolates are paired in a petri dish and the confron¬ tation zone is observed after a few weeks. When the mycelia from a local site are genetically identical, they intermingle in a pairing to a single homogeneous colony. When mycelia from a site are genetically differ¬ ent, they form a permanent demarcation line between each other in a pairing. The reaction can be intensified by cultivating the fungi in wood blocks (Hood and Morrison 1984). The test based on somatic incompatibility is a very use¬ ful method for identifying fungal genotypes. Some res¬ ervations in its usefulness are necessary, however. It has been found in experiments carried out with several spe¬ cies of Basidiomycetes that this method does not always distinguish between closely related heterokaryons, espe¬ cially sibcomposed heterokaryons (products of compat¬ ible matings between single-spore mycelia originating from one genotype) or their parent heterokaryon (Adams and Roth 1967, Barrett and Uscuplic 1971). In Armillaria, the situation is comparable: sibcomposed diploids, although genetically different, produce a dis¬ tinct line of demarcation in only about half the pairings (Kile 1983b, Korhonen 1978). The occurrence of sibcomposed diploids in the neighborhood of an inten¬ sively sporulating parent mycelium is possible, at least, if not likely. Furthermore, the reactions between differ¬ ent diploid genotypes of the same species should not be confused with reactions between diploids of different species. In the latter case, the paired mycelia usually produce a black line along the demarcation zone. The black line is usually absent in pairings between two genotypes of the same species. The most serious reservation about the use of vegetative demarcation lines for distinguishing strains is that the genetic basis for these vegetative reactions in Armillaria is not known. Because the intensity of the reaction var¬ ies among genetically different diploid strains, the reac¬ tion is probably determined by many loci with allelic variation. The demarcation lines are most useful as indi¬ cators of clonal identity when they are checked against other criteria (Kile 1983b, Korhonen 1978). The third, and least ambiguous, method used in identi¬ fying Armillaria genotypes is the use of mating-type alleles as genetic markers (Berthelay and Guillaumin 1985; Kile 1983b, 1986; Korhonen 1978; Ullrich and Anderson 1978). Because many A and B alleles occur in the population, it is unlikely that two outbred diploids contain identical alleles. However, sibcomposed dip¬ loids and their parent mycelium always contain identi¬ cal alleles. Using mating-type alleles as markers is considerably more laborious than using demarcation reactions because haploid cultures, and often a large number of matings between them, are necessary. More sophisticated methods, such as investigation of 18 Life Cycle isozymes or other protein spectra, and especially of nucleic acids, will undoubtedly open new perspectives for studies on intraspecific variation. For example, a recent study by Smith and others (1990) showed that several clones of A. ostoyae (NABS I) and A. gallica (NABS VII) in a local area each had a unique mitochon¬ drial genotype that was stable during vegetative growth. The Non-Heterothallic Armillaria Species The African species A. heimii (synonym A. fuscipes, table 1.1) forms basidiomes easily in pure culture (Mohammed and others 1989). Monospore isolates of this species become crustose after 10-15 days in culture. When grown on an agar medium, they are identical to each other and also to the isolate (presumably diploid) that gave rise to the basidiome. Matings among a series of different monospore isolates from the same basidiome do not show any mating reactions. It can thus be assumed that A. heimii, at least in the conditions of artificial culture, is not heterothallic and tetrapolar as are the European, North American, and Australian species. Additional evidence for this difference in sexu¬ ality is that some monospore isolates have given rise to basidiomes that were morphologically identical to the basidiome from which the monospore originated. Monospore isolates from these first-generation basidiomes are also crustose and identical to each other, to the parent monospore, and to the original wild isolate. Again, no mating reactions can be shown among cultures of the same series. Such a sexual behavior can be explained either by ho- mothallism or by parthenogenesis. Cytological obser¬ vations support homothallism: the basidia are clampless, the dikaryons are lacking, and each young basidium receives a single, large (presumably diploid) nucleus. However, the sequence of the nuclear divi¬ sions in the basidium is similar to that of the heteroth¬ allic species, indicating that meiosis (and not a succession of "normal" mitoses) occurs in the ba¬ sidium. Some other tropical Armillaria species from Africa or the West Indies could have a similar sexual behavior, according to the preliminary results of Mohammed and others (1989). The most plausible scheme for the life cycle of these tropical Armillaria species would be that the basidiospores are haploid and the young germinants convert to diploidy early. The remaining parts of the cycle would be diploid. However, the nu¬ clei of the basidiomes of these species have not yet been studied by photometry. The non-heterothallic behavior of the tropical species could affect their dispersal. The self-fertile spores of the homothallic species do not require mates in order to complete the life cycle, and therefore may be better colonizers than those of the heterothallic species. The quite rare A. ectypa, a non-tropical Armillaria spe¬ cies which grows in arctic and alpine peat bogs of Eu¬ rope, might also have a non-heterothallic behavior. It forms basidiomes easily in pure culture at 18°C. The monospore cultures from such a basidiome are identi¬ cal to each other and, when paired, do not exhibit any mating reaction (Guillaumin unpubl.). The same is true of the monospore cultures isolated from basidiomes of natural origin (Korhonen unpubl.). Moreover, as with A. heimii, some single cultures are able to form basidiomes in vitro (Guillaumin 1973). In contrast with the tropical species, however, the basidia of A. ectypa are clamped and dikaryotic, whether the basidiomes are of natural origin (Lamoure 1965) or originate from in vitro culture (Guillaumin 1973). Thus, the life cycle of A. ectypa might be homothallic with a dikaryotic stage (the homothallic equivalent of a heterothallic species with a dikaryotic subhymenium like A. ostoyae) while A. heimii would be homothallic and lacking a dikaryotic stage (the homothallic equivalent of the heterothallic species with a monokaryotic subhymenium, A. mellea). Conclusions Genetic and cytological investigations of Armillaria have made reliable species identification possible, and demonstrated the value of the biological species con¬ cept for the genus. Moreover, recent studies have pro¬ vided new information about the caryological cycles. The mating system of Armillaria species is generally tetrapolar, but the genus also contains homothallic species, especially from the tropics. The caryological cycle is exceptional in that Armillaria is the only hymenomycete known to have a persistent and wide¬ spread diploid vegetative stage in the field. Most spe¬ cies have a dikaryotic stage in compatible matings, but it is short and unstable with diploidization occurring in hyphal tip cells. Although vegetative diploids are very stable, benomyl will induce somatic haploidy. A phe¬ nomenon analogous to the Buller phenomenon is found between diploid and haploid mycelia of Armillaria, but its underlying genetic mechanism is unclear. Despite the predominance of diploidy in the vegetative stage, the basidiomes of most species contain dikaryotic hyphae with clamp connections; the clamped basidia arise from dikaryotic cells. The origin Life Cycle 19 of this dikaryotic stage is still unclear. The basidiomes of other species do not contain dikaryotic hyphae, and the clampless basidia arise directly from uninucleate diploid cells. The adaptive consequences of caryological variation among different species remain unknown. Although much recent progress has been made in un¬ derstanding the genetic mechanisms in Armillaria, we see four areas that await investigation. First, with re¬ spect to life cycles, the nature and timing of the puta¬ tive non-meiotic haploidization (if indeed it occurs at all) and the mechanisms of homothallism need to be resolved. We believe that appropriate experiments can help to clarify these aspects of the Armillaria life cycles. Second, because species and even individual genotypes can now be accurately identified, we can expect better resolution of epidemiological patterns, from long-range dispersal through local spread and infection in forests. Third, with sexual and parasexual crosses now avail¬ able in the laboratory, it may even be possible to iden¬ tify the determinants of pathogenicity. Finally, because of the considerable background on breeding relation¬ ships, morphology, ecology, and distribution of well- delineated species, Armillaria offers an excellent opportunity to use molecular characters to reconstruct phylogenetic relationships and to assess the relative roles of geographic isolation and intersterility in fungal speciation. 20 Life Cycle CHAPTER 3 Ontogeny and Physiology Michael O. Garraway, Aloys Hiittermann, and Philip M. Wargo he Armillaria life cycle, as with other mem¬ bers of the Agaricaceae, involves many de¬ velopmental events which lead to the expression of several morphological forms. Specific structures include fruiting bodies or basidiomes, basidiospores, mycelia, pseudosclerotial tissue, and rhizomorphs. These structures enable Armillaria to accommodate various habitats and allow, directly or indirectly, various species and isolates to survive in the wild and to infect and colonize diverse hosts and substrates. This adaptability strongly influ¬ ences the pathogenicity of Armillaria (see chapter 6), and we therefore discuss these structures and their de¬ velopment. Structure and Morphogenesis Armillaria resembles other agaricaceous fungi in the ca¬ pacity of its hyphae to differentiate into various struc¬ tures. Several of these structures enable this fungus to adapt to various environmental regimes and to exploit habitats and substrates which, without the structures, would be inaccessible. The structures in consideration include: (1) basidiomes, the main generative structure (fig. 3.1); (2) mycelia (fig. 3.2); (3) melanized cells (pseudosclerotia); (4) zone lines which Armillaria forms after interacting with other fungi and with tissues of in¬ fected hosts; and (5) rhizomorphs (fig. 3.3). Structural differentiation and development in Armillaria are invariably preceded and accompanied by a series of intracellular changes which redirect meta¬ bolic pathways, redistribute organelles, and rearrange structural materials. Studies which would elucidate how differentiation and development are regulated in Armillaria would benefit microbiologists, ecologists, plant pathologists, and others who wish to control the survival, spread, and pathogenesis of this fungus. For reasons such as these, we review the nutrition and physiology of Armillaria. As a root disease fungus, Armillaria is one of the most prominent killers and decayers of deciduous and conif¬ erous trees and shrubs in natural forests, plantations, orchards, and amenity plantings throughout the world. Its roles include primary pathogen, stress-induced sec¬ ondary invader, and saprophyte. Yet, the physiological bases for the varied roles are not well understood. Acknowledging this limitation, we discuss the physiol¬ ogy of the pathogen as it relates to host-parasite interactions. The following presentations on Armillaria structures and their development, nutrition and physiology, and host-parasite interactions are intended to support the discussions of biology, ecology, and pathology in other chapters. Development of Basidiomes Descriptions of basidiome ontogeny in agaricaceous fungi, including an Armillaria, were given by Hoffman (1861). Later, Hartig (1874), Beer (1911), and Atkinson (1914) studied basidiome development in material identified as A. mellea. The latter two authors contra¬ dicted Hartig's observation on the developmental pat¬ tern. Fischer (1909a,b) studied Armillaria mucida, a species now placed in Oudemansiella (see chapter 1). FIGURE 3.1 — Basidiomes of Armillaria (probably mellea) at the base of a dead red oak tree. (P. Wargo) Ontogeny and Physiology 21 FIGURE 3.2 B — Mycelial fans of Armillaria at the base of a fumigation-damaged red pine tree. (P. Wargo) (see chapters 1 and 2). While Singh and Bal (1973) stud¬ ied basidiome ultrastructure in an Armillaria sp., fur¬ ther work contrasting a wider range of Armillaria species and using modern morphological, cytological, and biochemical methods would advance our under¬ standing of this differentiation which is essential for the completion of th e-Armillaria life cycle. Production of Basidiomes in Culture Molisch (1904) first reported the formation of Armillaria basidiomes in culture when he grew the fungus on au¬ toclaved bread. Falck (1907) grew A. mellea from basid- iospores to basidiomes and reported that light was required for basidiome development (Falck 1909). That basidiomes of several Armillaria species may be pro¬ duced in vitro has been confirmed by many subsequent studies (Bothe 1928; Falck 1930; Fox and Popoola 1990, Guillaumin and others 1984,1985,1989a; Jacques-Felix 1968; Kiangsu Research Group 1974; Kile and Watling 1981; Kniep 1911,1916; Lisi 1940; Long and Marsh 1918; Manka 1961b; Raabe 1984; Reitsma 1932; Rhoads 1925,1945; Rykowski 1974a; Shaw and others 1981; Shaw 1989a; Siepmann 1985; Tang and Raabe 1973; Terashita and Chuman 1987). These numerous reports, however, somewhat obscure the fact that basidiome production in vitro is not yet reliably achieved, al¬ though techniques are improving. This difficulty has been noted as an important limitation to some studies (Ullrich and Anderson 1978). Many substrates have been found suitable for basi¬ diome development. These include bread (Falck 1930; Kniep 1911,1916; Molisch 1904); pieces of autoclaved wood or woodchips (Guillaumin and others 1989a, Molisch 1904, Raabe 1984; Siepmann 1985, Terashita and Chuman 1987); filter paper soaked in nutrients (Reitsma 1932); oranges (Guillaumin and others 1989a, FIGURE 3.2 A — Mycelial fans of Armillaria (probably calvescens) on the root collar of a defoliated sugar maple sapling. (From Wargo and Houston 1974) FIGURE 3.3 — Rhizomorphs of Armillaria gallica on a white oak root. (P. Wargo) Reijnders (1963) and Watling (1985) classified the basidiome developmental pattern of the few Armillaria species so far studied as monovelangiocarpic (only a universal veil encloses the hymenial primordium) as in exannulate species, or bivelangiocarpic (when the hy- menium is enclosed by a partial and a universal veil) as appears to be the case in annulate species. Some of the latter species could possibly also be metavelan- giocarpic (hyphae from various tissues proliferate to grow and cover the developing hymenium), but this re¬ mains to be established. Hymenophore development is probably ruptohymenial (differentiated from the back¬ ground tissue) and the overall development pattern is stipitocarpic in which the young primordium is a stipe- group or bundle (fascicle) of hyphae lacking an -real urea of differentiated cells. ailed morphological description of early opment in Armillaria remains that of Atkim-on >r one of the North American species 22 Ontogeny and Physiology Jacques-Felix 1968, Shaw 1989a); maize kernels (Kile and Watling 1981); nutrient solutions or agars with various amendments including fruit or plant extracts (Kiangsu Research Group 1974, Manka 1961b, Reitsma 1932, Rhoads 1925, Rykowski 1974a, Shaw and others 1981, Tang and Raabe 1973, Terashita and Chuman 1987). Basidiomes have apparently not been produced on a synthetic culture medium. While a complex carbo¬ hydrate source appears necessary to sustain mycelial growth and basidiome development, the role of inor¬ ganic nutrients, vitamins, or other compounds in stimulating basidiome production is poorly under¬ stood. Rykowski (1974) found that the fungicide so¬ dium pentachlorophenolate at low concentrations stimulated basidiome development, a result confirmed by Shaw and others (1981). Incubation conditions appear to affect in vitro develop¬ ment of basidiomes. Kile and Watling (1981) and Raabe (1984) noted that basidiome development in cultures coincided approximately with the natural basidiome season although other authors have not observed such an association (Rykowski 1974, Shaw and others 1981, Tang and Raabe 1973). However, most success seems to have been achieved when cultures are incubated in the dark after inoculation and then exposed to fluctuating temperature/light regimes (Guillaumin and others 1984, 1985,1989a; Kiangsu Research Group 1974; Kile and Watling 1981; Rhoads 1925; Rykowski 1974; Terashita and Chuman 1987). While Tang and Raabe (1973) claimed light was not necessary for basidiome initiation, most authors conclude that both initiation and basidiome development require light (Rykowski 1974; Guillaumin and others 1984,1989a). In this re¬ gard, Armillaria resembles other agarics (Lu 1974, Niederpruem 1963, Niederpruem and others 1964). However, significant scope exists to better define the light and temperature conditions that control basidiome initiation and maturation. Some species of Armillaria appear to form basidiomes more readily in culture than others (Guillaumin and others 1984,1985,1989a; Rhoads 1925, 1945; Shaw and others 1981; Terashita and Chuman 1987). Apart from research by Guillaumin and others (1984, 1985, 1989a) using European Armillaria species, little comparative study has been undertaken of the basidiome develop¬ ment of different species under standard conditions, al¬ though Reaves (unpubl.) has produced basidiomes of NABS I, VII, IX, and X under standard conditions. In¬ traspecific variation in basidiome development also re¬ quires more quantitative assessment. Pseudosclerotial Plates and Zone Lines Since Hartig's first description, almost every paper on wood-destroying fungi or wood decay mentions or dis¬ cusses the dark lines which are characteristic for wood degraded by fungi (for general reviews, see Bavendamm 1939, Rayner and Todd 1979). These dark lines also form in wood infected by Armillaria. When wood is incubated under sterile con¬ ditions with a single isolate of Armillaria, zone-line for¬ mation can be obtained reproducibly within 2 months (Hansson and Seifert 1987), a process which is even considered as an economically feasible method to ob¬ tain special veneers (Hansson and Seifert 1987). The compartmentalization of decayed wood in living trees, first described by Falck (1924) and further elucidated by Shigo and his co-workers (Shigo and Tippett 1981), is a completely different phenomenon and will not be discussed here. Campbell (1934) conducted the first systematic study on zone-line formation in wood decayed by Armillaria. He showed that zone lines can also form in sterile wood blocks. Since then, the physiology of zone-line formation has been studied by several authors, some of whom worked with Armillaria. They can be produced not only in wood blocks but also in sawdust cultures (Hopp 1938) and, during intra- and interspecific pair¬ ings of different isolates, in agar culture (Mallett and Hiratsuka 1986) or wood (Hood and Morrison 1984). Three different mechanisms appear to promote pseudosclerotial plate or zone-line formation: mechani¬ cal and physical factors, antagonistic interaction of dif¬ ferent mycelia (incompatibility reactions), and genetic factors within a species. Mechanical and physical factors which have been sug¬ gested to induce pseudosclerotial plate formation in¬ clude: — fluctuating moisture content (Campbell 1934, Lopez- Real and Swift 1975, Radzievskaya and Bobko 1985a); — gas phase composition (Lopez-Real and Swift 1977); — wounding respiration-induced damage to hyphae (Lopez-Real and Swift 1977). Incompatible reactions between vegetative mycelia of different species or different isolates of the same spe¬ cies resulting in the formation of black lines have been observed frequently on decayed wood (Radzievskaya and Bobko 1985b; Rayner and Todd 1977,1979), and during pairings of different isolates in culture (see chapter 2). Leslie and Leonard (1979) analyzed the genetics of in¬ jury-induced fruiting in Schizophyllum commune Fr. and found that mechanical injury may stimulate the forma¬ tion of either pseudosclerotial plates or basidiomes. Ontogeny and Physiology 23 The first (serendipitous) genetic analysis of zone-line formation was performed by Holt and others (1983). During their genetic analysis of basidiome formation in Heterobasidion annosum (Fr.) Bref., they found that zone lines were formed only in those crosses that also formed basidiomes. Although some conflicting results remain when differ¬ ent studies are compared, the formation of pseudo- sclerotial plates is, in general, a genetically determined feature of many wood-destroying basidiomycetes which is induced by various external stimuli. The morphological changes in hyphal structure caused by zone-line induction appear to be always similar re¬ gardless of the species, the mode of induction, or the substrate on which they are formed, either inside de¬ cayed wood or in culture (Hopp 1938, Lopez-Real 1975, Mallett and Hiratsuka 1986, Rayner 1976). The process of morphogenesis of pseudosclerotia can be divided into three distinct phases (Campbell 1934, Lopez-Real 1975): proliferation of hyphae, hyphal swelling and aggregation, and pigmentation and melanization of hyphae. The pseudosclerotial plate of Armillaria is thus charac¬ terized by melanized, bladder-like cells which, espe¬ cially in sawdust cultures, form a brittle plate. In such cultures, infrequently two types of rhizomorphs were produced (Lopez-Real 1975). Ribbon-shaped rhizomorphs were formed in deeper parts of the culture whereas round, pigmented rhizomorphs occasionally were generated directly from the surface crust. This association between the black crust and the pigmented rind of the round rhizomorphs indicates a close similar¬ ity between these two, differentiated structures (Campbell 1934, Lopez-Real 1975). Rhizomorphs Rhizomorphs and mycelial cords are examples of spe¬ cial morphological adaptations. They are discrete, fila¬ mentous aggregations which are formed by some fungi growing on the forest floor or, as in the case of the my¬ celial cords of Serpula lacrymans Pers.:F.S. Gray, even on concrete (Thompson 1984). Rhizomorphs differ from mycelial cords in that they are highly differentiated, are fully autonomous, and grow apically; typical mycelial cords are aggregations of parallel, relatively undifferen¬ tiated hyphae. In addition, rhizomorphs grow out from )od base into substrates that may not support their h This feature has been described for only one fungus, S. lacrymans (Thompson 1984). certain fungi to produce rhizomorphs ceral advantages (Thompson 1984). !' tion against deleterious external agents, translocation of resources, growth from a suit¬ able food base into an environment which initially does not support growth, enhancement of inoculum poten¬ tial, and amplification of individual hyphal sensitivity to external stimuli enabling directed growth responses. Because of their frequency in some forest soils and their wide distributions, rhizomorphs had already attracted the attention of many mycologists by the middle of the nineteenth century. Moreover, because they were somewhat self-contained units they were described by taxonomists of that time as a separate fun¬ gus species: Rhizomorpha fragilis Roth. This species was further divided into two subforms, R. subterranea, which is found within soils, and R. subcorticalis, which grows beneath tree bark. An early description of the different forms of R. fragilis was published by Schmitz (1848). He is probably the earliest investigator to de¬ scribe the remarkable stability of these structures and their ability to endure prolonged desiccation after which they appear to be dead, but revive when moist¬ ened. Schmitz inferred from observing rhizomorphs in rotted timber that the fungus was probably established in the trees before felling and utilized the timber as a food base following transfer to other locations such as mine shafts. He gives an "excellent description" of Armillaria rhizomorphs (quoted from Hartig 1874) and their effect on standing trees. Like most of the leading mycologists of his time, Schmitz did not fully understand the cause-and-effect relationship between the occurrence of the fungus and the disease (Ainsworth and Sussman 1965, pp. 154-156; Huttermann 1987). De Bary (1887, pp. 28-29) gives a record of the different views on the nature of rhizo¬ morphs which were held at that time by such out¬ standing mycologists as Roth, Persson, deCandolle, Eschweiler, Acharius, Fuckel, Otth, Palisoth de Beauvais, Caspary, and Tulasne. It was Robert Hartig who resolved these differences by providing decisive proof that the rhizomorphs found in forest soils belonged to the Honey Fungus (Hallimasch), Agaricus melleus, now known as Armillaria (Hartig 1874). He carefully observed the transition between the two rhizomorphic growth forms and prepared precise illustrations of this important morphological feature of the fungus. His suggestions that different environmental conditions and differences in availability of space, in either soil or beneath the bark of living trees, influence the development and morphology of the subcortical and subterranean forms of the rhizomorphs are still valid. His early observa¬ tions that browning occurs only in rhizomorphs that have been exposed to air and not in those located un¬ der tree bark have been affirmed and explained in re¬ cent work, as has his observation that the browning 24 Ontogeny and Physiology process, through the formation of a dense rind, inhibits further lateral growth. Cytology of Rhizomorphs De Bary (1869,1887) presents a schematic drawing of mycelial aggregation and the resulting conspicuous form of a primitive Armillaria thallus (fig. 3.4). A much more detailed description of rhizomorph organization is given by Hartig (1870,1874). He clearly described the organization of the thallus (fig. 3.5) with its three layers (cortex, subcortex, and medulla); and he described and illustrated the three forms of hyphae which are charac¬ teristic of these layers. He also observed the mucila- genous nature of the rhizomorph tip and the differen¬ tial formation of the cell walls in the different layers of the rhizomorph. This work was followed by that of Brefeld (1877), who first described the apical growing region as a meristem. This view of rhizomorph mor¬ phology was not improved upon until methods of tis¬ sue preparation improved and electron microscopes were employed to study fungal structures. Motta (1969) examined thin sections of rhizomorph tips with the electron microscope and discerned the structure in more detail than Hartig or Brefeld were able to do (fig. 3.6). He confirmed Brefeld's earlier findings concerning FIGURE 3.5 — Early drawing of rhizomorph (Hartig 1874). qs FIGURE 3.6 — Diagram of rhizomorph apex, illustrating the distribution of tissues and their origins: ah, apical hyphae; ac, apical center; Im, lateral meristem; pm, primary medulla; c, cortex; gs, gelatinous sheath; sm, secondary medulla; sc, subcortex. (From Motta 1969.) Ontogeny and Physiology 25 the presence of a primary meristem in the rhizomorph apex. But he noted two types of meristematic activity: (1) the primary meristem that is located in the apical center, near the rhizomorph tip in which new hyphal elements are formed from apical initials; and (2) sec¬ ondary meristems in the lateral regions of the apex where secondary cross wall formation takes place. Differentiation of the apical initials involves synchro¬ nous nuclear divisions accompanied by segmentation in many planes. The apical initials are highly cytoplas¬ mic, possessing non-membrane-bound fibrous bundles. Otherwise, they exhibit all the features normally found in hyphae of most basidiomycetes. The cells in the re¬ gion assumed to be the primary meristem were shown by Motta to have dense cytoplasm with abundant ribo¬ somes distributed throughout. Very few vacuoles were present and they were rather small. The number of nu¬ clei per cell varied but could be quite high. The thick¬ ness of the initial wall remained constant during cell enlargement, indicating that the wall material was con¬ tinuously synthesized in these cells. Schmid and Liese (1970) and Motta (1982) subsequently confirmed these findings. Motta (1971) studied the histochemistry of the rhizomorph system. He found very high amounts of protein and nucleic acids, especially RNA, in the rhizomorph apical region. This discovery agrees with the view that this region is a true meristem. Large stores of glycogen were found in the cells adjacent to the meristem (i.e., apical center) and in the primary me¬ dulla (Motta 1971). Wolkinger and others (1975) and Granlund and others (1984) studied rhizomorphs with the scanning electron microscope and discerned basically the same morphol¬ ogy described by Motta (1969) and Schmid and Liese (1970). Granlund and others (1984) used critical point drying which enabled them to better preserve the structure of the mycelia. This technique avoided hy¬ phal collapse and allowed them to demonstrate that a loose network of hyphae (which they call the periph¬ eral cover) covers the mature parts of the rhizomorphs. The method also enabled them to measure hyphal di¬ ameters in different regions of the rhizomorph and to calculate the resistance to solution flow through these hyphae (table 3.1). Obviously, from the values given in table 3.1 the hyphae of the medulla are the most likely candidates for solute flow-mediated transport in the rhizomorphs as was speculated by other previous au- i Layner (1983) studied the ultrastructural 'age production by rhizomorphs. Using polyethylene bags to incubate rhizomorphs from logs of infected trees, they produced substantially larger rhizomorph apices with more clearly defined layers. The outer layers and the apical region were more den¬ sely packed with cells compared to those obtained by earlier studies. Their analyses of the morphology and formation of the mucilagenous layer confirmed the re¬ sults of Hartig (1874), who described both long hyphae and swollen cells with a dense interior in the mucilage. Mucilage was produced in tightly packed cells at the interface between the mucilagenous and cellular re¬ gions of the rhizomorphs. In this region, mucilage-con¬ taining vesicles coalesced with the plasma membrane, creating a mucilage-filled space between the membrane and all parts of the cell wall, with the septal plate being traversed by membrane-bound protoplasmic protruberances. After partial or complete digestion of the cell wall, this mass was released outside the cells. Powell and Rayner (1984) found a specialized layer of cells, up to several cells wide, in the apical dome. These cells were biochemically very active as judged by their numerous mitochondria; and they were characterized by axial bundles of microfilaments, several of which occurred in each individual cell. These microfibril bundles were described earlier by Motta (1969). Powell and Rayner (1984) discussed the likelihood that this specialized layer of cells may provide a short-term sup¬ ply of growth materials to the apical dome. Some disagreement exists regarding the mechanism of rhizomorph growth which can be 19 mm or more per day. Brefeld (1877) and later Motta (1969) concluded that rhizomorph extension is due to a meristematic api¬ cal center containing actively dividing cells which give rise to the various other layers. This view was chal¬ lenged by Rayner and others (1985), who suggested that rhizomorph extension might be analogous to the balanced lysis mechanism which has been proposed for TABLE 3.1 — Diameter (pm), cross-sectional area (pm) 2 , and calculated resistance to solution flow (pm 2 ) of hyphae of different rhizomorph layers. Tissue Diameter Cross-sectional area Resistance to solution flow Cortex 2.3 4.15 0.24 Sub-cortex 4.7 17.35 0.085 Medulla 13.9 151.75 0.0065 Source: Granlund and others (1984) 26 Ontogeny ami Physiology hyphal extension (Bartnicki-Garcia 1973). In this model, extension is possibly mediated by a plasticized apical dome which is driven forward by pressure generated within a tube with rigid side walls (rind) and compen¬ sated for by branching and growth of the intercalated apical hyphae. Rayner and others (1985) considered that plasticization could be facilitated by mucilage produc¬ tion which disrupts the continuity of the hyphal mesh that covers the dome. Rigidity could be achieved by melanization and compaction of the outer (rind) crust, and forward pressure could be provided via osmotically driven flow through the medullary region. However, too little evidence is available to support conclusively the hypotheses of Rayner and others (1985). Also, the basically filamentous organization of the rhizomorph apices might be obscured in thin sections of the dense cells of the apical dome (Schmid and Liese 1970). For ex¬ ample, compare the scanning electron micrographs ob¬ tained by Granlund and others (1984) with Motta's (1969) transmission electron micrographs of thin sec¬ tions. We must conclude that the mechanisms underly¬ ing growth and extension of rhizomorphs are far from being completely understood. Organization of the Differentiated Rhizomorph All authors agree on the basic structure of the differenti¬ ated rhizomorph: the outer layer consists of mucilage and a loose network of hyphae surrounding a mela- nized and densely packed cortex. The cortex is the main structure which protects the rhizomorph in soil from be¬ ing colonized by fungi and bacteria. Presumably, the melanin content of the outer cell walls confers the pro¬ tection (Bloomfield and Alexander 1967, Khuo and Alexander 1967). Below the cortex lies the subcortical layer which forms the transition to the medulla. A loose mesh of wide-diameter hyphae, the medulla, is the main structure responsible for the transport of water and nu¬ trients (Jennings 1984). Towards the center of the rhizomorph, the medullary hyphae become more and more loose, forming finally a central canal which is the main structure of oxygen translocation (Smith and Grif¬ fin 1971). At the substrate-air interface, growing rhizomorphs can form "breathing pores" (Smith and Griffin 1971) that al¬ low oxygen to diffuse through the intertwining hyphae into the central canal. These structures resemble buds of rhizomorph branches but have a completely different morphology. They are formed by tufts of hyphae, per¬ haps of aborted side branches, that have burst through the rind of the rhizomorph. The apices of these branches are composed of loosely intertwined hyphae with no or¬ ganized meristem and are directly connected with the central canal. Uptake and Transport of Nutrients and Water The earliest studies on the nature and physiology of mycelial cords proposed a definite role for them in the uptake and especially the transport of nutrients and wa¬ ter (e.g., Falck 1912). The importance of rhizomorphs for transporting oxygen to growing parts of the fungus was first elucidated by Munch (1909), whose data were con¬ firmed by Reitsma (1932). Schiitte (1956) demonstrated that when fluorescein was applied to the base of rhizomorphs, it was transported to the tips. Morrison (1975) studied the uptake of radioactively labeled chlo¬ ride and phosphate plus the uptake of ammonium ions. The two labeled ions were readily taken up by rhizomorph tips. When applied to their bases, these ions were translocated to the tips, but not in the oppo¬ site direction. The immersion of rhizomorph tips into a medium containing ammonium stimulated production of amino acids. Anderson and Ullrich (1982b) basically confirmed Morrison's observation that the transport in actively growing rhizomorphs is acropetal. Using C-14 labeled glucose and P-32 labeled phosphate as isotopic markers, they showed that diffusion was not a mecha¬ nism of transport. Only rhizomorphs living under aero¬ bic conditions were able to absorb and to transport the nutrients, suggesting that the mechanism of transport is dependent upon aerobic respiration. Rhizomorphs liv¬ ing under anaerobic conditions were able to absorb the radioactive label but not transport to it. Eamus and Jennings (1984) determined the water, sol¬ ute, and turgor potentials in Armillaria rhizomorphs and found a considerable gradient of water and turgor po¬ tential from the tip to the base of the rhizomorphs. From these data and cytological evidence, the three cri¬ teria that Zimmermann (1971) said must be fulfilled for pressure-driven flow to be accepted as a translocation mechanism in plants are fulfilled in Armillaria rhizo¬ morphs. These criteria are: (1) the conducting channel must be relatively impermeable to water in a lateral di¬ rection; (2) it must be very permeable to solutes and wa¬ ter in a longitudinal direction; and (3) turgor gradients must exist between source and sink. Eamus and others (1985) measured the internal structure and hydraulic conductivity of rhizomorphs. Their data support the view that long-distance transport occurs predominantly by solutes moving along vessel hyphae of the medulla. Granlund and others (1985) measured the velocity of translocation, estimating it to be 0.55-10.8 cm.lv'; the flux of carbon and phosphate was 0.07-3.8[nMcm 2 s' ]. They could not determine the chemical form in which carbon is translocated because of a rather vigorous lat¬ eral transfer, metabolism, and metabolic compart- mentation of the label away from the stream within the rhizomorph. By changing the source-sink relations, they were able to demonstrate basipetal transport. In addi¬ tion, bidirectional transport was observed. Ontogeny and Physiology 27 The kinetics of phosphate uptake by rhizomorphs of A. mellea was studied by Cairney and others (1988). A biphasic mode of phosphate uptake indicated two different carrier systems with different Km and Vmax values. By chemically analyzing the homogenized rhizomorphs together with nuclear magnetic resonance studies of the intact system, they could discern be¬ tween two orthophosphate pools, cytoplasmic and vacuolar, with most of the orthophosphate located in the vacuole. A significant portion of the cytoplasmic phosphorus was present in the rhizomorph as polyphosphate. Concluding Comments on Rhizomorph Structure Although development of Armillaria rhizomorphs has been studied for over 150 years, this process is still not well understood. Considerable work has been done on the structural and morphological features of rhizo¬ morph development using both light and electron mi¬ croscopes. But as will be evident later, virtually nothing is known about the biochemical mechanisms or genetic events that accompany their differentiation. The morphology of rhizomorphs reveals a unique de¬ gree of differentiation. There are more than five types TABLE 3.2 — Specialized cells and regions of the Armillaria rhizomorph and their proposed functions. 1. Gelatinous sheet and mucilage layer at the apex: — protects the apex and facilitates its growth in the soil 2. Central region of the apex: — associated with mucilage production — includes a central meristem responsible for the growth of the rhizomorph 3. Circum-medullary cells of the apex: — provide a short-range supply of growth material for the apical dome 4. Lateral meristem: — originates lateral growth behind the apex 5. Melamzed cortex: — the outer rind of the rhizomorph which protects it against fungal and bacterial attack, owing to its melanin content 6. Subcortical layer: — the secondary meristem associated with lateral growth 7. Medulla: — large cells associated with solute-mediated transport of nutrients 8 Breathing pores: r - 'ions in the rhizomorph which facilitate oxygen uptake by the organ 9. Central canal: v, tt'-m the rhizomorph which enables it to translocate gases of tissues with different ultrastructures and functions in the organ. This makes rhizomorphs the most highly differentiated vegetative tissues of fungi, reaching al¬ most the degree of differentiation of a plant root. The order and function of the different specialized cells and cell regions are summarized in table 3.2. The picture that emerges so far is that of a highly dif¬ ferentiated organ with some specialization regarding solute transport and gas diffusion. Because of these structural features, Armillaria can grow in a hostile en¬ vironment and compete with the microbiota in the for¬ est floor. In addition, this structure enhances the pathogenic potential, including the capacity to enter the intact surfaces of a tree (Woeste 1956). It may also confer some competitive advantage over other root dis¬ ease fungi, such as H. annosum (Shaw 1989b). Nutrition and Physiology In Armillaria, as with other fungi, factors that control growth and development of morphological structures may do so through the activation of key physiological and biochemical processes. Therefore, their appropriate manipulation may lead to the elucidation of underly¬ ing processes and mechanisms that determine growth and development. Since factors affecting growth and development of rhizomorphs and associated physi¬ ological and biochemical changes have been the focus of many physiological investigations of Armillaria, these topics will be emphasized. But because of the paucity of data concerning some aspects of Armillaria physiology, relevant research involving other fungi is included. Garrett (1953) was the first to systematically study the induction of Armillaria rhizomorphs in pure culture on defined media. Working with agar plates, he showed that the production of rhizomorph initials is controlled by nutritional factors. Below we discuss Armillaria nu¬ trition and physiology, including factors that affect rhizomorph development. We emphasize two themes: "factors" and biochemical changes affecting growth and development. Factors Affecting Growth and Development Nutritional Factors Carbon Sources Armillaria can utilize a wide range of carbon sources. This can be inferred from the reports of its wide host range (Raabe 1962a, 1979b; Rishbeth 1983; Singh and Carew 1983) and studies that show that some isolates 28 Ontogeny and Physiology can utilize organic substrates for maintenance and growth in soil (Garrett 1960, Morrison 1982a) and on plant hosts (Rishbeth 1972b, Wargo 1980b). This view also is confirmed by the numerous reports that Armillaria can grow in culture on various carbon sources including carbohydrates (Wargo 1981a, Weinhold and Garraway 1966), lipids (Moody and Weinhold 1972a,b), phenols (Cheo 1982; Shaw 1985; Wargo 1983b, 1984), and alcohols (Weinhold 1963, Weinhold and Garraway 1966). The capacity of this fungus to fix CO, (Schinner and Concin 1981) suggests that this, too, may be a source of carbon for growth un¬ der certain conditions. Despite the wide range of carbon sources they can uti¬ lize, Armillaria species seem to be selective in their abil¬ ity to maximally utilize them for growth. For example, when glucose, fructose, and sucrose were compared, mycelia grew but were very sparse (table 3.3). This in¬ dicates that under these conditions these carbohydrates were used primarily as sources of energy for perfor¬ mance of vital functions and only sparingly for growth. In contrast, ethanol, added as a sole carbon source or as a supplement to a medium containing glucose, fruc¬ tose, or sucrose, caused prolific growth of mycelia and rhizomorphs (table 3.3). Also, the fungus grew on etha¬ nol-supplemented media containing glucose better than on fructose, which in turn was better than sucrose. Studies with C-14 labeled sugars suggest that these dif¬ ferences were partly related to different rates of uptake and utilization (Garraway 1975). Examining the studies in which relative growth on various sugars was compared, one may conclude that Armillaria selectively utilizes carbon sources; glucose is the preferred carbohydrate. Moreover, when nutri¬ TABLE 3.3 — A comparison of ethanol, glucose, fructose and sucrose, with or without an ethanol supplement, as carbon sources for mycelial growth and rhizomorph production by Armillaria in liquid culture. Dry weight (mg.) Carbon Source (2.4 g/l) Ethanol (.24 g/l) Mycelium Rhizomorphs Total Ethanol 22 18 40 Glucose - 0.8 0.0 0.8 Glucose + 26 20 46 Fructose - 2.6 0.0 2.6 Fructose + 8 8 16 Sucrose - 0.9 0.0 0.9 Sucrose + 6 4 10 Source: Weinhold and Garraway (1966) tional conditions change, the carbon source can shift from one which primarily maintains vital functions to one that both maintains these functions and supplies carbon for synthesis of compounds needed for growth and development. As described later, such observations may help pa¬ thologists and ecologists interpret and explain certain in vivo aspects of Armillaria behavior. Presumably, when the interaction between Armillaria and a host is quiescent, there is limited access to host nutrients and growth promoters. Conversely, conditions associated with aggressive colonization of the host are likely to in¬ volve high access to host nutrients and growth promot¬ ers. Support for this view comes from studies such as those of Wargo (1972). Nitrogen Sources Besides a carbon source, Armillaria needs a suitable and adequate nitrogen source to grow and develop effec¬ tively. Garrett (1953) noted that Armillaria is not able to use nitrate as its sole nitrogen source. Also, although it grows on ammonium tartrate, the best growth was ob¬ served with amino acids. Similarly, Weinhold and Garraway (1966) studied how nitrogen sources affect growth and development of Armillaria in culture with glucose (0.5%) as a carbon source and ethanol (0.05%) as a growth stimulant. Casein hydrolyzate was the most effective nitrogen source followed by individual amino acids, several of which were more effective than inorganic nitrogen sources such as ammonium and ni¬ trate (table 3.4). TABLE 3.4 — A comparison of nitrogen sources for mycelial growth and rhizomorph production by Armillaria in liquid culture with ethanol (2.4g C/I) as carbon source. Dry weight (mg.) Nitrogen Source (0.4 g N/1) Mycelium Rhizomorphs Total Casein 12 102 114 L-Aspartic acid 25 80 105 DL-Glutamic acid 13 89 102 L-Alanine 23 75 98 L-Asparagine 20 75 95 L-Glutamine 21 61 85 Glycine 35 36 71 DL-Leucine 15 46 61 (nh 4 ) 2 hpo 4 10 47 57 kno 3 3 0 3 Control—no nitrogen 7 0 7 Source: Weinhold and Garraway (1966) Ontogeny and Physiology 29 The effectiveness of casein hydrolyzate is related to its composition of mixed amino acids, including glutamic acid and leucine, which support vigorous growth of the fungus. Also, its effectiveness may be related to amino acid uptake which, in fungi, is governed by amino acid specific transport systems (Pateman and Kinghorn 1976). Transinhibition or transport system shutdown occurs as system-specific amino acids accu¬ mulate inside hyphae (Horak and others 1977). The va¬ riety of amino acids supplied by a substrate such as casein hydrolyzate would permit more transport sys¬ tems to operate, resulting in a greater total nitrogen up¬ take. The capacity of a fungus to utilize the available nitrogen source is largely determined by the amount and type of carbon source. For example, Garrett (1953) noted that the optimal concentration of nitrogen to in¬ duce rhizomorphs increased as the carbohydrate con¬ centration in the medium increased. Rykowski (1976a) studied the interrelations between carbon and nitrogen levels in culture media on myce¬ lial growth and rhizomorph production in several iso¬ lates of Armillaria. He found that at an appropriate nitrogen level, more carbon increased the mycelial dry weight. However, at a given carbon level, an increase in nitrogen above a certain level inhibited growth. Thus, the C:N ratio which varies for different isolates was found to be decisive for rhizomorph development. Inorganic Nutrients The requirements of Armillaria for inorganic nutrients are assumed to be comparable to those reported for other fungi. On this basis, relatively large quantities of magnesium, phosphorus, potassium, sulfur, and to a lesser extent, calcium may be required whereas copper, iron, magnesium, zinc, and in some instances, molyb¬ denum may be required in minute quantities. These nutrients may play the same physiological roles in Armillaria as in other fungi (Garraway and Evans 1984). Although no systematic study has addressed the effects of various concentrations of these essential inorganic nutrients on Armillaria growth and development, Morrison (1975) recognized that the availability of inor¬ ganic ions affected its behavior in soil. Vitamins The importance of certain vitamins for growth was studied systematically by Garrett (1953), who com¬ pared the responses to thiamine and biotin. He noted that thiamine was required for growth but biotin was not. Also, Garraway (1966) noted that one isolate of *’■milla h! grew optimally in a synthetic culture me- mented with ethanol when the only vita- . l was thiamine. When this medium was deprived of thiamine, growth was reduced by 85%. In contrast, growth of this isolate was insensitive to either the presence of absence of biotin. Thus, except for thia¬ mine, Armillaria appears to have the capacity, in com¬ mon with many other decay fungi, to synthesize required vitamins from simple precursors (Garraway 1966). Thiamine, as thiamine pyrophosphate, serves as the re¬ quired coenzyme for several enzymes of intermediary metabolism that catalyze the removal or transfer of al¬ dehyde groups. These include pyruvate carboxylase, transketolase, pyruvate dehydrogenase, and alpha- ketoglutarate dehydrogenase. Fungi are more often auxoheterotrophic for thiamine than for any other vita¬ min (Garraway and Evans 1984). Organic Growth Factors Several organic compounds produce rather dramatic effects on the growth and development of Armillaria. These compounds produce a response at concentra¬ tions substantially above those produced by typical vi¬ tamins, but far below those of nutrients such as carbon and nitrogen. Compounds which promote growth and development of Armillaria in this way include alcohols, auxin and related compounds, fatty acids, and phenols and related compounds. Prior to 1963, optimal growth and development of Armillaria in defined media could be accomplished only by supplementation with undefined substrates such as yeast or figwood extract (Raabe 1962b, Weinhold and others 1962). In 1963, Weinhold discov¬ ered that low-molecular-weight alcohols and related compounds enhanced the fungus' growth and develop¬ ment (table 3.5). This made it possible to grow Armillaria on a completely defined medium and opened the way for critical studies on the nutrition and physiology of the fungus. Thus, in addition to being carbon sources, low-molecular-weight alcohols serve as organic growth factors in the sense described above. Growth was poor and rhizomorphs failed to develop on a synthetic medium containing glucose (0.5%) as the sole carbon source. But adding a small quantity (0.05%) of either ethanol, 1-propanol, or 1-butanol to the glu¬ cose medium stimulated prolific growth and rhizomorph formation (Weinhold 1963, Weinhold and Garraway 1966). Several other low-molecular-weight alcohols were shown to enhance growth and rhizomorph formation, but Armillaria isolates varied greatly in their ability to respond to different alcohols (Allermann and Sortkjaer 1973). These observations are of potential interest to those who study Armillaria ecol¬ ogy because soil microorganisms produce sufficient ethanol to promote rhizomorph development 30 Ontogeny ami Physiology TABLE 3.5 — Effect of ethanol-related compounds containing two carbon atoms, and other alcohols, in different concentrations, on rhizomorph production by Armillaria. Length (cm) at 14 days* Cone. (mmole/ liter) Ethanol Acetal¬ dehyde Potassium acetate Methanol 1-Propanol 1-Butanol 10.8 59.8+2.8 17.5+1.1 < 1.0 36.5+1.3 79.3+2.9 2.6 60.3+4.3 21.3+3.7 11.2+6.0 < 1.0 54.5+3.5 54.2+4.7 1.08 28.9+3.2 15.7+3.5 2.5+0.3 49.0+4.4 43.7+4.7 0.0 < 1.0 < 1.0 < 1.0 < 1.0 < 1.0 < 1.0 * Each value is the mean of at least six replications; standard error is indicated Source: Weinhold (1963) (Pentland 1965,1967); and ethanol may also be present in tree roots (Coutts and Armstrong 1976, Crawford and Baines 1977). With the Armillaria isolate used by Weinhold, growth on a glucose medium supplemented with ethanol was equivalent to that on a medium containing ethanol (0.5%) as the sole carbon source (Weinhold and Garraway 1966). Analysis of the glucose culture me¬ dium at various times during the incubation period, however, showed that most of the growth occurred after the ethanol supplement was depleted from the medium (Garraway and Weinhold 1968b). This indi¬ cated that glucose was effectively used as a carbon source after a period of adapting to the ethanol supplement. When extra ethanol was added to a syn¬ thetic medium after 7 days (Garraway and Weinhold 1970) or 15 days (Sortkjaer and Allermann 1972) of in¬ cubation, the growth rate rose significantly. An in¬ creased growth-rate response to ethanol accompanied a decreased short-term uptake and utilization of glu¬ cose (Garraway and Weinhold 1968a, 1970) and an in¬ creased uptake of nitrogen and phosphate (Sortkjaer and Allermann 1973). Also, Sortkjaer and Allermann found that the rate of DNA and RNA accumulation increased as ethanol was added (fig. 3.7). These obser¬ vations may provide clues to the mechanism(s) by which low-molecular-weight alcohols promote growth and development in Armillaria. Several compounds with auxin activity promote growth and development of Armillaria. For example, synthetic media supplemented with 10 mg/1 or more of indole-3-acetic acid significantly increased rhizo¬ morph production (Garraway 1970,1975). Also, 2,4- dichlorophenoxyacetic acid (2,4-D) stimulated the growth rate and amount of rhizomorphs produced by several isolates (Pronos and Patton 1979). Such observations suggest that there is value in assess¬ ing models proposed to explain how auxins act on higher plants (Key 1969, Key and others 1967, Rayle 1973) to stimulate the growth of Armillaria rhizo¬ morphs. The proposed response to auxin involving nucleic acid and protein synthesis might relate not only to the effects of auxin but to those of ethanol as well. According to this proposed mode of auxin action, the interaction of auxin with the plasma membrane re¬ leases a factor that moves through the cytoplasm and into the nucleus. The factor controls the activity of RNA polymerase in the nuclei and stimulates the syn¬ thesis of rnRNA. The new mRNA is translated in the cytoplasm, resulting in new proteins which enhance cellular growth (Key 1969). Lipids and fatty acids (Moody and Weinhold 1972a,b) and ortho- and para-aminobenzoic acid (Garraway 1970) strongly stimulate rhizomorph development when added to a defined basal medium. Since ethanol is linked metabolically to lipids and fatty acids (Garraway and Weinhold 1968a) and ortho- and para- aminobenzoic acids are linked metabolically to auxin, the possibility exists that all of these organic growth factors promote rhizomorph development by a com¬ mon mechanism. Further molecular research will help to establish whether or not a common mechanism is in¬ volved in the response of Armillaria to these various growth factors. Plant Extracts and Phenolic Substances Many studies on Armillaria have reported that unde¬ fined media such as yeast extract or potato-dextrose- agar stimulate rhizomorph formation. Raabe (1962b) reported on the suitability of wood-based culture me¬ dia for their stimulatory effect on rhizomorph induc¬ tion. Also, Weinhold and others (1962) observed that a Ontogeny and Physiology 31 DAYS DAYS FIGURE 3.7 — DNA content (A) and RNA content (B) in Armillaria as a function of time following the addition of a boost of ethanol to culture media. The dashed lines show the content of DNA or RNA after the addition of extra ethanol; partially purified extract of figwood stimulates rhizo- morph initiation and growth even though chemical analyses suggested that some factors other than etha¬ nol or related compounds might be involved. More re¬ cently, Lin and others (1985) studied the induction of rhizomorphs by substances present in bark. Their ob¬ servations that various plant constituents are able to in¬ duce rhizomorphs have been confirmed by more recent studies with auxins and phenolic compounds. During the last decade, Armillaria has been reported to variously respond to phenolic compounds. Perhaps these studies received some impetus from earlier work which concluded that ethanol may enhance rhizomorph devel¬ opment by inhibiting glucose uptake and its conversion to phenolic inhibitors (Garraway and Weinhold 1970). The "phenol inhibitor theory" received added support when Vance and Garraway (1973) found that ethanol ai¬ red the phenol composition and lowered phenol con- • n 1 Guns > n the fungus. Moreover, they noted that ex! • :ts 1 n Armillaria thalli grown on glucose media had 1 iigh l \ i Is of phenol and inhibited growth whereas ex¬ arrows indicate the time of addition of the extra ethanol. 0 = Diphenylamine method for DNA, orcinol method for RNA, A = UV method for DNA or RNA (from Sortkjaer and Allermann, 1973). tracts of thalli grown on ethanol-supplemented media had lower phenol levels and were non-inhibitory. This theory received further support when Oduro and oth¬ ers (1976) partially characterized from Armillaria a phe¬ nolic compound with antibiotic properties. Elevated levels of certain phenols may stimulate growth and rhizomorph production whereas other phenols may be inhibitory. Thus, mycelial growth was enhanced by as little as 10 mg/1 of shikimic acid (a pre¬ cursor of phenol synthesis), protocatachualdehyde, and p-hydroxy benzoic acid (Garraway 1970). Also, guaiacol (Edwards 1981, Edwards and Garraway 1981), tannic acid (Cheo 1982, Shaw 1985) and substances rich in lig¬ nin (Guillaumin and Leprince 1979) promoted growth and rhizomorph development. But gallic acid, a deriva¬ tive of oak bark tannin, inhibited certain isolates of Armillaria (Cheo 1982, Shaw 1985, Wargo 1980a). Al¬ though Armillatox, a proprietary phenolic fungicide, has been shown to inhibit rhizomorph development from wood blocks (Rahman 1978), it w T as ineffective as a control agent (see chapter 11). 32 Ontogeny and Physiology In recent years, attempts have been made to use growth on phenol-amended culture media as an aid in distinguishing species and genotypes of Armillaria. Wargo (1980a) reported the reactions of several isolates grown on a gallic acid medium both with and without ethanol. He suggested that growth differences on gallic acid-amended media may indicate differences in pathogenicity or virulence of isolates. However, this method of testing pathogenicity was found to be unre¬ liable (Shaw 1985). Efforts to distinguish Armillaria spe¬ cies according to their growth habits on culture media amended with phenolic compounds have been re¬ ported (Rishbeth 1982,1986). Shaw (1985) found differ¬ ences in the growth habits of 21 isolates of several Armillaria species depending on whether the phenol amendment used was gallic acid (the hydrolyzed form of tannic acid) or tannic acid. This could reflect differ¬ ences in the permeability of fungal cell membranes to these phenols. Such differences could confound efforts to use phenol-amended medium as an aid to distin¬ guish among species. Environmental Factors Growth and development of Armillaria involves a complex interplay of metabolic processes and other intracellular events. Therefore, environmental factors should help shape the expression of metabolic events leading to morphological changes. In the previous sec¬ tion, effects of nutritional factors on growth and devel¬ opment were emphasized. Below, we discuss effects of environmental factors such as temperature, aeration, pH, light, soil organic matter, and soil organisms. Temperature The earlier studies of Benton and Ehrlich (1941) and Bliss (1946) may have prompted the more recent sys¬ tematic studies of the effects of temperature on myce¬ lial and rhizomorph growth (Rishbeth 1968). Such studies provide information useful in predicting the fungal behavior on natural substrates and in soil. In this regard, Rishbeth (1968) noted the optimum growth rates of Armillaria mycelia and rhizomorphs on malt agar were 0.75 mm/day and 9.8 mm/day, respectively, at 28°C. The optimum growing temperature varied with the conditions but was about 22°C for rhizomorph growth from woody inocula through tubes of soil and for mycelial sheets growing along woody stems. Rhizomorphs produced by Armillaria isolates from dif¬ ferent parts of the world grew maximally at 20°C and minimally at either 10°C or above 26°C (Rishbeth 1978a). How temperature affects field behavior of Armillaria is discussed in chapter 4. Aeration The vigor of Armillaria growth in soil and on natural substrates is related to aeration and, to a lesser extent, CO, levels. For example, the dry weight of rhizomorphs was reduced when the concentration of O, was lowered or that of CO, raised (Rishbeth 1978a). These studies and those of Ono (1970), Singh (1981b), and Morrison (1976) suggest that aeration strongly affects the distribution of rhizomorphs in soils (see chapter 4). Smith and Griffin (1971) reported that oxygen affects both the rate of growth and the form of rhizomorphs. They acknowledged that maximum growth depends on high rates of oxygen diffusion within the rhizo- morph's central canals. However, a partial pressure of oxygen of 0.04 atm on their outside surfaces inhibits rhizomorphs. They believed this occurred because high partial pressures of O, stimulated the fungus to pro¬ duce p-diphenol oxidase, and that catalyzed the forma¬ tion of a brown pigment in the rhizomorphs. This pigment overlays the walls of the cells and probably prevents growth by blocking the uptake of nutrients or the disposal of waste products by the cells. pH Benton and Ehrlich (1941) investigated how pH affects various Armillaria isolates in culture. The optimum pH for growth on malt agar was 4.5 at 21 °C and 5 at 25°C. Stud¬ ies with other fungi suggest that pH influences a fungus' ability to absorb various nutrients (Garraway and Evans 1984). Accordingly, the pathogenicity and aggressiveness that Armillaria exhibits on soils with low pH (Redfern 1978, Singh 1983) may be related to the pH effect on nutri¬ ent uptake by the fungus. Light Light inhibits vegetative growth of Armillaria (Wein- hold and Hendrix 1963). Doty and Cheo (1974) found that mycelial and rhizomorph growth were inhibited by up to 80% when cultured in continuous light. Growth was reduced about 60% when cultures of the fungus were illuminated for 12 hr/day. Even exposure of only 2 hr/day inhibited growth by about 50%. The inhibitory effect of light occurred with several isolates. It was most inhibitory to isolates producing abundant rhizomorphs and less inhibitory to less productive iso¬ lates. Evidently, not all isolates or species of Armillaria are inhibited by light. For example, Benjamin (1983) showed that A. limonea produced rhizomorphs in the dark whereas A. novae-zelandiae would not produce rhizomorphs without light. This difference has been used as a diagnostic feature to separate isolates of the two species (Benjamin 1983, Hood and Sandberg 1987). Ontogeny and Physiology 33 Growth of other rhizomorphic fungi appears to be in¬ hibited by light. For example, Makambila (1978) noted that exposing cultures of Rosellinia cjuercina Hartig to light for 20 hr/day may inhibit rhizomorph growth up to 50%. Soil Organic Matter In vitro nutritional studies of Armillaria help validate the interpretation of field studies undertaken to evalu¬ ate the nutritional role of soil organic matter. Morrison (1976,1982a) indicated that rhizomorphs absorb and utilize nutrients from soil and that soils rich in organic matter supply more nutrients for rhizomorph growth. Effect of Other Organisms Pentland (1965) observed that rhizomorph develop¬ ment was stimulated in pure culture by Aureobasidium pullulans (de Bary) Arnaud and attributed this effect to ethanol produced by this fungus (Pentland 1967). Also, Watanabe (1986) tested 121 fungal isolates for their ability to stimulate rhizomorph production either by co-culturing them with Armillaria or by amending Armillaria culture media with culture broth of the tester strain. He observed that 37 of the isolates tested effec¬ tively induced rhizomorphs. The most effective genera were Macrophomina, Gliocephalis, Diploidia, and Sordaria together with two unidentified species of Deuteromycotina. His reports did not include informa¬ tion on the chemical nature of the stimulatory factors involved. Genetic Factors Most researchers now acknowledge that species of Armillaria that occur worldwide comprise a complex of populations with distinctive genetic compositions (see chapters 1 and 2). Since genetic factors determine the expression of physiological and biochemical processes, genetic variation in Armillaria could be involved with reported cultural (Raabe 1966b) and pathogenic varia¬ tions (Raabe 1967). Similarly, variation observed among Armillaria isolates in their responses to nutri¬ tional and environmental stimuli could be at least par¬ tially related to genetic differences. Examples cited previously include growth variation in response to low-molecular-weight alcohols (Allermann and Sortkjaer 1973), gallic acid (Cheo 1982, Shaw 1985, Wargo 1980a), and light (Benjamin 1983, Doty and Cheo 1974). However, nothing is known of the precise relationship between genetic control of responses to nutritional and environmental stimuli and the bio- hemical events involved. Also, the possible contribu¬ tion ot \ irus-like particles (Reaves and others 1988) to v > ition among Armillaria isolates should be consid¬ ered. Chapter 6 provides further discussion of genetics in relation to pathogenicity and virulence. Biochemical Changes Associated with Growth and Development Voluminous literature relates biochemical changes to growth and development in fungi (Burnett and Trinci 1979, Moore and others 1985, Smith and Berry 1978); but the precise ways in which these changes regulate these phenomena are not known. However, studies of how biochemical changes relate to development in fungi provide clues to the regulatory mechanisms in¬ volved. A scan of the published literature suggests that many aspects of Armillaria biochemistry are either un¬ known or poorly understood. Therefore, formulating a good working hypothesis that implicates biochemical mechanisms in the pathogen's growth and develop¬ ment is difficult. We now focus on two biochemical themes that could have relevance to the regulation of growth and development of Armillaria: cell-wall po¬ lysaccharides and other macromolecules, and phenoloxidizing enzymes. Cell-wall Polysaccharides and Other Macromolecules Because cell walls control the shape of fungal cells and thalli, their composition and structure have been given particular emphasis in developmental studies. Ethanol, at concentrations that promoted growth and rhizomorph development, increased the incorporation of glucose into cell-wall polysaccharides by over 50% (Garraway and Weinhold 1968a, 1970). This could mean that cell-wall polysaccharide biosynthesis plays a part in the growth response (i.e., basidiome or rhizomorph formation) to various stimuli, as indicated in studies with other fungi (Stewart and Rogers 1978, Sietsma and Wessels 1977, Wang and others 1968, Wessels 1966). For example, the ratio of R-glucans (al¬ kali-insoluble, highly branched beta-1,3- and beta-1,6 glucan) to S-glucans (alkali-soluble, alpha-1,3-glucan) was reported to change during basidiome development of Schizophyllum commune Fr. (Wessels 1965). Also, changes in cell-wall polysaccharide composition were correlated with geneticallv controlled changes in mor¬ phology in this fungus (Wang and others 1968). More¬ over, cell-wall polysaccharide fractions from an S. commune mutant that failed to develop fully formed basidiomes were resistant to enzyme solubilization, whereas the same fractions from the wild-tvpe isolate were soluble (Wessels 1966). Similar studies applied to Armillaria might help elucidate the role of cell-wall bio¬ synthesis in its growth and development. A complex carbohydrate was recovered from mycelial cultures of Armillaria and some of its components have been char- 34 Ontogeny and Physiology acterized (Bouveng and others 1967). But the impor¬ tance for morphogenesis, if any, is not known. Changes in large molecules not associated with the cell walls also occur during growth and development. For example, DNA and RNA contents of Armillaria in¬ creased at three times the rate of the dry weight in the first few days after ethanol was added to thalli (Sortkjaer and Allermann 1973). Also, similar increases in protein were observed in response to ethanol (Garraway unpublished). Thus, ethanol at concentra¬ tions which promote growth and development of Armillaria caused an early increase in constituents needed for nuclear division as well as for protein syn¬ thesis. An association between lipids and growth and rhizomorph production in Armillaria was suggested from studies with C-14 labeled ethanol (Garraway and Weinhold 1968a). Armillaria preferentially incorporated ethanol into lipids. Furthermore, lipids of the type which are assumed to be present in Armillaria and its natural substrates, including lecithin, oleic acid, and li- noleic acid, were able to replace ethanol as promoters of rhizomorph production (Moody and Weinhold 1972a,b). Enzymes Diverse enzyme studies have attempted to establish clues to the biochemical factors which regulate growth and development in fungi. Although changes in vari¬ ous enzymes have been reportedly correlated with morphogenesis, they are probably secondary to the more fundamental changes involved. This view is sup¬ ported by studies involving enzyme levels and isoen¬ zymes in S. commune (Bromberg and Schwalb 1978, Ullrich 1977). Work with Armillaria dehydrogenases are relevant in this regard. Mallett and Colotelo (1984) ana¬ lyzed the activity and isoenzyme pattern of alcohol de¬ hydrogenase during ethanol-induced rhizomorph formation. They found a significant increase both of the enzyme activity and the number of isoenzymes of alco¬ hol dehydrogenases in the rhizomorphs but not in the mycelium. The relevance of the biochemical event studied appears obvious: alcohol dehydrogenase is needed for the metabolism of ethanol. But the rel¬ evance of this biochemical event to rhizomorph mor¬ phogenesis is still an open question. Currently, some researchers are evaluating how the ob¬ served correlation between phenoloxidizing enzymes and rhizomorph development affects morphogenesis. The association of phenoloxidizing enzymes with rhizomorph growth received increased attention with the report that CF partial pressures above 0.04 atm at the rhizomorph surface enhanced accumulation of a brown pigment and inhibited its growth (Smith and Griffin 1971). Since high CF partial pressures stimulated the activity of p-diphenol oxidase, they proposed that the pigment formed as a result of enzymatic polymer¬ ization of phenols. Electron micrographs revealed that the pigment became localized in the intracellular spaces of the rhizomorphs. Smith and Griffin (1971). proposed that the pigment inhibited rhizomorph growth because an impermeable layer of polymerized phenol formed and it probably prevented the uptake of nutrients or the disposal of waste products by the cells. More recently, Worrall and others (1986) have pro¬ posed a stimulatory role for laccase in rhizomorph ini¬ tiation and development. Evidence supporting their claim includes several correlations. Ethanol and other substances that induced rhizomorphs in Armillaria also induced laccase (phenol oxidase) formation. In a range of isolates, rhizomorph production and laccase activi¬ ties were positively correlated. Laccase was first de¬ tected just before the appearance of rhizomorph initials. Laccase activity peaked when rhizomorph growth was highest and fell to near zero when rhizo¬ morph growth ceased. Laccase was not detected in cul¬ tures which were not induced to form rhizomorphs. Also, laccase activity and rhizomorph production, but not mycelial growth, were decreased by enzyme inhibi¬ tors with activity against laccase. The contrasting interpretations of the role of phenoloxidizing enzymes by Smith and Griffin (1971) on the one hand, and by Worrall and others (1986) on the other, could involve different species of Armillaria. But contrasts are commonly encountered in Armillaria research. Edwards (1981) and Garraway and Edwards (1983) found that on a synthetic medium with casein hydrolyzate as the nitrogen source, a supplement of guaiacol (200 mg/1) promoted rhizomorph formation and increased phenoloxidizing enzyme (presumably laccase) activity. In contrast, when casein hydrolyzate was replaced with L-asparagine as the nitrogen source the same guaiacol supplement increased phenol¬ oxidizing enzyme activity but not rhizomorph devel¬ opment. Adding an ethanol supplement to a medium containing guaiacol increased the activity of a laccase- like phenoloxidizing enzyme as well as rhizomorph growth. Thus, phenoloxidizing enzyme activity in Armillaria is apparently correlated with, but is not caus- atively related to, rhizomorph production in response to ethanol and other substances. Marsh and Wargo (1989) observed a similar association of laccase activity and rhizomorph formation among isolates of five spe¬ cies of Armillaria. Among the isolates that produced rhizomorphs, there was an association of higher laccase activity with greater rhizomorph production. Some iso- Ontogeny and Physiology 35 lates, however, had laccase activity but produced no rhizomorphs (Marsh and Wargo 1989). Phenoloxidizing enzymes have been implicated in the regulation of morphogenesis and differentiation of sporulating and resting structures in basidiomycetes and other fungi including S. commune (Leonard 1971, 1972, Phillips and Leonard 1976, Wessels and others 1985), Coprinus congregatus (Bull, ex St. Amans) Fr. (Choi and others 1987, Ross 1982), Lentinus edodes (Leatham and Stahmann 1981), Podospora anserina (Ces.) Niessl (Esser 1968, Molitoris and Esser 1971), Sclerotium rolfsii Sacc. (Chet and others 1972, Miller and Liberta 1977), and Sclerotinia sclerotiorum (Lib.) deBary (Wong and Willetts 1974). Very likely, they are important in these processes in Armillaria as well. Nature of Phenoloxidizing Enzymes Produced by Armillaria Because of the proposed causative association between rhizomorph morphogenesis and phenoloxidizing en¬ zymes, the nature of these enzymes and their produc¬ tion by Armillaria needs to be reviewed. We do so giving consideration to the terminology for describing phenoloxidizing enzymes and the substrates used in their assay (Mayer 1987, Mayer and Harel 1979). The commission on enzymes refers to monophenol monoxygenase (tyrosinase) as 1.14.18.1, diphenol oxi¬ dase (catechol oxidase, diphenol oxygen oxidoreduc- tase) as 1.10.3.2, and laccase as 1.10.3.1 (Mayer 1987). This new classification differentiates between two reac¬ tions of the same enzyme, 1.14.18.1 for the cresolase ac¬ tivity and 1.10.3.2 for the catecholase activity of the same enzyme, catechol oxidase (Mayer 1987). Mayer proposes the general terms of "catechol oxidase" and "laccase" as the least confusing terms to use. Catechol oxidase can oxidize monophenols (tyrosinase or cresolase activity) or o-diphenols (catecholase activity); it cannot oxidize p-diphenols and this is diagnostic (Mayer and Harel 1979). Laccase can oxidize a wide range of substrates including mono-, di-, and tri¬ phenols. It can oxidize both o- and p-diphenols and its ability to oxidize p-diphenols is diagnostic (Mayer and Harel 1979). Catechol oxidase (tyrosinase) in fungi is primarily an intracellular enzyme and may have a role in melanin formation. Laccase is commonly excreted by fungi and has roles in lignin oxidation and degradation and detoxificiation of antifungal phenols in plant tis¬ sues (Mayer and Harel 1979). Peroxidase (1.11.1.7) also catalyzes the oxidation of 'aols by hydrogen peroxide (H,OJ and is non-spe- pi'i iols. Much of the polyphenol oxidase ac- i in the Armillaria literature could idase activity if H,O z commonly present in cell-free preparations was not removed. For ex¬ ample, Mallett and Colotelo (1984), using 4-amino- antipyrine, a substrate specific for peroxidase, detected peroxidase in exudates from Armillaria rhizomorphs. Also, they used catechol to detect phenol oxidase activ¬ ity in the exudates. Since catechol is oxidized by tyrosi¬ nase, laccase, and peroxidase, a proportion of the phenol oxidase activity detected included peroxidase. These workers also noted the presence of beta-glucosi- dase, acid protease, and alkaline protease in the exu¬ dates. Peroxidase activities were also reported in rhizomorph extracts of Armillaria by Lanphere (1934) and Lyr (1955). However, no substrate specific for peroxidase activity was used nor was catalase added to extracts to destroy H n O n and eliminate peroxidase activity. Both tyrosinase and laccase activities have been re¬ ported in mycelial extracts of Armillaria (Kaarik 1965); but laccase can oxidize both tyrosine and guaiacol (p- and o-diphenols), the two substrates used. Both tyrosi¬ nase (catechol oxidase) and laccase activities were based on visual color development in tubes with agar and either guaiacol or tyrosine as substrates in the growth medium. Stronger evidence for laccase (p-diphenol) activity was reported in rhizomorphs of A. mellea (Jacques-Felix 1968) and A. elegans (Smith and Griffin 1971), the latter now known to be A. luteobubalina. Worrall and others (1986), working with several Armillaria species, de¬ tected true laccase activity in culture liquid using 2,6- dimethoxyphenol and p-phenylenediamine as substrates. They found a general relationship of laccase production and species of Armillaria related to the pro¬ clivity of each species to produce rhizomorphs. Armillaria mellea isolates tended to have relatively high laccase activity and rhizomorph production, A. ostoyae isolates had low laccase activity and low rhizomorph production, and A. gallica had a broad range of laccase activities and rhizomorph production. No peroxidase activity was detected in these studies; however, only one of the isolates was screened for peroxidase activity (Worrall and others 1986). Recently, Marsh and Wargo (1989) assayed phenol oxi¬ dases over time in three isolates from each of five bio¬ logical species of Armillaria: NABS I (A. ostoyae), NABS III (A. calvescens), NABS V (A. sinapina), NABS VI (A. mellea), and NABS VII (A. gallica). Laccase (tetra- methyl-benzidine=TMB=substrate) and peroxidase (TMB with and without catalase=substrates) were de¬ tected in extracts from mycelium and rhizomorphs and in the extra-cellular growth medium. Peroxidase activ¬ ity was confirmed by the lowering of oxidase activity when H,Ck in the extract was destroyed by adding 36 Ontogeny and Physiology catalase, and by assay with a substrate specific to per¬ oxidase activity, aminoantipyrine. Peroxidase activity was not detected in all isolates, and a broad range of activities among the isolates with detectable peroxidase activity did occur. Tyrosinase activity (dihydroxyphenylalanine=L-DOPA= substrate) was found only intracellularly. They detected a general re¬ lationship of higher laccase activity with greater rhizomorph production among rhizomorph-producing isolates. However, laccase activity was also present in some isolates that produced no rhizomorphs. Conclusions The foregoing discussion of nutritional and environ¬ mental factors affecting Armillaria indicates that prin¬ ciples of fungal nutrition and physiology may be applicable to some aspects of its behavior in soil and on infected hosts. On the other hand, the discussion of biochemical factors that regulate growth and develop¬ ment indicates major information gaps for fungi in gen¬ eral and Armillaria in particular. More basic information at the molecular and biochemical levels is needed to develop a good working hypothesis to ex¬ plain regulation of growth and morphogenesis in re¬ sponse to nutritional and environmental factors. When this information becomes available, more effective ap¬ proaches to manipulating Armillaria in culture, in soil, and on its many hosts may be forthcoming. Miscellaneous Themes in the Physiology of Armillaria Protease A protease with unique properties has been recovered from Armillaria (Broadbent and others 1972). This en¬ zyme cleaves peptide bonds which are N-terminal to lysine residues in proteins (Hunneyball and Stanworth 1975, Lewis and others 1978). This specificity for lysine residues in the protein is maintained even when the positive side chain of the lysine is formylated and thus neutral in charge (Barry and others 1981). The enzyme is very stable in the presence of denatur¬ ing detergents such as sodium dodecylsulfate. Because of this feature, the enzyme can be used to fragment proteins which are insoluble in water but can be solubi¬ lized by the addition of detergent (Barry and Doonan 1987). No information is available on the biological role of the enzyme. Whether it is secreted into the environ¬ ment or present at unique points in the developmental cycle, such as during basidiome formation, is not known. Antibiotics and Other Metabolites In 1951, Armillaria was observed to exhibit considerable antibiotic properties when cultivated either on wood, solid media, or liquid media (Oppermann 1952). Armillaria antibiotics inhibited other fungi as well as bacteria. These findings were confirmed by Richard (1971). Later, Ohr and Munnecke (1974) found that the production of these antibiotics was considerably re¬ duced when Armillaria was fumigated with sublethal concentrations of methyl bromide. The authors sug¬ gested that this is one reason for the effect of soil fumi¬ gation. It may predispose Armillaria to attack by biological control agents such as Trichoderma that would otherwise be restricted by the fungus' own anti¬ biotics (see chapter 11). The chemical nature of the antibiotic substances and other metabolites produced by Armillaria was eluci¬ dated in subsequent years by several groups of scien¬ tists. Oduro and others (1976) isolated four chloroform-soluble substances for which antibiotic ac¬ tivities were determined by bioassays with either Bacil¬ lus sp. isolated from fumigated citrus roots naturally infected by Armillaria or cultures of Cladosporium cucumerinum Ellis and Arth. The authors were able to show that antibiotic activity was produced by all 17 Armillaria isolates used. Detailed studies by several authors (Ayer and MacCauley 1987, Donnelly and others 1982, Jungshan and others 1984, Midland and others 1982, Obuchi and others 1990) have revealed that various isolates of Armillaria have at least 10 different compounds with antibiotic properties. Two aspects of the chemical na¬ ture of these substances are rather interesting. First, they are mostly complicated sesquiterpenoid esters, some belonging to the protolludane group. The organic acid to which they are bound is, suprisingly, the same substance which has been identified as the antibiotic substance of Sparassis crispa Wulf.: Fr. (Falck 1907,1909, 1924, 1930). Second, these compounds contain a rather simple aromatic, Sparassol or orsellinic acid, which in all tests exhibits high antifungal and antibacterial activ¬ ity (Cwielong 1986). Apparently, Armillaria uses the same chemical weapon as does S. crispa with the modi¬ fication that sesquiterpenoids are attached to the aro¬ matic group. Thus, the Armillaria antibiotics would penetrate more easily through membranes and would probably be more toxic than the unsubstituted Sparassol. The variety of antibiotic substances produced by Armillaria and their high toxicity against microbes may explain, in part, why this fungus is so successful in its natural habitat and also some of its medicinal proper¬ ties. For example, folklore of early American loggers Ontogeny and Physiology 37 tells of woodsmen who would wrap their wounds from accidental cuts in an Armillaria fan. This pro¬ tected them from further irritation and enhanced healing. Also, tablets containing artifically cultured mycelia of Armillaria are used in China for treating of dizziness, headache, neurasthenia, insomnia, numb¬ ness in limbs, and infantile convulsions (Jungshan and others 1984). Bioluminescence Bioluminescent fungi have interested biologists for some time (Glawe and Solberg 1989). More recently, at¬ tention has been given to the biochemical mechanisms involved (Airth and others 1966, Danilov 1987). Armillaria is one of several bioluminescent basidiomy- cetes (Guyot 1927). Airth and Foerster (1960) prepared a self-portrait of a 15-day culture of Armillaria that showed high luminescence in the peripheral region (young cells) and less in the central area (older cells). A similar, more precise study using photomicrography and a different species of Armillaria (Berliner and Hovnanian 1963) showed light emission occurred throughout the entire cell. The characteristics of the light emitted by Armillaria and other fungi have been investigated. Airth and Foerster (1960) noted the emission maximum of 528 nm was similar to that of other fungi but different from that of bacteria. They found that the energy of activa¬ tion for emission in Armillaria is 17,500 calories with a temperature optimum of 26°C. Berliner (1961) sug¬ gested that fungi which exhibit bioluminescence may emit some waste energy of oxidation as light instead of heat. Also, Berliner noted that Armillaria took a longer time than other fungi studied to attain maximum light emission values, but sustained luminescence of 10 weeks equaled or exceeded that of other fungi. Effect of Environment, Nutrition, and Growth Factors The effects of temperature, exposure to X rays and ul¬ traviolet light, nutrition, and growth factors on lumi¬ nescence in Armillaria and other fungi have been reported. Temperature Ultraviolet and X-irradiation Ultraviolet irradiation inhibited light emission from Armillaria and other fungi (Airth and Foerster 1960, Berliner 1963, Berliner and Brand 1962). The effects ob¬ served varied with the wavelength of incident radia¬ tion, the time elapsed, and the fungal species used. In contrast, X-irradiation enhanced luminescence from Panellus (Panus) stipticus (Bull: Fr.) P. Karst. (Berliner 1961) and probably would produce a similar effect on Armillaria. Nutrition The relationship between light emission and nutrition has been reviewed (Flarvey 1952). Airth and Foerster (1965) reported a specific pH and nitrogen source for optimum light emission by Collybia velutipes (Fr.) Sing. On this basis, optimal nutritional conditions for maxi¬ mum light emission presumably exist for Armillaria as well. Growth Factors Luminescence in Armillaria responds to growth factors according to the concentration and type of factor used. For example, the light output was intensified more than 150% when Armillaria was grown on a medium containing 0.75 mg/1 of biotin. Also, kinetin at 0.25 mg/1 increased light output, but 6-benzylaminepurine had no effect (Berliner and LaRochelle 1964). The ef¬ fects of antibiotics on light emission have also been studied (Berliner 1965). Mechanism of Fungal Bioluminescence Studies with Armillaria and other fungi have identified the key biochemical steps involved in fungal biolumi¬ nescence. For example, Airth and Foerster (1962) pre¬ sented evidence that fungal bioluminescence involves the following: (a) either reduced nicotinamide adenine dinucle¬ otide (NADH) or reduced nicotinamide adenine di¬ nucleotide phosphate (NADPH); (b) an electron acceptor found in hot water extracts; (c) soluble dehydrogenases; (d) molecular oxygen; (e) the particulate enzyme luciferase. idit emission was low at -10°C and low or non-exis- ' e U)°C (Airth and Foerster 1960) with the opti- mfure in the range of 18-26°C. Berliner lilar optimum temperature for light al basidiomycete fungi including The proposed reaction involved in light emission is: 2NADH + X soluble enzyme > XH, + 2NAD + yu , 1/0 particulate 2 1 2 luciferase > X + HOH + light 38 Ontogeny and Physiology The similarities and differences of light emission be¬ tween fungi and bacteria have been noted (Airth and others 1966). However, fungal and bacterial biolumi¬ nescence and chemoluminescence may have close links not only in their physical nature but in their biochemi¬ cal nature as well. Physiology of Host-Pathogen Interactions Understanding the physiological bases for pathogen¬ esis and the interactions of Armillaria species with their hosts is the key to understanding the variation in pathogenicities among and within the species of Armillaria that we now know. Unfortunately, much of the work that has been conducted in this area lacks es¬ sential taxonomy of the fungus. Results of these stud¬ ies, therefore, may reflect the physiology of a single species, one or several genotypes within a species, or several different species all interacting with hosts that may or may not be resistant. Our current understand¬ ing, and hence what is presented herein, of what stimu¬ lates and controls penetration and colonization of a substrate by Armillaria is incomplete for any single spe¬ cies. What we know is probably a composite of several different Armillaria species interacting on susceptible and resistant hosts. Genetic Control The infection processes, resistant reactions, pathogenic¬ ity and virulence, and disease development within the host tree are discussed in chapters 4, 5, and 6. These processes represent host-pathogen interactions and in¬ volve the physiology of metabolic regulation of the fungus and host. Metabolic control of these interactions is determined by the genetic control of the physiologi¬ cal processes as modified by the environment (Daly 1976). The reaction of host and fungus, therefore, depends on the host species that is attacked, the species and per¬ haps genotype of Armillaria that is attacking, and the environmental conditions under which host and fun¬ gus are growing. Most historical information on host- pathogen interactions focuses on the differences in response among host species. Little attention has been paid previously to differences in the pathogen since it was considered for the most part to be a single species. Now that several species of Armillaria are recognized with different pathogenic capabilities on different hosts (Davidson and Rishbeth 1988; Rishbeth 1982,1985b), previous reports on host-pathogen interactions must be re-examined. The infection process is both mechanical and enzy¬ matic. Since penetration of the outer bark is reportedly similar in both the susceptible and the resistant reac¬ tions, subsequent colonization of the inner bark and cambial zone tissues differentiates the susceptible from the resistant reaction (Thomas 1934). These observa¬ tions are based on reactions of hosts with single isolates of unknown species of Armillaria, although some at¬ tempts have been made to assign species names to some isolates used in these historical studies (see chap¬ ters 4 and 6). Whether all species of Armillaria can suc¬ cessfully penetrate the outer bark is not known. Wounding of the roots can enhance infection by Armillaria (see chapters 4 and 7), and perhaps some species of Armillaria are unable to penetrate intact bark. Metabolic Control Little work on the metabolism of Armillaria species in association with their hosts has been conducted. Therefore, mostly metabolic capabilities of Armillaria and their potential for interacting with hosts are re¬ ported here. Pathogen Factors Suberinase Bark apparently offers limited resistance to penetration by Armillaria. Even periderms formed in response to the penetrating hyphae are unable to contain its growth (Rykowski 1975, Thomas 1934). The fungus can appar¬ ently grow faster than developing periderms and in¬ vades around them (Rykowski 1975) or penetrates directly through the periderms, probably by enzymatic activity (Arthaud and others 1980, Rykowski 1975, Thomas 1934). Armillaria can degrade suberin. Swift (1965) reported that the fungus, grown on ground bark of Brachystegia spicaeformis, caused a 59% loss in suberin content of the bark. Armillaria also produced hydrolytic enzymes when grown for 10 months on 0.5% raspberry suberin medium supplemented with salts, thiamine, and ethanol (Zimmermann and Seemiiller 1984). Con¬ centrated enzyme preparations from culture fluids caused up to 1% dry weight loss of suberin prepara¬ tions after 16 hr incubation. Gas chromatographic analyses of the released material indicated that the components constituted a major part of the aliphatic monomers present in suberin (Kolattukudy and others 1981). How important suberin degradation is in the in¬ fection process is uncertain. Polyphenol Oxidases Armillaria produces phenol oxidases during the infec¬ tion process. Discoloration, especially browning of Ontogeny and Physiology 39 tissues, has been observed commonly during the infec¬ tion and colonization process (Rykowski 1975; Thomas 1934; Wargo 1977,1984a). Discolored bark in advance of colonized bark in black and white oaks had signifi¬ cantly less total phenols and more oxidized phenols than contiguous or noncontiguous healthy bark (Wargo 1984a). In colonized bark, total phenols were only 22% and 46%, respectively, of that in healthy bark of black and white oaks; and oxidized phenol levels were 3 and 3.5 times greater than in healthy bark (table 3.6). Phenol levels in discolored bark from wounded only bark tissues were also lower after 4 weeks than in healthy contiguous bark, but not as low as in colonized bark. Levels of oxidized phenols in discolored bark from wounded-only tissues did not increase as much as in colonized tissues. Oxidation of the phenols in root tissues can result from both fungal and host polyphenol oxidases. No reports distinguish between host and fungus-mediated phenol oxidation. Fungal enzymes can oxidize phenols as a result of separate or combined effects of peroxidase, tyrosinase, or laccase depending on the phenolic sub¬ strates. Annillaria possesses all three enzyme activities and peroxidase and laccase can be secreted to oxidize phenols extracellularly, as described previously in this chapter. Very limited information details the role of phenoloxidizing enzymes in the pathogenic process. Marsh and Wargo (1989) screened three isolates each of A. ostoyae, A. calvescens, A. sinapina, A. mellea, and A. gallica for production of constitutive phenol oxidases. Many, but not all, of these isolates were rated by other researchers in pathogenicity studies. The pathogenicities of the remaining isolates were rated by Marsh and Wargo as high, moderate, or low, based on their association with the host tree from which they were isolated. No obvious correlations of constitutive enzyme levels with pathogenicity were detected. Phenols and other host substances can inhibit hydro¬ lytic enzymes of fungi, thus restricting their activities on host cell walls and membranes and preventing in¬ fection and colonization. Polyphenol oxidases cause the oxidation and polymerization of compounds that are potentially toxic to the fungus, allowing infection and colonization to proceed in tissues rich in phenols. This reaction is apparent at the leading edge of mycelial fans colonizing living tissue. Here, an advancing band of oxidized (browned) tissue precedes the advancing mycelium (fig. 3.8). There is some evidence that these brown pigments induce wilt in infected plants. Thornberry and Ray (1953) isolated a dark brown pro¬ tein-like pigment produced by ArmiUaria in liquid me- TABLE 3.6 — Changes in mean concentrations of soluble phenols and their oxidation products effected by ArmiUaria in bark of roots of black and white oak trees naturally colonized by the fungus. Tannins 1 Total Hydrolyzable Condensed Species and tissue state Phenols 1 total mg phenols/g tissue Phenols 2 oxidized Black Oak 238a Healthy, control 167a 128a 143a 13a Healthy,contiguous 161 ab 124ab 136a 13a 243a Discolored 145b 107b 61b 11a 306a Colonized 37c 31c 22c 8b 731b SE ±5 ±5 ±3 ±2 ±30 White Oak 352a Healthy, control 196a 147a 147a 15a Healthy,contiguous 170a 136a 160a 8b 62 lab Discolored 158a 124a 107b 9b 742b Colonized 90b 67b 63c 1 lab 1235c SE ±10 ±10 ±8 ±2 ±80 Vargo (1984a) p.' and hydrolyzable tannins - mg tannic acid equivalents/g freeze-dried bark: condensed tannin - mg catechin equivalents/g. Significant differences by ANOVA and Tukeys studentized range test (P<0.05) indicated by different letter. 100 mg bark in 10 ml water at 450 nm and 1 cm light path used as estimate of oxidized phenols 40 Ontogeny and Physiology FIGURE 3.8 — Discolored brown zone in both bark and wood in advance of the mycelium. Note rhizomorphs on surface of primary root. (P. Wargo) FIGURE 3.9 — Advanced decay of root wood by Armillaria (also note discolored brown zone in advance of mycelium). (P. Wargo) dium. The pigment induced wilt in tomato seedlings and peach twigs at low concentrations. There is, how¬ ever, no evidence that this mechanism operates in large mature trees. These phenoloxidizing enzymes are also important in wood degradation (fig. 3.9). Armillaria is classified as a white-rot fungus because it degrades and removes the lignified material from the cells, leaving the white cel¬ lulose and hemicelluloses somewhat intact (Campbell 1931,1932). Campbell also found that decay of wood by Armillaria was somewhat atypical of most white-rot fungi in that lignin degradation in laboratory tests was limited compared to cellulose degradation. Scurti (1956), however, grew Armillaria in vitro on pure cellu¬ lose and pure lignin, and observed that lignin was de¬ graded but not cellulose. Whether these results reflect differences among species of Armillaria cannot be an¬ swered. The ability to decay wood is probably quite different among species of Armillaria, and studies with known species are necessary. Marsh and Wargo (1989) found that some species of Armillaria produced high constitu¬ tive levels of an H n O,-enhanced phenol oxidase in vitro. This enzyme may be a lignin-degrading enzyme similar to the one found in the decay fungus Phanerochaete chrysosporium Burds (Tien and Kirk 1984). This ability of Armillaria to decay wood after it has pen¬ etrated and killed the cambial tissues allows the fungus to maintain itself in woody tissues. Here it may build up inoculum potential and overcome the resistant reac¬ tions in the living cambial zone tissues, or infect and kill additional tissue when the tree is weakened by stress (fig. 3.9). Host Factors Physical barriers probably slow the penetration and in¬ fection of root tissue by Armillaria, but they do not pre¬ vent infection. Resistance is therefore mostly chemical as either preformed constituents in the bark or as mobi¬ lized constituents in response to penetration by the fungus. Limited work by Wargo (1984a) indicated that no increase in concentration of total or specific phenols occurred in bark tissues contiguous with bark naturally colonized by Armillaria or wounded and inoculated with the fungus. Since total bark was analyzed, the in¬ crease in phenols may have been masked. Other work indicates that phenol accumulation in bark tissue in re¬ sponse to fungal colonization occurs primarily in the inner bark regions (Ostrofsky and others 1984, Wargo 1988). Preformed phenolics and other constituents can prob¬ ably act as effective chemical barriers to penetration and infection by Armillaria. In vitro studies with Armillaria have shown that some phenols commonly found in both coniferous and deciduous hosts can in¬ hibit fungus growth. Fifteen North American isolates representing at least the four species A. mellea, A. gallica, A. ostoyae, and A. sinapina (Wargo unpubl.) were challenged with hydrolyzable tannin (tannic acid, gallotannin) and gallic acid (Wargo 1980a). The isolates were both stimulated and inhibited depending on the phenol, the concentration of glucose, and the presence or absence of ethanol in the growth media (fig. 3.10). In general, gallic acid was more inhibitory to growth while hydrolyzable tannin was more stimulatory com¬ pared to the control. The ability to oxidize the pheno¬ lics seemed to be the key to inhibition or stimulation. Growth was inhibited if the isolate could not or only slightly (as determined by browning of the medium) oxidize the phenol. Growth was stimulated greatly where oxidation occurred readily; oxidation was initi¬ ated or accelerated by the addition of glucose and etha- Ontogeny and Physiology 41 FIGURE 3.10 — Growth of an Armillaria isolate on gallic acid GA, C+ET, C). Top: 1 g glucose/I; Middle: 5 g glucose/I; (GA) and control (C) media amended or not amended with Bottom: 10 g glucose/I. (P. Wargo) ethanol (ET) and with three glucose levels (left to right, GA+ET, nol. Isolates of A. gallica oxidized gallic acid and grew better in its presence with or without ethanol than did isolates of A. ostoyae. Wargo (1981 d) also observed that some ponderosa pine isolates of A. ostoyae from the Western United States that were pathogenic on the pine (Shaw 1977) could not i i/e gallic acid and did not grow at all on malt agar .Jed - ith 0.5% (w/v) gallic acid. Some less patho- m hardwood isolates (probably A. gallica, ■re able to oxidize gallic acid and re- :o eastern hardwood isolates that be- ; ?r tress has altered the tree ' Shaw (1985) could not confirm these reactions to gallic acid. He found that gallic acid both with and without ethanol inhibited most (20/21- dry weight, 21/21-colony diameter) of the 21 Armillaria isolates tested representing A. mellea (4), A. ostoyae (4), A. gallica (5), NABS V (3), A. luteobubalina (3), and A. no- vae-zelandiae (2). Variation within a species was as great as among the species. Growth of all isolates was stimu¬ lated on tannic acid medium (hydrolyzable gallo-tan- nin) without ethanol; with ethanol, a few isolates (4/ 21) grew less. The different response of A. ostoyae isolates to gallic acid in both studies (Shaw 1985, Wargo 1981 d) may have resulted from the different inocula used. Wargo 42 Ontogeny and Physiology used inoculum growing on water agar and Shaw used inoculum from 3% malt agar. The isolates on malt agar may have been conditioned to produce laccase (malt agar turns brown when Armillaria isolates grow in it, indicating oxidase activity) and were able to oxidize some gallic acid immediately. Also, Shaw amended 3% malt agar with gallic acid while Wargo used 2% malt agar. The difference in nutrient concentration could have affected the abilities of the various isolates to oxi¬ dize gallic acid (Wargo 1980a). Cheo (1982) also ob¬ served a carbohydrate effect on Armillaria growing on tannin-supplemented media. Growth of a single isolate with 0.5% tannin was 1.5 to 5 times greater when glu¬ cose was added to the medium. The stimulation of Armillaria species by tannic acid and the inhibition by gallic acid suggests that the concentra¬ tion of gallic acid and the rate at which it can be oxi¬ dized controls the response of the fungus. Tannic acid has approximately one glucose molecule for every five gallic acid molecules. The fungus may hydrolyze tan¬ nic acid to gallic acid, which it then oxidizes and poly¬ merizes immediately. This prevents the gallic acid concentration from becoming inhibitory. Alternatively, the fungus may oxidize the tannin without hydrolyz¬ ing it, thus preventing gallic acid from building up in the substrate. No work has been conducted on degra¬ dation of tannins by Armillaria. Analyses of phenols and tannin degradation in oak bark tissues colonized by Armillaria showed that gallic acid did not occur in colonized tissue (Wargo 1984a). Gallic acid and various polymers (di, tri, etc.) of gallic acid were present in the healthy and discolored tissues contiguous with the colonized portion but these materials decreased in the colonized bark compared to healthy tissues. This sug¬ gests that Armillaria oxidizes tannic acid and other polymers of gallic acid but does not hydrolyze them to gallic acid. However, this needs to be verified with more critical experiments. The ability of Armillaria to oxidize gallic acid, tannic acid, and other phenols in bark tissues is also influ¬ enced by carbon and nitrogen concentrations (Wargo 1983b). The growth rate and hence oxidation rate of phenols in extracts from root bark of black oak de¬ pended on supplemental glucose and nitrogen. Growth was directly proportional to the decrease in level of to¬ tal phenols in a culture medium, and was five times greater in the phenol plus supplement medium than in supplement alone. Phenols other than gallic acid and gallotannins also can inhibit Armillaria species. Both A. ostoyae and A. gallica were inhibited by various monophenols and alpha pinene, a terpene in conifer resins (Entry and Cromack 1989). Low levels of these phenols (<1 mg gT) stimu¬ lated rhizomorph production. No differences occurred between the two Armillaria species in response to the various phenols or pinene; variation of growth re¬ sponse to each compound was as great within as be¬ tween species. These results must be accepted very cautiously because the compounds were dissolved in 50 ml ethanol and added to 1 1 of medium. This concen¬ tration of ethanol is 30 to 100 times greater than con¬ centrations used in other studies. Results could be confounded by these high concentrations. Alkaloids are also known to inhibit Armillaria. Greathouse and Rigler (1940) found that alkaloids from several plant families inhibited growth of Armillaria in vitro. Other plant constituents have been found highly stimulatory to Armillaria. Lipids from roots of pon- derosa pine, Douglas-fir, white fir, incense-cedar, and peach promoted vigorous growth in vitro of an Armillaria isolate from California, probably A. mellea (Moody and Weinhold 1972a,b). The fatty acid fraction of the lipids was the active portion. Resin acids from ponderosa pine also were highly stimulatory and pro¬ moted twice as much rhizomorph growth as the fatty acid fraction from the same amount of root tissue. Abietic acid, a commercially available resin acid, stimulated rhizomorph production when it was steril¬ ized by autoclaving but not by filtration, suggesting that breakdown products of the acids are the stimula¬ tory factors. Fresh or autoclaved wound resin from ponderosa pine also stimulates in vitro growth of Armillaria (Shaw 1975) and has been used in medium prepared for cultural paring tests (Shaw and Roth 1976). Predisposition Effects Stress Susceptible or resistant responses of the host to a fun¬ gal pathogen depend on the genetic makeup of the host and the pathogen, and the environment in which they exist. Stress can alter the relationship and change the balance in the interaction between host and pathogen, resulting in root disease. Stresses obviously affect the pathogen, but few studies report on these effects. We know that drought and wa¬ terlogging sometimes increase the incidence and sever¬ ity of Armillaria root disease (see chapter 7). However, we have no idea how drought or waterlogging affect the fungus when it occurs as rhizomorphs in the soil or as mycelium inside tree tissues. For example, we do not know how turgor pressure in the rhizomorph influ¬ ences penetration of the root bark; nor do we know how moisture extremes influence this relationship. Nechleba (1915), in his conclusions regarding the pathogenic relationship of trees and Armillaria, specu- Ontogeny and Physiology 43 lated that dry conditions in forests promoted infection and colonization by inducing rhizomorphs of the fun¬ gus to colonize other substrates for water and nutri¬ ents. He proposed that the rhizomorphs "find their way instinctively (hydrotropism) toward living roots" and colonize them. Armillaria species infect roots of healthy trees by rhizomorph contact, from diseased tissue, or by direct mycelial contact from diseased roots (see chapters 4 and 6). Hyphae penetrate the outer bark and "chal¬ lenge" the inner bark tissue; it is here where stress in¬ fluences the reaction. Chemical changes induced in the host by stress may promote susceptibility by (1) remov¬ ing fungal inhibitors, (2) releasing nutrients and me¬ tabolites required by the fungus for pathogenesis, (3) providing the fungus with growth stimulators that al¬ low it to overwhelm the capacity of the host root sys¬ tem to resist harmful fungal metabolites, or (4) reducing the capacity of the host tissues to tolerate or control the metabolites produced by the fungus (Wargo 1984b). All or any combination of these relationships may occur. Many stresses predispose trees to Armillaria and initiate root disease or accelerate root disease in the host (see chapter 7). However, our knowledge about how stress specifically affects the relationship between Armillaria and its hosts is mostly about the host and is limited predominantly to the effects of drought and insect damage on a few host tree species (Wargo 1983a,b; 1984a,b). Nutritional Changes Both drought and defoliation affect the carbohydrate and nitrogen levels in the root tissues colonized by Armillaria (Gregory and Wargo 1986, Parker 1979, Parker and Houston 1971, Parker and Patton 1975, Wargo 1972, Wargo and others 1972). Defoliation can substantially decrease the starch content in the root wood (fig. 3.11) and decrease sucrose levels in both bark and cambial tissues of sugar maple roots (Wargo 1972,1981b). Reducing-sugar levels increase especially in cambial zone tissues. Concentrations of reducing- sugar can be 4-5 times higher in defoliated trees than those in non-defoliated trees at the same time of the year, and 3-4 times higher than the normal spring high when carbohydrates are mobilized for growth (Wargo 1971). Since Armillaria predominantly uses glucose (Garraway 1975, Wargo 1981a), this increase is poten¬ tially important to the fungus. Growth on glucose or polymers of glucose, such as maltose and starch (fig. 3.12), can be 1.5-3 times higher than growth on other carbon sources (Wargo 1981a). Enhanced growth of A. calvescens (Wargo unpubl.) on root extracts of defoli¬ ated sugar maples was related to higher levels of glu¬ cose in the extract (Wargo 1972). DATE OF DEFOLIATION DATE OF TREE HARVEST dine in sucrose and starch content in the Sucrose level in the inner bark; B: Starch level in wood. (From gar maple roots caused by defoliation. A: Wargo 1972) 44 Ontogeny and Physiology GROWTH OF ARMILLARIA MELLEA 0 N DIFFERENT SOURCES O - STARCH £ - SUCROSE 0 - FRUCTOSE — GLUCOSE TOTAL GRAMS CARBOHYDRATE GROWTH OF A RH I L LA R 1 A MELLEA q OM HEDIA CONTAINING 5g GLUCU C E D AND OTHER SUGARS X _ starch GROWTH OF ARM ILLARI A MELLEA Q OM MEDIA CONTAINING 5g SUCROSE AND OTHER SUGARS FIGURE 3.12 — Growth in vitro of Armillaria on various carbohydrates that demonstrate the stimulation of growth by glucose. A: Growth on various carbohydrates; B: Growth on Drought and defoliation also increase both total amino nitrogen levels and certain individual amino acids in sugar maple trees (Wargo 1972) and seedlings of black and red oak (Parker 1979, Parker and Patton 1975). Both individual amino acids and total animo nitrogen supplements were very satisfactory nitrogen sources for in vitro growth of Armillaria (Weinhold and Garraway 1966), as discussed previously. Also, noted earlier, ethanol is a potent stimulator of Armillaria, especially rhizomorph production and growth (Weinhold 1963, Weinhold and Garraway 1966). In the presence of ethanol, the fungus can me¬ tabolize phenolic compounds that would otherwise in¬ hibit growth (Longworth and Garraway 1981; Wargo 1980a, 1981d). Ethanol enhances laccase production by the fungus (Worrall and others 1986) and improves its ability to utilize carbon sources other than glucose (Weinhold and Garraway 1966). Ethanol could be an important factor in stressed trees. Stress from flooding or defoliation can stimulate etha¬ nol production and accumulation in woody roots (Wargo unpubl.). On poorly drained sites and more mesic areas, seasonally high water tables often occur and cause anaerobic conditions about tree roots. Defo¬ liation, because it reduces transpiration, promotes or glucose media supplemented with various carbohydrates; C: Growth on sucrose media supplemented with glucose and fructose. (From Wargo 1981a) prolongs wet soil conditions. In oak forests in Con¬ necticut, soils in stands defoliated by the gypsy moth (.Lymantria dispar L.) were wetter and defoliated trees contained more water than soils and trees on adjacent nondefoliated sites (Stephens and others 1972). Signifi¬ cant amounts of ethanol can be produced in roots de¬ pending on the duration of the anaerobic conditions and tree species (Coutts and Armstrong 1976, Crawford and Baines 1977). Injection of ethanol into roots of black and white oaks promoted colonization of the roots by Armillaria. Colonization, however, was related more to tissue necrosis caused by the ethanol rather than to the ethanol alone (Wargo and Montgom¬ ery 1983). Phenol Degradation Stress-induced chemical changes in roots may also de¬ termine how well Armillaria can oxidize phenols. Inhi¬ bition of Armillaria growth by gallic acid was lessened or reversed by adding more glucose to the medium (Wargo 1980a). Growth in bark extracts from black oak roots depended on phenol oxidation, which was greatly enhanced by adding glucose and nitrogen to the extract (Wargo 1983b). Additional growth studies using commercial sources of phenols found in oak bark (quercetin, quercitrin, catechin, and tannic acid) indi- Ontogeny and Physiology 45 Host-Induced Lysis FIGURE 3.13 — Growth of an Armillaria isolate on an extract from red oak bark. Upper flask—extract + glucose + ethanol. Lower flask—as above + 500 ppm ascorbic acid. (P. Wargo) cated that if the fungus could oxidize the phenol, the phenol no longer inhibited the fungus (Wargo unpubl.). Growth was also stimulated, suggesting that the oxidized phenols were being utilized as carbon sources or growth regulators. If oxidation of the phenols were inhibited by adding a reducing agent (fig. 3.13), growth significantly declined (Wargo unpubl.). Successful colonization of root tissues in stressed trees may depend on the fungus' ability to oxidize phenols and the inability of the tree tissue to prevent the oxida¬ tion reaction. In healthy deciduous trees, Armillaria ap¬ pears to be confined to wounded and necrotic tissue; contiguous healthy tissues are not "browned" or colo¬ nized by the fungus. In weakened trees, contiguous liv¬ ing tissues are "browned" in advance of the fungus, probably by extracellular secretions of laccase and per¬ oxidase, and then colonized (Wargo 1983b, 1984a). This interaction has similarities to that proposed for the re¬ dox theory of hypersensitivity reaction (Goodman and others 1986) where necrosis in response to fungal inva¬ sion occurs when the balance between reductive and oxidative processes shift in favor of the latter. In healthy tissues, necrosis induced by Armillaria is inhib¬ ited or contained, probably by a highly reductive state in contiguous tissues. Perhaps stressed tissues cannot confine the oxidative processes and necrosis begins and ■■oreads as oxidative and other enzymes are secreted by the fungus. Host-produced enzymes that may potentially assist bark tissue in resisting Armillaria are also affected by stress from defoliation (Wargo 1976). The hyphal walls of Armillaria contain chitin and beta-1,3-glucan, and are vulnerable to lysis by chitinase and beta-1,3-glucanase (Ballesta and Alexander 1972, Bouveng and others 1967, Wargo 1975). These enzymes are found in bark and sap of several oaks and sugar maples, and their ac¬ tivities are lowered by defoliation (Wargo 1975,1976). Lysis of Armillaria hyphae in vivo has been reported for species associated with orchids (Hamada 1940, Kusano 1911) and the description of fungal digestion in orchid species suggests a host-mediated lysis (Burges 1939). Complete dissolution of the hyphae is not necessary to disrupt growth. Hyphal tips grow by a delicate balance between wall synthesis and wall lysis, and bursting of the hyphal tips can occur when the balance shifts to¬ ward the lytic stage (Bartnicki-Garcia and Lippman 1972). Extrahyphal enzymes in host cells that can dis¬ solve hyphal wall components could alter the wall for¬ mation balance, disrupt hyphal-tip growth, and provide a defense mechanism against invasion by fun¬ gal pathogens. More recent work on these enzymes in¬ dicates that they are indeed potent inhibitors of fungal growth (Schlumbaum and others 1986). The fungus is not defenseless against lysis by host-pro¬ duced enzymes. The phenol oxidase enzymes, espe¬ cially tyrosinase, produced by the fungus are linked to melanin synthesis by fungi (Mayer and Harel 1979). As noted earlier, Armillaria is capable of producing mela¬ nin-like pigments in rhizomorphs and probably to a limited extent in hyphae (Chet and Hiittermann 1977, Smith and Griffin 1971). Phenol oxidase-catalyzed for¬ mation of extracellular pigments may be related to the formation of melanin-like pigments in hyphae. They may strengthen hyphae (Bell and Wheeler 1986) and protect them from dissolution by lytic enzymes (Bloomfield and Alexander 1967). Conclusions Host-pathogen interactions ultimately depend on the relationship of fungal species, host species, and the en¬ vironment in which they interact, including the distur¬ bances induced by stress. Much of the information on the physiological and chemical interactions of Armillaria species and their hosts is fragmented, and the characteristics of the events for any one species of 46 Ontogeny and Physiology Armillaria and its host are incomplete. The fungus pen¬ etrates generally through intact bark, interacts with the inner bark, is stimulated to colonize and kill the inner bark, and either invades the cambial zone or is inhib¬ ited by as yet unknown mechanisms. The interaction with phenols present in the bark tissues is probably a major event in determining resistance or susceptibility and the pathogenic process. Stress from a variety of sources influences the resistance mechanisms and en¬ hances penetration, colonization, and killing by Armillaria. The concepts discussed in this section are based on fragments of information concerning the many interac¬ tions that can occur among the many Armillaria species and host species. Studies using clonal host material, known species, and genotypes of Armillaria and stressed and non-stressed systems must be conducted to elucidate the kinds and sequence of pathogen and host changes that occur in resistant and susceptible re¬ actions. Some of the morphological and anatomical in¬ teractions have been characterized. These must be verified in the host-pathogen system described above, and the chemical changes associated with these interac¬ tions must be characterized. This area of research is ripe for much work by the students of host and fungal physiology and their interactions. Ontogeny and Physiology 47 CHAPTER 4 Inoculum and Infection Derek B. Redfern and Gregory M. Filip A ll Armillaria species survive saprophyti- cally in woody substrates in soil, and the majority form the most highly organized rhizomorphs of any fungus. By extension of these rhizomorphs through the soil, the fungus can colonize additional woody material. Varying degrees of pathogenicity may be exhibited during this phase. Robert Hartig (1873b) was the first to not only make the link between the spread of infection and the presence of nearby trees previously killed by the fungus, but also to suggest that rhizomorphs cause infection. Descriptive terms such as "food base" and "invasive potential" have obvious application to the rhizomorph- forming Armillaria species. "Inoculum potential" is a similar term. This concept was explored by Garrett (1970), partly through a series of experiments with A. mellea (sensu lato) (Garrett 1956b). The term was not new, but he redefined it (1970) as "the energy of growth of a parasite available for infection of a host, at the surface of the host organ to be infected." The defi¬ nition encompasses the net effect of variables such as the surface area of fungus in contact with unit area of host, the vigor of the invading hyphae, and environ¬ mental effects on the fungus. This chapter deals primarily with factors that affect the success of infection through their effect on inoculum potential. First, the nature of the inoculum capable of causing infection and the quality of the substrate pro¬ vided by different tree species are considered. The sec¬ ond part concentrates on those factors which affect the success of infection through their effect on the fungus, particularly the rhizomorphs, which provide the means of infection and spread in most Armillaria species. Inoculum Source of Inoculum poses, wood provides the only ef- : which Armillaria can spread and cause infection. Tree roots constitute the major source of inoculum, but logging debris may also be colonized and act in the same way (MacKenzie and Shaw 1977). The fungus becomes established in roots and stumps by infecting live trees and by colonizing stumps cre¬ ated during felling operations. If a tree is killed, the en¬ tire root system may become inoculum. The fungus colonizes newly created stumps in three ways: by rapid extension from pre-existing lesions in which it was for¬ merly held in check by host resistance (Kile 1980b, Leach 1939); by invasion from an epiphytic position on the roots; or by invasion from outside by newly arrived rhizomorphs. Based on Garrett's work (1960,1970), the series of cir¬ cumstances under which Armillaria becomes estab¬ lished in substrates can be taken to represent a requirement for a decreasing parasitic ability and an in¬ creasing competitive saprophytic ability. Logging resi¬ dues constitute an extension to the series because, apart from being less readily available for colonization than stumps by virtue of position, their tissues are likely to die more rapidly and be available earlier for coloniza¬ tion by competing saprophytic organisms. Where stumps provide potential sources of inoculum, they are most commonly colonized by vegetative spread, but the cut surface can also provide an avenue for colonization by basidiospores (Rishbeth 1970, 1978b, 1988). A number of researchers have failed to in¬ fect stumps in this way, however (Kile 1983b, Leach 1939, Podger and others 1978), while others have had very limited success (Swift 1972). It is apparently an uncommon event but may be important to disease de¬ velopment in certain crops (Horner 1988). Even though basidiospore-infected stumps probably constitute a mi¬ nor portion of the total inoculum, spore infection is im¬ portant for providing a source of genetic diversity, for facilitating long-range spread, and also for infecting forests established on arable land. Some work on geno¬ type identification (Hood and Sandberg 198/, Horner 48 Inoculum and Infection 1988, Kile 1983b, Ullrich and Anderson 1978) provides indirect evidence for spore infections, but similar work by others provides less support (Shaw and Roth 1976). No evidence indicates basidiospores can directly infect living roots, presumably because the inoculum poten¬ tial provided by the limited resource within spores is inadequate. Hartig (1874) suggested that basidiospores may colonize dead organic matter and subsequently form rhizomorphs, but no experimental evidence sup¬ ports this. In experiments, most successful infections have been achieved using woody inocula prepared either from naturally infected roots (Leach 1937) or by culturing the fungus in various ways on woody stem or root seg¬ ments (Patton and Riker 1959, Redfern 1975, Shaw 1977, Thomas 1934). Cultures established on non- woody substrates such as nutrient agar, bran, or bean pods have been generally unsuccessful as inocula (Bliss 1941, Plakidas 1941). Wood is not an absolute prerequi¬ site for infection; inocula derived from less substantial substrates may be adequate. For example, Guyot (1927) caused infection using cultures on an agar medium containing acorns and horse chestnuts. Nevertheless, only a woody substrate is able to provide an inoculum which is sufficiently durable and potent to cause dis¬ ease reliably. Under experimental conditions, infection has been achieved even without a substrate by means of excised rhizomorphs. These pieces can be large enough to form new growing tips with an inoculum potential high enough to infect healthy seedlings (Redfern 1973, Rykowski 1984). Holdenreider (1987) caused infection in a similar way but found wounds to be an apparent prerequisite. Other reports concerning the infective po¬ tential of detached rhizomorphs have been negative (Bliss 1941). In common with other root-rot fungi, Armillaria inocu¬ lum is generally confined to infested sites. However, roots may become fragmented and transported by wa¬ ter, thus potentially creating new foci of infection (Hewitt 1936). Colonized logging debris could be trans¬ ported in the same way. The rhizomorph-forming abil¬ ity of most species would enable Armillaria to exploit such an event much more effectively than other root pathogens such as Heterobasidion annosum (Fr.) Bref. and Phellinus weirii (Murr.) Gilbn. Substrate Quality—Conifers Versus Hardwoods Armillaria mellea sensu lato was considered to be a highly variable species long before the present under¬ standing of speciation in the genus and of the ecology of these species. In spite of this, much of the observed variation in disease was attributed to factors other than variation in pathogenicity. Prominent among these was the nature of the substrate providing the inoculum. Disease is now known to be associated with stumps of many species, ranging from Australasian hardwoods (Kile 1981, Podger and others 1978, Shaw and Calderon 1977) to European and North American conifers (Redfern 1975, Shaw and others 1976a). Early records, however, largely associated mortality with hardwood stumps. A possible reason for this is that until rela¬ tively recently the disease attracted most attention in fruit orchards and in plantations of tea, coffee, rubber, and exotic conifers, all established on land cleared of indigenous forest where Armillaria was endemic. In the tropics and sub-tropics, this original forest comprised a mixture of broadleaved species (Leach 1939). In tem¬ perate regions, hardwoods would probably have been at least a major component on the richer soils where such plantation crops were grown. Many early reports of disease concern losses in these circumstances (Butler 1928, Dade 1927, Gibson 1960, Hendrickson 1925, Horne 1914, Lawrence 1910, Nechleba 1915, Rhoads 1956, Wallace 1935). In California, the disease occurred so consistently in orchards planted on land cleared of oaks that for many years articles in Californian agricul¬ tural journals referred to Armillaria as the "oak root fungus" (Kimball 1949, Raabe and others 1967). In Europe, Hartig (1874) and Nechleba (1915) observed that serious disease may occur where conifer planta¬ tions replace hardwoods, whereas damage is generally unimportant in crops replacing conifers. This had a major influence on early thinking about how substrate affects disease development. The prevailing view was that hardwood stumps provide a superior substrate to conifer stumps. Peace (1962), for example, commented that Armillaria is essentially a fungus of areas with a hardwood history, and suggested that where conifers replace hardwoods damage is likely to be absent or much reduced in the second conifer rotation. During the first rotation, conifer stumps left after thinning are readily colonized by Armillaria (Greig 1962, Low and Gladman 1962), but Peace (1962) believed the fungus acts purely saprophytically in this situation and there is no increase in parasitic activity. The implication was that conifer stumps have little or no significance in sus¬ taining attacks. By contrast with observations implicating hardwood stumps as sources of infection, the first reports in which disease was clearly identified as being associ¬ ated with conifer stumps are relatively recent. Weiss and Riffle (1971) recorded killing of ponderosa pine fol¬ lowing a crop of the same species, and Swift (1972) re¬ ported losses in slash pine planted as a second rotation Inoculum and Infection 49 on a site formerly occupied by indigenous hardwoods. Ono (1965,1970) and Redfern (1975) reported serious disease where the major source of nutrition for the fun¬ gus was provided by conifer stumps. Initially, such ob¬ servations were rare among the continuing reports concerning hardwoods (Gladman and Low 1963, Huntly and others 1961, MacKenzie and Shaw 1977, Ono 1965, Pronos and Patton 1978, Swift 1972). They have become more numerous, particularly from natu¬ ral coniferous forests in North America (Morrison 1981, Wargo and Shaw 1985), as increasing interest in forest management draws attention to the impact of Armillaria losses. In the Northwestern United States, conifer stumps were shown to be effective inoculum sources (Filip 1979, Roth and others 1980) causing con¬ siderable infection and mortality in several indigenous coniferous species, especially in partially harvested for¬ ests (Filip 1977, Filip and Goheen 1984, Shaw and oth¬ ers 1976a). In experiments, trees have been successfully infected using inocula prepared from stems and roots of various coniferous and hardwood species, providing ample evidence of at least the short-term suitability of conifer¬ ous substrates as food bases for Armillaria. Species used include red pine and eastern white pine (Patton and Riker 1959); Japanese larch (Ono 1970); fig and citrus (Wilbur and others 1972); common beech, planetree and Scots pine (Redfern 1975, 1978); Sitka spruce (Singh 1980a); alder (Shaw 1977, Shaw and others 1981); and English oak (Morrison 1982b). While rhizomorph production may not be the best measure of substrate quality, particularly for those pathogenic species which produce few rhizomorphs, it has been commonly used. Thus, in experiments to de¬ termine the relative value of the substrate provided by roots of hardwood and coniferous species, Redfern (1970) found that segments of red maple inoculated with Armillaria produced a greater number, total length, and dry weight of rhizomorphs than red spruce segments of equal volume. However, when corrections were made for differences in initial wood density of the two species, differences in length and weight were no longer evident, although maple segments still pro¬ duced a greater number of rhizomorphs than spruce. Working with several Armillaria isolates and several co¬ niferous and hardwood species as substrates, Morrison (1972) found that, with the exception of one isolate, hardwood segments produced a greater dry weight of ■ 'morphs than conifer segments. He made a similar fi i density. The number of rhizomorphs was : '. this experiment, but when stumps were he field, Morrison found that the number \ stems, as well as the total length of ump, was greater for hardwood or :vi stumps. In a similar study. which included measuring rhizomorph production by naturally infected stumps, Rishbeth (1972b) concluded that pines are inferior to English oak as substrates for Armillaria in terms of the number and weight of rhizomorphs produced. In comparing maritime pine with oak, Guillaumin and Lung (1985) obtained the same results as Rishbeth for both A. ostoyae and A. mellea. Redfern (1975) examined the effect of substrate on in¬ fection as well as rhizomorph production. Sitka spruce seedlings were inoculated with four isolates of Armillaria growing on root segments of either planetree or Scots pine. Gregory (1985) subsequently identified these isolates to species. Armillaria ostoyae and A. mellea killed more trees when growing on planetree than on pine, whereas the reverse was true for A. gallica. Sub¬ strate species had no effect on A. cepistipes. Rhizomorph production was significantly greater on planetree than on pine for three of the species {A. ostoyae, A. gallica, and A. mellea), but A. cepistipes produced more on pine. Armillaria ostoyae and A. mellea were both highly patho¬ genic in the experiment, whereas the other two species showed very low pathogenicity. Thus, for both patho¬ genic species, rhizomorph production and infection were favored by a hardwood rather than a coniferous substrate. Rykowski (1984) obtained similar results in experiments with Scots pine seedlings and inocula pre¬ pared from branch segments. Hardwood substrates, es¬ pecially oak and common beech, were superior to Scots pine and European larch for both rhizomorph produc¬ tion and infection. Three isolates were used, but only one produced rhizomorphs consistently and caused in¬ fection. The species was referred to as A. mellea, but evi¬ dence in the paper suggests it was A. ostoyae. In similar work with the Australasian species A. novae- zelandiae and A. limonea, Benjamin and Newhook (1984b) ranked a number of indigenous and exotic hardwood species and two exotic conifers, radiata pine and ponderosa pine, as substrates for rhizomorph pro¬ duction. The two conifers occupied an intermediate position among the hardwoods as food bases for A. no- vae-zelandiae, whereas they were equal or superior to most of the hardwoods for A. limonea. Interestingly, the native hardwood tawa provided the best substrate for both species. In pathogenicity trials using the two Armillaria species with radiata pine and eucalypt seed¬ lings, radiata pine and several hardwood food bases were equally effective substrates when tested against radiata pine seedlings. Some evidence indicated that tawa was superior to radiata pine against eucalypt seedlings. Pearce and Malajczuk (1990a) tested the quality of the food base provided by two common hardwood hosts of A. luteobubalina by measuring rhizomorph production. 50 Inoculum and Infection They found that stem segments of sunbush were supe¬ rior to those of karri. Three genotypes of A. luteobubalina behaved in the same way. With so few experiments on substrate quality, data are insufficient to suggest a general superiority of one wood type over the other as a food base, but some Armillaria species or isolates may be favored by par¬ ticular species. However, observations similar to those made by Nechleba (1915) concerning the association of killing attacks with former hardwood sites continue to be made (Rishbeth 1982, Rykowski 1984). In the field, factors other than the intrinsic quality of the substrate may determine a stump's effectiveness as an inoculum source. Morrison (1972) and Rishbeth (1972b) both con¬ cluded that the frequently reported association of hard¬ wood food bases with disease could be partially attributed to those broadleaved trees in which resis¬ tance to infection is maintained by regrowth after cut¬ ting. They are less quickly exhausted as food bases than conifer stumps, which die rapidly. The generally higher wood density and greater resistance to decay of hardwood species compared to conifers (Rykowski 1984) may also increase the longevity of hardwood inocula. The possible "field" superiority of hardwood food bases as inoculum, at least for some Armillaria species, is not great, and the association of disease with hard¬ wood stumps should not be over-emphasised. As dis¬ cussed by Redfern (1975), it may be a mistake to assume that damage will diminish appreciably in suc¬ ceeding conifer rotations. This is supported by recent survey data from second-rotation radiata pine stands established on land cleared of indigenous hardwood forest (MacKenzie and Self 1988). It is salutory to quote Hartig, who wrote in 1874: "The disease often occurs especially destructively where the planting of conifers has been carried out after the felling of hardwoods .... But it should not be maintained from this that the rhizomorphs attack only from hardwood stumps to the conifer woods since, as we said earlier, the mycelium grows for several years on all conifer stumps and roots; therefore, hardwood stumps are not necessary for the spread or origin of the disease." The nature of the substrate probably has far less direct influence on disease development in plantations than the pathogenicity of the Armillaria species present in the previous crops. However, an indirect substrate ef¬ fect may occur through species selection resulting from host specialization. Thus, Rishbeth (1985a) found that despite being rare on broadleaved trees and stumps, A. ostoyae caused death as commonly as A. me/lea in coni¬ fers established on sites previously occupied by broadleaved woodland. Where conifers replaced coni¬ fers, it was the predominant cause of mortality. The importance of variation in pathogenicity between species is suggested in the early North American litera¬ ture. In a notable paper. Piper and Fletcher (1903) de¬ scribed damage in prune orchards by two forms of A. mellea (sensu lato). One form, referred to as A. mellea, caused severe damage and was believed to have been introduced. The other, referred to as A. mellea bulbosa, was much less damaging. The latter was abundant on native trees, both conifers and hardwoods. Later, Childs and Zeller (1929) observed disease in apple or¬ chards established on sites cleared of oak, but found no disease on sites formerly occupied by Douglas-fir. Both site types were infested with Armillaria, which the au¬ thors suggested might exist as two strains differing in "pathogenicity" (see chapter 6). Substrate Specialization In common with other wood-rotting fungi that kill tree roots, Armillaria is polyphagous. Individual species or isolates grow on excised stems or roots of many tree species, including ones which they would not encoun¬ ter naturally (Benjamin and Newhook 1984b, Rishbeth 1978a). There is little prima facie evidence for substrate specialization. In the field, however, substrates are ac¬ quired both parasitically and saprophytically. Where several Armillaria species of different pathogenicity and competitive saprophytic ability are present in the same forest type, substrates are unlikely to be equally avail¬ able to them all. Our knowledge of the better-known species clearly shows that their association with par¬ ticular substrates reflects their ecology rather than a substrate specialization or preference. Armillaria ostoyae is highly pathogenic and occurs mainly on conifers throughout Europe and North America (see chapters 6 and 8). However, its associa¬ tion with conifers is not exclusive. In Canada, Morrison and others (1985a) found that broadleaved trees within disease centers were frequently attacked and killed. Elsewhere in North America, A. ostoyae kills cherry (Proffer and others 1987) and several other hardwood species (Harrington and others 1989). By contrast, Europe's other major pathogenic species, A. mellea, may be described as a "hardwood species" because it has a wide host range among hardwood trees and shrubs, and is common on hardwood stumps (Guillaumin and others 1985, Rishbeth 1985a). The association is not ex¬ clusive, however, as it also attacks young or weakened conifers and occasionally occurs on conifer stumps (Davidson and Rishbeth 1988, Rishbeth 1985a). Armillaria gallica also has a wide host range (Guillaumin and others 1985) and has been recorded as a weak pathogen on both hardwood and coniferous hosts, but it is most important as a cause of butt rot in hardwood trees and as a colonist of hardwood rather than conifer stumps (Rishbeth 1985a). Morrison and Inoculum and Infection 51 others (1985a) found A. gallica exclusively on living and dead hardwood hosts. Experiments show that whereas only A. mellea and A. ostoyae infect vigorous English oak and Scots pine, re¬ spectively, all three species colonize both hosts when resistance is reduced by suppression (Davidson and Rishbeth 1988). Neither host specialization by the fun¬ gus nor selectivity by the tree are apparently main¬ tained under these circumstances. Thus, for A. ostoyae and A. mellea, their host specialization as primary para¬ sites largely determines their substrates as saprophytes. Kile and Watling (1983, 1988) have discussed the ecol¬ ogy of the five known Australian species (see chapter 8). Four of these species, A. luteobubalina, A. hinnulea, A. novae-zelandiae, and A. fumosa, have extended geo¬ graphical distributions which include Tasmania. Arm ill aria hinnulea and A. novae-zelandiae also occur in New Zealand. Some species overlap ecologically, but the last two species occur in wet forests, whereas A. luteobubalina predominates in dry sclerophyll eucalypt forests. Armillaria fumosa has only been found on wet sites within these dry forests, and is therefore associ¬ ated with the particular species of these locations. Armillaria luteobubalina is the only Australasian species for which comprehensive information about substrate species is available, but it does not indicate substrate specialization among the hosts commonly present in the dry sclerophyll eucalypt forest. Both stumps and trees of the major eucalypt species groups are equally susceptible to infection (Kellas and others 1987, Pearce and others 1986, Shearer and Tippett 1988); its host range includes 81 species in 21 plant families (see table 8.1). In New Zealand, A. limonea and A. novae-zelandiae cause serious disease in radiata pine established on sites for¬ merly occupied by indigenous forest comprising host species such as tawa and rimu (MacKenzie and Shaw 1977). However, no evidence indicates that certain spe¬ cies provide superior substrates or that they are pre¬ ferred substrates for one Armillaria species or the other (MacKenzie and Shaw 1977, van der Pas 1981a). In general, there is little evidence for substrate special¬ ization within the natural range of each Armillaria spe¬ cies. The New Zealand example provides a dramatic illustration since the two species involved appear to nsferred successfully from indigenous hard- :hern-hemisphere conifer (MacKenzie ' ertheless, in northern temperate for- species express a degree of specializa- id A. gallica are generally ’ ved hosts and A. ostoyae with The infection of stumps by basidiospores offers, in a sense, a "free choice" of substrate. Rishbeth (1988) made the interesting observation that A. ostoyae and A. gallica most frequently colonized conifer and hardwood stumps, respectively, although both species also colo¬ nized the other substrate. Longevity of Inoculum and Persistence of the Fungus Most estimates of inoculum longevity are based on ob¬ servations made on single occasions, and refer to the ages of stumps which show evidence of viable Armillaria. Observations of this nature offer no informa¬ tion on the persistence of the fungus on the site and may underestimate its longevity in individual stumps. For example, survival in the stumps of hardwood trees showing regrowth may be greatly affected by the ex¬ tended period over which such stumps become colo¬ nized. When the fungus is already present as a perthophyte or as a butt rot, colonization may begin long before the tree is felled. Thus, longevity of the fungus in individual roots may give little idea of the time over which the stump may act as an inoculum source. Estimates vary widely but generally indicate fungal survival for decades in both broadleaved and conifer¬ ous stumps. Pronos and Patton (1978) found that oaks killed by herbicide produced rhizomorphs for at least 14 years, and Rishbeth (1972b) reported that wood from English oak stumps could do so 40 years after the trees were cut. Swift (1972) gave a figure of at least 20 years for survival in East African hardwoods. The only data available for conifers are from ponderosa pine in North America, and probably refer to A. ostoyae. Shaw (1975) found that wood cut from 30-year-old stumps contained viable Armillaria which could produce rhizomorphs; Roth and others (1980) isolated the fun¬ gus from large, old-growth stumps more than 35 years old. They estimated that it would remain viable in such stumps for at least 50 years. Few data are available for identified species. Kile (1981) suggested a longevity of 15-25 years for A. luteobubalina in messmate stringyb- ark. In contrast, he isolated A. hinnulea from 70-year- old stumps of the same eucalypt species (Kile 1980b and pers. comm.). Rishbeth (1985a) recently reported an example in which A. gallica remained viable in an oak stump 53 years after felling. Making valid comparisons between species based on field observations is difficult since longevity is likely to be affected bv the stump species, its size, and by envi¬ ronmental factors. The difference in longevity between A. luteobubalina and A. hinnulea quoted above might be attributable largely to site differences since the observa¬ tions were made in different forest types (G.A. Kile, 52 Inoculum and Infection pers. comm.). There are some indications from experi¬ ments with small inocula about the effects on survival of soil moisture (Pearce and Malajczuk 1990a), tem¬ perature (Bliss 1946), and competing fungi such as Trichoderma viride Pers.: Fr. (Garrett 1957) but further work is required. Inoculum size may not be a major factor. Even in the comparatively minute inocula used in experiments, the fungus remained viable in Sitka spruce for at least 4 years (Singh 1980a) and in pine for up to 3 years (Patton and Riker 1959). Armillaria can persist on a site for a very long time. For example, Shaw and Roth (1976) suggest that individual clones of A. ostoyae may survive for several centuries. Clearly this must involve a succession of substrates. For pathogenic species, these may be acquired either at the margins of expanding disease centers or among re¬ generating trees within disease gaps following a period of survival in stumps. The figures cited for longevity in individual stumps suggest this period may be suffi¬ ciently long to permit a resurgence of disease. For weakly pathogenic species, persistence may be aided by the behavior of the extensive rhizomorph systems some of them form. In unmanaged forests, longevity probably confers a survival advantage on all species, but it may be par¬ ticularly important for the less pathogenic ones since the opportunity for them to acquire additional sub¬ strates may be more limited than for more pathogenic species. The latter may benefit, particularly in forests of susceptible species, by survival in disease gaps until a new crop becomes established. In forests which are managed intensively and are subject to selection cut¬ ting or regular thinning, longevity may no longer be a survival trait, at least for weakly pathogenic species, since a regular supply of stumps would be available for colonization. Factors Affecting Growth of Rhizomorphs from Inoculum The abundance, type, and distribution of rhizomorphs on a site are primarily determined by the Armillaria species present, but environment exerts a major influ¬ ence through the effects of soil. Variation Among Species Whereas all Armillaria species form rhizomorphs to some degree in axenic culture, not all have been ob¬ served to do so in the field. No rhizomorphs have been reported for A. tabescens (Rhoads 1956, Rishbeth 1982, Ross 1970) although Rishbeth observed them on in¬ ocula buried in soil. In A. luteobubalina, they are either absent (Kile 1981, Shearer and Tippett 1988) or sparse under natural conditions (Pearce and others 1986, Podger and others 1978); other Australasian species, for example A. limonea and A. novae-zelandiae (Hood and Sandberg 1987), form rhizomorphs readily. Armillaria hinnulea forms rhizomorphs more prolifically than A. luteobubalina, but they are confined to root surfaces (Kile 1980b, Kile and Watling 1983). Among European and North American species, rhizomorph production is greater in A. gallica and A. cepistipes than in A. ostoyae and A. mellea (Gregory 1985, Guillaumin and others 1989a, Redfern 1975, Rishbeth 1985a). Information is lacking for some of the more recently described species such as A. pallidula and A. fellea (Kile and Watling 1988), but A. sinapina is reported to produce rhizomorphs abundantly in the field (Berube and Dessureault 1988). At the present time, information is insufficient to establish that the ability to produce rhizomorphs represents a continuum among species, but that may well be the case. Morrison (1989) studied rhizomorph production by an array of species from Europe, Australasia, and North America using woody inocula buried in pots contain¬ ing a mixture of forest soil, peat, and sand. While pro¬ ducing valuable information, such studies are not necessarily a reliable guide to field behavior. Thus, A. luteobubalina produced rhizomorphs more abundantly (fig. 4.1) under these circumstances than might have been anticipated from the field observations reported above. Podger and others (1978) reported similar re¬ sults from pot culture, suggesting rhizomorph forma¬ tion may be inhibited in the field by environmental conditions. For other species, observations under artifi¬ cial conditions do coincide with field behavior (Gre¬ gory 1985; Redfern 1975; Rishbeth 1985a,b). The growth habit of rhizomorphs in soil also varies among species; branching (fig. 4.1) is either monopo- dial or dichotomous (Morrison 1982b, 1989). This char¬ acter may have ecological significance since Morrison (1989) found that species with dichotomously branched rhizomorphs tended to be more pathogenic than those producing monopodially branched rhizomorphs, but the distinction was not entirely consistent. The Effect of Soil on Rhizomorph Growth Most observations about soil have concerned its influ¬ ence on the incidence and severity of disease, whereas the primary interest here is effect of soil on the fungus itself. The wide variety of soils associated with disease (Ono 1965,1970, Rhoads 1956, Ritchie 1932, Shields and Hobbs 1979) suggests Armillaria species tolerate a fairly broad range of conditions. Field observations on effects of soil on disease fre¬ quently conflict. Unfortunately, many are of limited value, and may be misleading, because they refer to A. Inoculum and Infection 53 FIGURE 4.1—Variation in rhizomorph growth habit among Armillaria species. (Adapted from Morrison 1989). mellea (sensu Into) when more than one species may be present. In these circumstances, differences in disease incidence due to the differing pathogenicity of the spe¬ cies involved may have been incorrectly attributed to soil factors. Similar misinterpretations may also arise through failure to appreciate the effects on disease de¬ velopment of the discontinuous distribution of inocu¬ lum. Experiments in which woody inocula containing Armillaria isolates have been allowed to form rhizomorphs in soil (Gramss 1983; Morrison 1976; Redfern 1970,1973,1975; Rishbeth 1985b) confirm field observations that Armillaria can grow in a wide variety of forest and agricultural soils. Soil seems to exert a major influence on rhizomorph growth only under un¬ usual or extreme circumstances. Thus, pure sand can partially inhibit rhizomorph production (Garrett 1956b, Redfern 1973, Rykowski 1984), whereas peat stimulates growth and branching (Redfern 1973). Certain tropical soils inhibit rhizomorph development (Dade 1927, Fox 1964, Rishbeth 1980, Swift 1968), and the paucity of M7omorphs associated with damage by A. :balma may also be soil-induced (Pearce and 1 990a, Podger and others 1978). ■ hods have been used to assess from woody inocula in soil. These total length or dry weight of rhizomorphs and repeatedly measuring individual rhizomorphs. Rishbeth (1968) used the last method for testing the effect of temperature, and discussed some problems associated with this type of work. Moisture Working with A. mellea (sensu lato), Garrett (1956b) and Redfern (1970) found soil moisture had no effect on growth within the ranges 40%-80% and 25%-75% of moisture-holding capacity, respectively. Growth of A. luteobubalina also occurs over a wide range of matric potentials (-0.0008 MPa to -7 MPa), but it is restricted below -0.6 MPa (which is roughly equivalent to 25% moisture-holding capacity). Seasonal drying may partly explain the paucity of rhizomorphs of this spe¬ cies in Australian soils (Pearce and Malajczuk 1990a). In Britain, Morrison (1976) concluded that seasonal drying may affect growth of A. mellea (sensu lato) in the upper soil layers. Waterlogging may restrict growth at depth indirectly through the soil atmosphere (Rishbeth 1978a) and can prevent rhizomorph formation by inocula in pot ex¬ periments (Guillaumin and Leprince 1979). Despite the reservations already expressed about field observa¬ tions, it is notable that Armillaria has rarely been re¬ ported from permanently wet soils with an appreciable peat accumulation. There is a single observation of A. ostoyae from Scotland (senior author and S.C. Gregory pers. comm.), and Hintikka (1974) commented that it seems to be largely absent from forested Sphagnum swamps, except where the peat is thin and the ground water is moving. Temperature The in vitro studies reported in chapter 3 provide a guide to the behavior of the fungus in soil, but caution should be exercised in extrapolating results since im¬ portant differences may exist. For example, growth oc¬ curs at higher temperatures in agar culture than in soil (Rishbeth 1968). Using woody inocula colonized by a suspension of ba- sidiospores and by measuring growth directly, Rishbeth (1968) found the optimum temperature for rhizomorph growth through soil was about 22°C. Some growth occurred at 5°C and 28°C but none at 30°C. He concluded that rates of spread of about 1.5 m per year observed at sites in southern Britain, where the soil temperature at a depth of 15 cm averages 1(TC, roughly corresponded with those determined from his experiments. Eater, working with a number of isolates and species, Rishbeth (1978a) found the dry weight of rhizomorphs produced by inocula in soil was usually maximal at 20°C and minimal above 26°C and below 54 Inoculum ami Infection 10°C. He suggested the lack of rhizomorph growth in forest soils at low elevations in tropical Africa (Dade 1927, Fox 1964, Swift 1968) may be due to high soil temperatures. By contrast, low temperatures may be limiting in many forest soils, particularly in the north temperate zone during winter (Rishbeth 1978a). How¬ ever, the production of rhizomorphs from inocula in¬ volves two processes: initiation and growth. Initiation may occur over a more restricted temperature range than growth (Rishbeth 1968). Thus, pre-existing rhizomorphs may grow at lower temperatures than in¬ dicated by experiments in which rhizomorph produc¬ tion rather than growth is measured. Temperatures below 10°C may therefore be less restrictive than has hitherto been suggested. Although rhizomorph initia¬ tion may be curtailed in winter, growth of those initi¬ ated at higher summer temperatures may continue. The effect of low temperatures receives some support from in vitro studies (Hintikka 1974, Pearce and Malajczuk 1990a, Rishbeth 1968), but as indicated ear¬ lier, they may not provide an entirely satisfactory guide to behavior in soil and further work is required. Rishbeth (1978a) found variation in the effect of tem¬ perature on rhizomorph growth in soil among a world¬ wide selection of isolates, but there is little information for different species. Pearce and Malajczuk (1990a) tested growth of A. luteobubalina over a limited range of temperatures and found maximum growth at the high¬ est temperature tested (20°C) with virtually no growth at 10°C. On agar, the optimum temperature for growth by this species was in the range 20-26°C, suggesting that it might be somewhat higher in soil. Also on agar, the more northern or high-altitude European species A. borealis, A. cepistipes, and A. ostoyae have a lower opti¬ mum for growth than the southern or low-altitude spe¬ cies A. gallica and A. mellea (Guillaumin and others 1989a). Thus, although there is some evidence for inter¬ specific variation in the temperature relations of Armillaria, further work is required in soil. Temperature may affect both the number and branch¬ ing of rhizomorphs initiated from woody inocula. Redfern (1973) found that an isolate of A. mellea (sensu lato) initiated more rhizomorphs in soil at 25°C than at 15°C, and each system had a greater branching fre¬ quency at the higher temperature. This effect requires confirmation and further study with a range of species. The possibility that growth patterns may vary in re¬ sponse to seasonal variations in soil temperature is of particular interest and may have implications for infec¬ tion and spread. The studies on A. luteobubalina by Pearce and Malajczuk (1990a) demonstrated that rhizomorph be¬ havior may be influenced by an interaction between temperature and moisture. This may well apply to other species, although the relative importance of the two factors may differ elsewhere. pH No body of field observations suggests that pH has a significant effect on Armillaria. Gard (1928) associated disease in Persian walnut with a reduction in lime con¬ tent of the soil, and Rishbeth (1982) recorded killing by A. ostoyae on acidic soils but not at comparable sites where soil was alkaline. By contrast, he found A. mellea often killed trees on alkaline soils. In an inoculation ex¬ periment, Redfern (1978) found that infection by one isolate of Armillaria was significantly greater in an acidic soil than in an alkaline soil of similar sandy tex¬ ture. However, in all these cases any pH effect may have been expressed through the host rather than through the pathogen. Other authors (Kawada and oth¬ ers 1962, Rhoads 1956) refer to killing on acidic soils but this probably only reflects the pH of most forest soils. Experimental studies of pH effects are hampered by the difficulty of adjusting soil pH. In England, a succes¬ sion of workers partly avoided the problem by taking advantage of a natural pH variation induced in uni¬ formly sandy soil by differences in the depth of under¬ lying chalk. In an initial experiment, rhizomorph production by a single isolate was greater at pH 7.5 than at pH 4.9 (Redfern 1970). Subsequently, more comprehensive work (Morrison 1974) with a range of isolates gave a variable response, with some isolates being unaffected. Further work by Morrison (pers. comm.) has shown that these differences were related to species. Armillaria mellea and A. ostoyae grew more in acidic than in alkaline soil, whereas A. gallica was either unaffected by pH or favored by alkaline soil. Rishbeth (1985b) tested three species in the same soils but de¬ tected no differences. Inhibitory Substances After several experiments with sterilized soil extracts. Swift (1968) attributed the absence of rhizomorphs from forest soils in Rhodesia (Zimbabwe) to a water- soluble inhibitor. Olembo (1972) found unsterile leachates of East African soils reduced the colonization of wood by Armillaria, but no further work has been done on this topic. Organic Matter and Soil Nutrient Status Accumulating evidence suggests soil nutrition affects rhizomorph growth. Rykowski (1984) confirmed the stimulating effect of peat (Redfern 1973) and observed a similar response to pine bark compost. Studying the Inoculum and Infection 55 influence of various organic soil amendments on rhizomorph development, including peat, Guillaumin and Leprince (1979) obtained rather different results but nevertheless concluded that the surrounding me¬ dium affects growth. Morrison (1975) investigated the peat effect and demonstrated that rhizomorph growing tips absorb nutrients. He suggested that the nutrients available from a food base may be supplemented by uptake from soil, and that rhizomorph development may be related to soil nutrient status. Nutrient balance may also be important. Rykowski (1984) found growth in one soil was increased by application of potassium and reduced by nitrogen and phosphorus. The Distribution of Rhizomorphs in Soil Soil moisture affects the vertical distribution of rhizomorphs in soil. Morrison (1976) found rhizomorphs grow towards the soil surface, and has suggested this behavior is a response to the oxygen gradient in soil. Vertical distribution is probably con¬ trolled by seasonal desiccation of the upper soil layers and by oxygen and carbon dioxide concentrations lower down (Morrison 1976, Rishbeth 1978a). Hartig (1873b) noted rhizomorphs lie at about 10 cm depth, and Lawrence (1910) observed them in "great abun¬ dance from 3 to 18 inches below the soil surface." Later authors reported a concentration in the upper soil lay¬ ers, generally within 10-20 cm of the surface (Day 1927b, Ono 1970, Redfern 1973). Where a humus layer is present, rhizomorphs are more common there than in the mineral soil below (Hintikka 1974, Singh 1981b), an interesting observation in view of the stimulating ef¬ fect of peat on rhizomorph growth. The concentration of rhizomorphs in the upper soil layers may be impor¬ tant epidemiologically because of the greater vulner¬ ability of trees to infections initiated on the root collar and proximal part of the root system compared to the deeper, more peripheral roots (Bliss 1946, Hintikka 1974, Patton and Riker 1959, Shaw 1980). Inoculum po¬ tential may also be greater than with a less stratified distribution. These field data on rhizomorph distribution are most likely to have been contributed by species which form rhizomorphs capable of extensive growth through soil. Little information is available for species with rhizomorphs which are more closely associated with roots. Pearce and others (1986) found rhizomorphs of A. luteobubalina were present on infested sites at depths between 5 and 15 cm. Experimentally, A. luteobubalina laced rhizomorphs from woody inocula buried at Y.irce and Malajczuk 1990a) although the num- tal length were small. For species which ■ by root contact, it seems likely that the nfection and spread would be maxi¬ mized by an ability to form rhizomorphs throughout the rooting depth of the host. The horizontal distribution of rhizomorphs can be exten¬ sive. Armillaria gallica forms a network of rhizomorphs over the surface of living roots (Rishbeth 1985a). Redfern (1973), who probably observed the same species, sug¬ gested that rhizomorphs branch and anastomose to form extensive, complex networks which envelop both living trees and the food bases from which they originated. In one new plantation, Redfern (1973) estimated that rhizomorphs had spread up to 35 m in 37 years from ad¬ jacent, long-standing woodland infested by Armillaria. Armillaria cepistipes and other prolific rhizomorph-form- ing species may behave in the same way. In North America, Lawrence (1910) observed that rhizomorphs growing from infected raspberry canes formed a "net¬ work by frequently branching and rebranching", and Childs and Zeller (1929) referred to "a complete network of rhizomorphs about the larger roots" of orchard trees on fir-cleared land infested by a non-pathogenic species. Several authors have estimated the abundance of rhizomorphs in soil (Hintikka 1974; Hood and Sandberg 1989; Ono 1965,1970; Rykowski 1984). Hintikka re¬ corded 121 cm of rhizomorphs per 100 cm 2 of soil sur¬ face. Inoculum Potential and Infection Rhizomorphs represent extensions of inoculum, and are important in the infection, spread, and persistence of many Armillaria species. In a minority of species, rhizomorphs are absent or are only sparsely formed, and in these species infection is confined to points of contact between host roots and the inoculum. The infection pro¬ cess may involve epiphytic rhizomorphs or the transfer of mycelium, but the most important feature for the epi¬ demiology of these species is the need for contact be¬ tween host and inoculum. Infection also occurs in this way under certain environmental conditions which pre¬ vent or restrict rhizomorph formation. Garrett (1970) concluded from his experiments with rhizomorph-form- ing Armillaria species that inoculum size, distance be¬ tween the inoculum and the host, and the influence of environment on the fungus were the major determinants of inoculum potential. However, where infection occurs at root contacts, only the first and last factors seem likely to be important. This section primarily addresses those factors which af¬ fect disease development through their effect on inocu¬ lum potential. Chapter 5 describes the infection process in detail; here, attention is confined to the way in which infection occurs and its effect on the epidemiology of disease. The role of wounds in the successful establish¬ ment of infection is also considered. 56 Inoculum and Infection Inoculum Potential Little work has been done on inoculum potential since Garrett's classical experiments (Garrett 1956b, 1958) and none with species lacking rhizomorphs. Garrett ex¬ perimented with model systems consisting of small woody inocula and potato tubers in soil. He found rhizomorph growth rate was related to inoculum size, and that the extent of infection in potato tubers in¬ creased with inoculum size and decreased with in¬ creasing distance between inoculum and tuber. Rhizomorph growth rate also declined with time, and he attributed this partly to nutrient depletion in the in¬ oculum and partly to competition for nutrients be¬ tween the main apex of a rhizomorph system and its subordinate branch apices. Rykowski (1984) recently confirmed Garrett's results, using larger inocula and Scots pine seedlings. He used indices derived from the number, length, and weight of rhizomorph systems produced from inocula, and the number of apices on those systems, to calculate the "potential infection threat" presented by inocula in various soils. Although the concept of inoculum potential is simple and of considerable biological importance, it is difficult to envisage its application to individual trees since field situations are frequently complex. Inocula are rarely discrete, and infection often is not readily associated with specific point sources. Also, inocula vary in size from parts of individual roots to entire stumps. The rhizomorph networks formed by some species may present an additional complication. The behavior of these systems requires study. Redfern (1973) sug¬ gested they may be relatively long-lived, being sup¬ ported by a succession of food bases as they become available to different parts of the network, and the di¬ rection of nutrient flows changing to maintain the en¬ tire system from different sources. This is apparently inconsistent with experiments on translocation (Ander¬ son and Ullrich 1982b, Schiitte 1956) which have shown that it only occurs towards growing tips. Morrison (1975) found that nutrients absorbed by growing tips were not translocated towards the food base. These ex¬ periments do not represent the behavior of an entire network, however, and they are not inconsistent with the possibility that the direction of translocation within a rhizomorph in a network may vary with time. Ander¬ son and Ullrich (1982b) commented that "if the (rhizomorph) base were converted to a sink for nutri¬ ents, as may be the case during fruiting or exhaustion of food reserves, rhizomorphs may transport nutrients from tip to base." This is supported by observations on severed rhizomorphs. Rhizomorphs forming part of a network and which are severed in situ initiate new rhizomorphs simultaneously from the cut ends, as do excised sections of large diameter rhizomorphs (Hintikka 1974, Redfern 1973, Rykowski 1984). The principle of a fungal corpus consisting of a network of colonized stumps and rhizomorphs may apply equally well to species such as A. mellea, A. ostoyae, and A. hinnulea with more restricted rhizomorph-forming abilities. Roots may simply predominate over rhizo¬ morphs in the network. However, some evidence indi¬ cates that, in contrast to A. gallica and A. cepistipes, rhizomorphs of A. mellea are short-lived and are pro¬ duced in successive waves (Guillaumin and others 1989a) which suggest these species are unlikely to form persistent networks. Clearly, much of the foregoing is speculative, but it is worth consideration since rhizomorph systems which behave in this way might create inocula consisting ef¬ fectively of several stumps. Despite this complexity, and notwithstanding the minute inocula used by Garrett (1956b) and Rykowski (1984) compared to substrates available naturally, there seems no reason to doubt the general applicability of the principle of inoculum potential to such large in¬ ocula. Inoculum potential is maximized where healthy roots and inoculum are in contact; where gaps are bridged by rhizomorphs, it diminishes with increasing distance between them. However, few detailed analy¬ ses of disease patterns in relation to the distribution of inoculum have been done. Understanding such pat¬ terns requires considerable knowledge of pathogen behavior in the circumstances of each outbreak, par¬ ticularly the relative importance of rhizomorphs and root contacts as the means of spread in the species involved. Shaw (1980) and Shaw and others (1976a) described a relatively straightforward situation in young pon- derosa pine involving a single species, A. ostoyae (Shaw 1984), spreading essentially by root contact from dis¬ crete sources of inoculum. On the other hand, disease development following replacement of indigenous for¬ est comprising many host species and more than one Armillaria species by a susceptible monoculture (MacKenzie and Shaw 1977) is much more complex. Under these circumstances, the pattern of mortality as¬ sessed on a single occasion (van der Pas 1981a) may be difficult to interpret (Roth and others 1979). MacKenzie and Shaw (1977) recorded decreasing mortality with increasing distance from infected stumps. Though such a pattern suggests the stumps were acting as the initial infection sources, interpretation of subsequent events in terms of inoculum potential is not possible. As sug¬ gested by Roth and others (1979), the effect could be caused by rapid, early killing within the rooting zone Inoculum and Infection 57 of the stumps acting as inocula, followed by a slower rate of mortality outside this zone as growing roots or rhizomorphs bridge the gaps between potential hosts and inocula. In another example, similar patterns of mortality among saplings around eucalypt stumps infected by A. luteobubalina (Pearce and others 1986) may simply have reflected the time when the developing sapling root systems made contact with stump roots. Alternatively, disease centers in young conifers (probably caused by A. ostoyae) show reduced extension rates because of in¬ creasing host resistance (Redfern 1978) rather than de¬ clining inoculum potential with increasing distance from a point source. On a large scale, the effect of an increase in inoculum can be appreciated readily. Forestry operations such as clear-felling, selective cutting, thinning, the treatment of indigenous crops with herbicides, or events such as fire provide opportunities for a massive increase in in¬ oculum (Kile 1980b, Pronos and Patton 1978, Rishbeth 1972b, Shaw and others 1976a, Swift 1972, van der Pas 1981b). For pathogenic species, more inoculum typi¬ cally results in more disease. Thus, A. luteobubalina causes disease in unlogged eucalypt forest (Kile 1983b), but the greatest incidence and severity of disease is as¬ sociated with logging (Kellas and others 1987, Pearce and others 1986). Concomitantly, natural regeneration or planting repositions hosts relative to the inoculum. Thus, in former disease centers which are devoid of hosts, or in plantation systems where trees are planted beside stumps which may subsequently become colo¬ nized, the distance between potential hosts and inocu¬ lum may be small. Physical disturbance of the soil by logging, plowing, scarifying, or even planting may sever rhizomorph net¬ works, which respond by initiating new growing tips from the cut ends. Besides increasing the amount of in¬ oculum and perhaps the availability of new and more susceptible hosts, harvesting disturbance can also stimulate the production of rhizomorph growing tips and locally increase the chance of infection (Redfern 1973, Rykowski 1984, see chapter 11). Inorganic fertilizers may influence inoculum potential through the soil environment. The effect of some macro-nutrients on rhizomorph production by inocula in soil has already been mentioned (Rykowski 1984). The inoculum may also be directly affected. Work by io (1970-71), Garrett (1953,1970), and Rykowski "t. igests the possibility that crop fertilization so inoculum potential by changing sub- hen roots with an enhanced nutrient . oecome inoculum. Both of these inter¬ esting possibilities merit further study. To assess the need for control in Armillaria-mfested ar¬ eas (see chapter 11), forest managers must estimate the inoculum potential of the species present in addition to knowing their pathogenicity and distribution. How¬ ever, even in the simplest situation involving only one species, there seems little possibility that the inoculum potential of Armillaria on a site could be assessed by ca¬ sual observation. For example, although it has been used for modeling purposes (see chapter 10), stump size may be a poor guide unless colonization is com¬ plete. The circumstances under which complete coloni¬ zation may be achieved include the invasion of living, susceptible conifers by highly pathogenic species, the colonization of freshly felled conifers by growth of the same species from root lesions, and the colonization of healthy conifer stumps by species capable of forming extensive rhizomorph systems. In hardwoods, how¬ ever, colonization may be restricted in those species which tend to regrow after cutting (Rishbeth 1972b). In some eucalypts (Kile 1980b), and possibly oaks, the heartwood is resistant to decay and remains uncolonized. Pearce and others (1986) found a signifi¬ cant relationship for A. luteobubalina between an esti¬ mate of how much inoculum was provided by individual, infected stumps and mortality in nearby saplings; assessments like this are unlikely to be fea¬ sible in commercial forestry, however. A reduction in inoculum potential or the prevention of inoculum buildup provides the basis for many control measures (see chapter 11). Under natural conditions, the amount of Armillaria inoculum on a site may be re¬ duced by competition from other fungi and by fire. In the case of wood-rotting fungi which are also parasites, such as Phellinus weirii (Morrison and others 1988) and Heterobasidion annosum (Greig 1962), competition is not beneficial; but some saprophytic decay fungi are also able to compete successfully and may be useful for bio¬ logical control (Pearce and Malajczuk 1990b, Rishbeth 1976). The soil-inhabiting fungus Trichoderma viride may exert a degree of control which can be enhanced by soil fumigation (Bliss 1951; Garrett 1957,1958; Ohr and others 1973). Fire may kill rhizomorphs in soil (Hood and Sandberg 1989), but its effects on inoculum survival and subsequent rhizomorph activity are unknown. A massive inoculum is not a prerequisite for infection if the distance between inoculum and host is minimal. Many experiments have demonstrated that successful infections can be established on small trees by means of small woody inocula, some weighing as little as a few grams (Patton and Riker 1959, Rykowski 1984). This has particular relevance for control by inoculum re¬ moval since root fragments inevitably remain after destumping and root raking operations (Morrison and others 1988). Although a high level of control can be 58 Inoculum and Infection achieved by destumping, certainly a level which would return an infested site to normal productivity, residual root fragments may nevertheless permit the re-estab¬ lishment of disease. Damage may be confined to a few early losses, but it could be extended by secondary, tree-to-tree spread (Rykowski 1984). Although small and large inocula may both cause in¬ fection, Rykowski (1984) has suggested that each repre¬ sents a different type of threat. In experiments with small, woody inocula, rhizomorph production per unit volume of inoculum was inversely related to total in¬ oculum volume. This suggests that rhizomorph pro¬ duction is delayed until the substrate has been fully colonized and certain nutritional requirements have been met (Benjamin and Newhook 1984b, Garrett 1953, Patton and Riker 1959, Rykowski 1984). Rykowski con¬ cluded that whereas in small substrates the phases of colonization, rhizomorph production, and exhaustion are accomplished rapidly, the same process takes longer in larger inocula. He argued that stumps may behave in the same way, presenting short-term and long-term infection threats, respectively. Infection As already discussed, rhizomorphs are formed in soil to a greater or lesser extent by most Armillaria species; the absence of rhizomorphs is apparently uncommon among species in the genus. In some species, they are restricted to root surfaces or to the close proximity of roots, whereas others form abundant rhizomorphs which ramify freely through soil. Without rhizo¬ morphs, infection is confined to points of contact between host roots and the inoculum; with increasing rhizomorph production, infection can also take place at greater distances from the inoculum. Because rhizomorphs are often abundant, much of the early literature from temperate countries emphasized the importance of rhizomorphs growing freely through soil as a means of spread. Indeed, some authors consid¬ ered them essential (van Vloten 1936). However, a number of authors either observed infection at root contacts (Kawada and others 1962, Prihoda 1957, Zeller 1926) or inferred its occurrence from their observations (Marsh 1952, Molin and Rennerfelt 1959). Working in black currant plantations, Marsh (1952) found the pat¬ tern of disease spread was best explained by root con¬ tact infection rather than by rhizomorphs growing in soil unoccupied by roots. Molin and Rennerfelt (1959) concluded that spread occurs mainly by root contact, and rhizomorphs only play a secondary role except over distances less than 1 m. In Czechoslovakia, Prihoda (1957) referred specifically to infection of Nor¬ way spruce both by rhizomorphs and by the transfer of mycelium at root contacts where rhizomorphs were ab¬ sent. He commented that although soil rhizomorphs were present on one site, they were sparse and weak and the bulk of infection was by mycelium transfer. He discussed the possibility that rhizomorph formation might be inhibited by alkaline soils, but he concluded that soil was unimportant and that some "forms" of Armillaria do not produce rhizomorphs whereas others do so abundantly. Without our present understanding of Armillaria spe- ciation and ecology, earlier authors did not appreciate the difference between spread of the more pathogenic species among susceptible hosts and the spread of less pathogenic species on stumps and weakened trees. Prihoda's comments (1957) were therefore particularly percipient. These European observations of spread by root contact probably referred to either A. ostoyae or A. mellea, which are pathogenic and form fewer rhizo¬ morphs than the weakly pathogenic species A. gallica and A. cepistipes (Guillaumin and others 1985,1989a; Rishbeth 1985a). Abundant rhizomorph production by the latter species may also prompt misinterpretation where they occur with pathogenic species if it is as¬ sumed that any rhizomorphs observed in soil are those of the disease-causing species. For species such as A. tabescens, A. hinnulea, and A. luteobubalina in which rhizomorphs are either absent or confined to root surfaces, infected roots must be in con¬ tact with potential hosts, or very close to them, for in¬ fection to occur (Kile 1980b, 1981; Kile and Watling 1983; Pearce and others 1986; Shearer and Tippett 1988). Nevertheless, interlocking root systems can pro¬ vide highly effective pathways for spread by patho¬ genic species among susceptible hosts. Surveying dieback in messmate stringybark and mountain ash as¬ sociated with A. hinnulea (Kile 1980b, Kile and Watling 1983), Kile (1980b) found that 74% of living trees had infections or epiphytic rhizomorphs on the root system. By contrast, species which form extensive rhizomorph systems, such as A. gallica and A. cepistipes, are not re¬ stricted in this way. Armillaria mellea and A. ostoyae have a lesser ability to form rhizomorphs in soil than A. gallica, but they are not confined to root surfaces and these species may occupy an intermediate position. In ponderosa pines, A. ostoyae spread between roots near to each other as well as at contacts (Shaw 1980). In New Zealand, free-growing rhizomorphs are com¬ mon in soil where both A. limonea and A. novae- zelandiae are present (Hood and Sandberg 1987), but the relative contribution of each species to the rhizomorph population is unknown. However, both species readily produce rhizomorphs in pot culture (Benjamin and Newhook 1984b), so it is likely that rhizomorph spread is important in both cases. Inoculum and Infection 59 Recent work has suggested a relationship between rhizomorph production and pathogenicity among some European species. The more pathogenic species tend to produce smaller rhizomorph systems than less pathogenic species (Gregory 1985; Guillaumin and oth¬ ers 1985, 1989a; Redfern 1975; Rishbeth 1985a). Some evidence indicates the relationship may also apply to North American and Australasian species (Morrison 1989). Further information, particularly about rhizomorph production, is required for many of the more recently described species, but differences appear to be large (Gregory 1985, Redfern 1975) and may have considerable ecological significance. For a weak patho¬ gen, a strategy involving a wide dispersion of inocu¬ lum offers the advantage of position when potential food bases become available. Thus, weakly pathogenic species which form extensive rhizomorph systems and infest roots in a network of rhizomorphs are able to ex¬ ploit this advantage in the acquisition of substrates, which may consist of stumps or living trees with de¬ clining resistance. More pathogenic species, by con¬ trast, do not require such a strategy and are able to spread among susceptible hosts through root contacts. It follows from this discussion that spread in patho¬ genic species is likely to be influenced more by factors affecting the distribution of tree roots than by those which affect rhizomorph development. Thus, for pur¬ poses of disease management, pathogenic species in North America and Europe should perhaps be consid¬ ered to have a greater affinity with Phellinns zveirii or even Heterobasidion annosum than they traditionally have been. Before our present understanding of speciation and pathogenicity in the genus, considerable debate fo¬ cused on the environmental conditions required for in¬ fection and on the need for infection courts provided by root wounds or debilitated roots. The distinction is important since otherwise healthy roots which are physically wounded, perhaps by abrasion against stones, by animals, or by harvesting machinery, differ greatly from roots debilitated by, for example, poor soil aeration. From the many inoculations which have been done on wounded roots, little doubt remains that infection can take place through wounds; but their importance as natural infection courts, however, has not been estab¬ lished clearly. Dimitri (1969) concluded that although i ilection in Norway spruce can take place through undamaged roots, it occurs primarily through 1 dead roots. Buckland (1953) reported that to detect infection through healthy bark ugl s-fir, observing it only in roots mechanically damaged or physiologi- ; kka (1974) believed root collar in¬ juries caused by snow bend promoted rhizomorph penetration at this point. However, it is difficult to de¬ termine by observation alone the role of wounded or stressed roots in the establishment of infection. In one of the few inoculation experiments designed to test the effect of wounding. Weaver (1974) found that it in¬ creased the number of isolates of A. tabescens which were able to infect peach roots. Invasion was also more extensive in injured roots. More recently, Whitney and others (1989b) found wounding increased infection in balsam fir inoculated with A. ostoyae. Evidence from natural disease outbreaks, and the ease with which unwounded trees can be infected in inocu¬ lation experiments, suggest that, at least for the more pathogenic species, wounds are unlikely to increase the success of infection. Wounds and debilitated roots could be important infection courts for less pathogenic species such as A. gallica, but no evidence supports this. Gregory (1985) showed that the length over which rhizomorphs became attached to the host surface was greater for species of low pathogenicity than for those of high pathogenicity. This could be expected to pro¬ vide weakly pathogenic species with a greater opportu¬ nity to encounter wounds than would be available to pathogenic species. Conclusions Wood, mainly tree roots, provides the major source of inoculum for Armillaria. Many older observations of disease supported the view that hardwoods provide a superior substrate for Armillaria than conifers. In gen¬ eral, little experimental evidence substantiates an in¬ trinsic difference between the two substrates but stumps of broadleaved trees may exhibit greater lon¬ gevity as inoculum. Some Armillaria species may sub¬ sist better on particular food base species, but there is no evidence for substrate specialization. However, a degree of ecological specialization is known for some north-temperate species. All species form rhizomorphs in culture, and almost all do so in forest soils, but they vary greatly in the amount of rhizomorph growth. Some species are epi¬ phytic or restricted to the close proximity of roots, whereas others grow freely through soil, forming net¬ works which link both colonized stumps and living trees. Infection is probably caused by rhizomorphs in most species, either at contacts between host roots and the inoculum or at some distance from the inoculum. For species lacking rhizomorphs, or where soil condi¬ tions prevent their formation, infection is restricted to contacts and occurs by the transfer of mycelium. Spe¬ cies with epiphytic rhizomorphs are similarly re¬ stricted, but infection can be either by mycelium transfer or by rhizomorphs. The relative importance of 60 Inoculum and Infection the two methods for these species is unknown. No in¬ formation is available about the influence of soil on in¬ fection by mycelium transfer. The environment can have a major effect on Armillaria through the effects of competing fungi on survival in woody substrates and through the influence of soil on rhizomorph growth. The fungus can grow in soils de¬ rived from a wide variety of lithologies, but more fer¬ tile soils may be particularly favorable since nutrient uptake from the soil may supplement nutrients from the food base. Soil moisture, temperature, and pH all affect rhizomorph growth, and there is some evidence for an interaction between moisture and temperature which may also be important. Species differ in their re¬ sponse to temperature and pH, but little information is available. The inoculum potential of Armillaria is influenced by the amount of inoculum, by the distance between the inoculum and the host, and by environmental effects. Forestry operations such as felling and thinning increase inoculum on a site, but patterns of mortality should not be interpreted simply in terms of inoculum potential. Interaction between, among other things, the amount and distribution of inoculum, the method of spread by the Armillaria species involved, and root system development by the host may be equally important. The more pathogenic Armillaria species may produce smaller rhizomorph systems than less pathogenic spe¬ cies. Further information is required, particularly for more recently described species, but such a tendency may have considerable ecological significance. Thus, the extensive rhizomorph systems produced by weakly pathogenic species may represent a strategy for the wide dispersal of inoculum in order to gain the advan¬ tage of position when potential substrates become available. By contrast, the interlocking root systems of susceptible hosts may provide an effective means of spread for more pathogenic species, and obviate the need for extensive rhizomorph systems. Wounds may be important infection courts for weakly pathogenic species, but they are unlikely to increase the success of infection by more pathogenic species. Inoculum and Infection 61 CHAPTER 5 Infection, Disease Development, Diagnosis, and Detection Duncan J. Morrison , Ralph E. Williams , and Roy D. Whitney T he first comprehensive description of Armillaria root disease, including the causal fungus and its life cycle, was made by Rob¬ ert Hartig (1874). He recognized that Rhizo¬ morpha fragilis Roth with its two chief forms, R. subter- ranea and R. subcorticalis, composed part of the mycelial body of Agaricus (Armillaria) melleus. Rhizomorpha subterranea and R. subcorticalis were the binomials ap¬ plied to the cylindrical brown to black mycelial strands found in soil and on root surfaces and the flattened white to cream colored mycelial felts (fans) found between the bark and wood of hosts, respectively. Hartig observed the basidiomes of A. melleus develop¬ ing on rootstocks with R. subcorticalis under the bark and on rhizomorph apices in soil. He also described infection and disease development in several conifer species. Since Hartig's work, more than 600 species of woody plants have been recorded as hosts of Armillaria species (Raabe 1962a). The infection process and disease devel¬ opment have been described for several hardwood and coniferous hosts. A wide variety of symptoms, signs, and host responses resulting from disease have been recorded, reflecting the wide host and geographical ranges and number of Armillaria species. This chapter describes the infection process and disease develop¬ ment in photosynthesizing (green) plants, the symp¬ toms and signs on diseased plants, and how these symptoms can be used to detect Armillaria root disease in forests and orchards. The Infection Process and Disease Development The Infection Process (1934) defined infection by Armillaria root dis- •tration of the fungus into the host, with or : uent colonization. The roots of woody ted following contact between a 'hizomorph or diseased root. Al¬ though many records document many different hosts being attacked (Raabe 1962a), the literature on the in¬ fection process is sparse. All detailed studies of the in¬ fection process predate acceptance by pathologists that Armillaria mellea sensu lato represents many species. Di¬ verse host responses and sometimes conflicting reports about the same host are evident in accounts of disease occurrence. These apparent discrepancies may be partly attributable to different Armillaria species having been involved. Current knowledge of the geographical distribution and host preferences of Armillaria species helps clarify the identity of Armillaria species reported in early studies. For example, the Armillaria on pine (Hartig 1874) is probably A. ostoyae (H. Marxmiiller pers. comm.) and Thomas' (1934) studies on hardwood trees probably involved A. mellea sensu stricta. The first account of the Armillaria infection process was given by Hartig (1874). He wrote, “The killing of roots is brought about by Rhizomorpha fragilis which bores into the root, spreads out in all directions as R. sub¬ corticalis and thus from the point of attack continually approaches the root stock until this is reached. General agreement exists among the detailed studies of coniferous (Day 1927b, Rykowski 1975, Woeste 1956) and hardwood hosts (Guillaumin and Rykowski 1980, Thomas 1934) about the infection process by rhizomor- phs. A rhizomorph becomes attached to a root initially by hardening of the mucilagenous substance which covers its growing tip. Then, single hyphae developing from the rhizomorph tip and penetrating the outer lay¬ er of cork cells anchor the rhizomorph to the root. On suscepts with smooth bark, branches which will form the root-penetrating rhizomorph develop at points of firm contact with the root surface. The branches origi¬ nate in the inner cortical cells of the rhizomorph when hyphae divide and spread laterally. These hyphae force their way through the outer cortical cells of the rhizo¬ morph and emerge as a branch. Branches may be nu¬ merous and always develop on the side of the rhizomorph contacting the host (Thomas 1934). 62 Diagnosis and Detection Thomas (1934) studied how Armillaria infected fleshy and woody roots of susceptible and resistant hosts. Penetration of the rhizomorph was essentially the same for both groups. The lateral branch, acting as a unit, not as individual hyphae, begins to penetrate by mechani¬ cal force. The host cork cells under the rhizomorph are pushed in and slightly compressed (figs. 5.1A,B). At this stage, tissues below the cork cells appear disorga¬ nized, which is attributed to secretions from the rhizomorph. Penetration continues by a combination of chemical and mechanical means. Beneath the cork, the rhizomorph branches spread laterally and radially into bark tissues. The descriptions of this process by Day (1927b) and Woeste (1956) indicate that more chemical destruction of tissues occurs in conifers than in hard¬ woods. Enzymatic breakdown of suberin may also be involved in bark penetration (Swift 1965, Zimmermann and Seemiiller 1984). In suscepts with scaly bark, the rhizomorphs (R. subterranea) run tangentially under bark scales becom¬ ing R. fragilis (Woeste 1956); that is, white strands with¬ out a rind. Rhizomorphs may emerge successively from beneath bark scales along a root. Rhizomorpha fragilis (as R. subcorticalis) may penetrate the bark scales and develop infection wedges beneath each one. Cell walls turn brown and cell contents become disorga¬ nized some distance from the infection wedge. Day (1927b), Thomas (1934), and Woeste (1956) con¬ cluded that rhizomorphs of Armillaria need neither wounds nor anatomical points of weakness to attack healthy, vigorously growing suscepts. However, root injuries caused by stones and wind-induced root move¬ ments, wounds made by insects and scarification equipment, and rootlets killed by excessive moisture could all serve as infection courts (Basham 1988, Dimitri 1969, Kile 1981, Rizzo and Harrington 1988b, Whitney 1961). Two years after inoculation with A. tabescens, most isolates had infected injured roots of peach, whereas only a few isolates had infected uninjured ones, and invasion of injured roots was usu¬ ally more extensive (Weaver 1974). Zeller (1926) described infection of suscept roots by mycelial transfer across points of contact with diseased apple roots. He suggested that infection of the suscept root begins when its healthy bark is acted upon by toxic substances produced by Armillaria in the contact¬ ing diseased root. Shallow brown spots appear in the bark's outer parenchyma, and these eventually coa¬ lesce. Flakes of dead cork are sloughed as new cork lay¬ ers are formed. Armillaria mycelium was not found in the spots until two or more plates of cork had been sloughed. Eventually, the fungus reaches the cambium and a canker develops. Conifers may become infected in a similar manner (Morrison unpubl.). Initially, myce¬ lial fans of A. ostoyae grow in a root's outer bark. As the area of colonized bark increases, mycelial fans pen¬ etrate to the cambium. Bark tissue becomes necrotic in advance of the mycelial fans. Host Response to Infection Host responses to Armillaria root disease fall into three categories: exudate production, meristematic activity, and biochemical interaction. At the biochemical level, fungal infection involves an interaction between com¬ pounds already present in the host or induced by infec¬ tion and extracellular fungal metabolites. These biochemical interactions are discussed in chapter 3. Here, responses involving meristematic activity and exudates are discussed. Meristematic activity leading to cork and callus forma¬ tion and, frequently, adventitious roots is a common B FIGURE 5.1 — Penetration of the bark of a walnut root by Armillaria mellea. A: Subterranean rhizomorph with a develop¬ ing lateral branch; B: Infection wedge penetrating host bark (1- rhizomorph; 2- rhizomorph branch; 3- infection wedge; 4- host bark). (J.J. Guillaumin) Diagnosis and Detection 63 host response to Armillaria infection on roots and at the root collar. Most descriptions of the infection process by rhizomorphs indicate that all living, vigorous suscepts responded to bark penetration by forming one or more secondary cork layers beneath the point of penetration. Thomas (1934) noted that in resistant hosts the lesion produced by initial penetration was walled off by the secondary periderm; this cork layer then widened with root growth. In susceptible hosts, pen¬ etrating rhizomorphs breach these secondary cork lay¬ ers. Rykowski (1975) observed similar reactions in Scots pine roots. On some roots, the penetrating rhizomorph reached the cambium whereas on others secondary cork isolated the infecting mycelium from living host tissues and caused infected bark to be sloughed (fig. 5.2). Observations on plum rootstocks showed that their resistance to A. mellea was mainly due to post-infection reactions, because the success rate in penetration by the fungus was similar for susceptible and resistant rootstocks (Guillaumin and others 1989b). Mycelial fans in the bark and sapwood grew consider¬ ably less in resistant rootstocks, and the slower growth was associated with pink or purple discoloration of bark and wood tissues surrounding lesions (fig. 5.3). FIGURE 5.2 —Armillaria ostoyae lesion on a Douglas-fir root in which secondary bark has isolated the infecting mycelium (1-xylem; 2-bark; 3-infected bark). (D.J. Morrison) ia mellea lesion on a resistant Prunus sp. . oloration of bark around the lesion. Perhaps this post-infection reaction, which only occurs in living tissues, kills the mycelium thus preventing disease development. Many hosts respond to Armillaria attack by exuding resin, gum, or kino. In hardwood hosts in which a pathogenic Armillaria species had penetrated to the cambium, Thomas (1934) observed that the xylem be¬ came brown ahead of penetrating hyphae. This reac¬ tion appeared to result from a gummy deposit in the vessels, perhaps secreted from the affected cells or a product from their walls. Resin production in pines was confined to areas of bark where mycelium had penetrated to the cambium and was not associated with ectotrophic spread in bark scales (Redfern 1978). On 5-year-old Corsican pine, the resin response was much more vigorous on trees inoculated with A. mellea than with A. ostoyae (Rishbeth 1982). Copious resin formed beneath bark tissue infected by A. mellea, forc¬ ing the tissue away from the root wood. Some mycelial sheets of A. mellea containing the resin were not viable, while those of A. ostoyae appeared to be unaffected (Rishbeth 1982). The effects of conifer resin or resin components on Armillaria growth in culture vary. Pinene inhibited growth of A. ostoyae and A. gallica (Entry and Cromack 1989) and volatiles in Scots pine oleoresin reduced the growth of Armillaria by one-half (Rishbeth 1972a). However, powdered wound resin from ponderosa pine, when added to malt extract agar, significantly in¬ creased Armillaria growth compared to the basic me¬ dium (Shaw and Roth 1976). Similarities may exist between Heterobasidion annosum (Fr.) Bref. and Armillaria in how oleoresin affects myce¬ lial growth. Oleoresin did not affect H. annosum growth in agar culture. Prior (1976) suggested that resin-im¬ pregnated root wood of Corsican pine was a physical impediment to the fungus, reducing mycelial growth rate by more than one-half compared to non-resinous roots. Rykowski (1975) observed that resin-soaked wood and callus around root lesions on Scots pine lim¬ ited spread of Armillaria ; hyphae were not found in the resinous wood. Similarly, in roots of young, vigorously growing Douglas-fir trees, the host checked infections by laying down a callus and resin barrier (Buckland 1953), thus forming a latent canker (fig. 5.4). Cankers were 2-3 cm long, covered with resin, bounded by cal¬ lus tissue, and often several years old. Within cankers, A. ostoyae either remained dormant or grew into the woody core of the root. Lesions at the root collar devel¬ oped from one or more diseased lateral roots (Day 1927b). After killing the cambium at the root collar and on the lower bole, further spread of Armillaria was checked and callusing occurred (figs. 5.5A,B). In conifer roots, a central column of decay caused by Armillaria 64 Diagnosis and Detection FIGURE 5.4 — A rmillaria ostoyae lesion on a Douglas-fir root. Note callusing at the margin of the lesion. (G.W. Wallis) was compartmentalized by a barrier zone consisting of complete rings of resin ducts and parenchyma or nu¬ merous resin ducts separated by tracheids (Tippett and Shigo 1981). In eucalypts, the development of decay in the roots and lower bole reflects differences in their response to A. luteobubalina (Shearer and Tippett 1988). Roots of jarrah often had bark lesions which were confined by new layers of periderm. Barrier zones formed in the xylem as a result of infection and were obvious boundaries between xylem produced before and after infection (fig. 5.6). Armillaria luteobubalina often girdles wandoo at the root collar because the tree does not resist tangential spread of the fungus in the inner bark. In contrast, cal¬ lus tissue formed by jarrah and messmate stringybark restricted tangential spread, causing inverted V-shaped lesions (Kile 1981, Shearer and Tippett 1988). On citrus trees attacked by A. tabescens, living roots had bark lesions up to 30 cm long, some of which were de¬ limited by callus (Rhoads 1948). Adventitious roots arising from callus tissue (fig. 5.7) may compensate for roots killed by Armillaria root dis¬ ease (Cooley 1943, Kile 1980b, Riggenbach 1966, Rishbeth 1985b). The incidence of mortality by Armillaria root disease of¬ ten decreases with increasing plant age, particularly in conifers (Buckland 1953, Gibson 1960, Johnson and oth¬ ers 1972, MacKenzie 1987). This decrease usually is at¬ tributed to increased host resistance with age, which could be associated with physiological or biochemical changes in the host. In lodgepole pine, resin production increases with age until about 50 years (Shrimpton 1973). The ability of conifers to form callus where le¬ sions form on lateral roots and the root collar increases between age 5 and 20 years (Johnson and others 1972). Post-Infection Development Post-infection development of Armillaria root disease in a host root system depends upon the susceptibility, size, and age of the host (see below), the pathogenicity (see chapter 6) and inoculum potential (see chapter 4) B FIGURE 5.5 — A: Armillaria ostoyae lesion on the lower bole of a 21-year-old Douglas-fir. Note loosened bark and blackened resin. B: Cross section through lesion in (A). Note active callusing of lesion. (D.J. Morrison) Diagnosis and Detection 65 FIGURE 5.6 — Armillaria luteobubalina lesion on a greatcone banksia root. One side of the root was killed by the fungus. The area of xylem discoloration is bounded by a barrier zone. (Figure 5E from Shearer and Tippett 1988) FIGURE 5.7 — Adventitious roots arising from a callused A ostoyae lesion on a Douglas-fir root (1- living root; 2- adventi¬ tious roots; 3- A. ostoyae-killed root). (D.J. Morrison) of the fungus, and the influence of environment on host-fungus interaction (see chapters 7, 8). In suscep¬ tible hosts, the rhizomorph which causes infection pen¬ etrates to the cambium, becomes R. subcorticalis, and spreads laterally in all directions through the cambial zone (Woeste 1956). Growth of mycelial fans in the outer bark may precede that in the cambium; that is, growth is ectotrophic. The extent of ectotrophic growth is variable. In messmate stringybark mycelium of A. luteobubalina in the outer bark was up to 1 m ahead of cambium infection (Marks and others 1976). In Scots pine (Redfern 1978), mycelium of A. ostoyae was only 2 cm ahead of established infection proximal to the infec¬ tion point. As occurs with the penetrating rhizomorph branch, mycelial fans act as a unit, and host tissues are affected ahead of them. Schmid (1954) described the in¬ vasion of spruce bark by R. subcorticalis. In the xylem. mycelium penetrates the rays and spreads from them laterally into the xylem elements (Dade 1927, Woeste 1956). Continued killing of host tissues in the cambial zone girdles the root. The fungus spreads distally and proximally from the point of infection, and on reaching the root collar it spreads to other primary roots. The location of infections is an important factor in dis¬ ease development. Whether the result of contact with rhizomorphs or diseased roots, infections at the root collar or on the tap root (if present) usually kill the host more rapidly than infections on lateral roots (Barss 1913, Gadd 1930, Shaw 1980). However, infections at either location may be lethal (Rhoads 1948). On sapling and pole-sized ponderosa pines, Shaw (1980) found that rhizomorph-initiated infections on lateral roots were common, although the fungus rarely advanced proximally more than a short distance from a girdling root lesion. Armillaria infections on lateral roots may have failed to spread proximally because of host re¬ sponse, because rhizomorphs and distal portions of small roots may have provided inadequate inoculum potential, or both. Lethal attacks occurred high on the tap root or on the root collar. Similar observations were made on young Douglas-fir (Buckland 1953), on red pines and eastern white pines, and on white spruce (Patton and Riker 1959). Rykowski (1975) described the development of disease in the root systems of Scots pines, showing seven distinct patterns of infection. Where rhizomorphs cannot establish progressive infec¬ tions or for species which do not form them in forest soils, infections develop at contacts between healthy and diseased roots. Contacts are more likely to occur on lateral roots than at the root collar. On cacao (Dade 1927), citrus (Rhoads 1948), Douglas-fir (Morrison 1981), and eucalypts (Pearce and others 1986, Podger and others 1978, Shearer and Tippett 1988), infections originating this way on lateral roots spread to the root collar (fig. 5.8) and then to the tap root and other lateral roots, eventually girdling the trunk. When Armillaria girdles a root, the portion distal to the infection is colonized rapidly by mycelial fans growing in the cambium (Redfern 1978, Shaw 1980). Redfern (1978) observed maximum spread of 110 cm (mean 62 cm) in 10 months in inoculated roots which had been severed. Effects on the Host In agricultural crops, Armillaria root disease may re¬ duce the quantity and quality of produce prior to a plant's death. In forest crops, the disease may reduce height and diameter growth, cause decay of the bole, or cause death of the host, directly or indirectly. 66 Diagnosis and Detection FIGURE 5.8 — Armillaria ostoyae infection spreading along a Douglas-fir root (1- mycelial fan In outer bark; 2- bark necrosis in advance of the mycelial fan; 3- cambial necrosis). (D.J. Morrison) Reduction in height and diameter increment is a conse¬ quence of partial killing of the host's root system. Ten- year-old radiata pine showed a highly significant difference in cumulative mean increment between healthy trees and those with more than 65% of root col¬ lar circumference showing symptoms of A. limonea or A. novae-zelandiae (Shaw and Toes 1977). Diameter growth of 70- to 80-year-old Norway spruce affected by Armillaria root disease was reduced from one to six times compared with healthy trees (Sokolov 1964). An¬ nual growth increment of diseased 80- to 120-year-old Norway spruce was about one-half that of healthy trees (Molin and Rennerfelt 1959). Kile and others (1982) ob¬ served reduced growth in messmate stringybark with over 25% of their root collar circumferences infected by A. luteobubalina. Norway spruce (110 years old) which were classified as heavily infected by Armillaria had wider growth rings early in the rotation than trees which were healthy or lightly infected (Hrib and others 1983). This suggests that faster growing trees become infected earlier and more frequently due to their more extensive root system and greater probability of con¬ tacting inoculum (Bloomberg and Reynolds 1985). Later in the rotation, ring widths of trees in the two highest infection classes were 1 mm or less compared to 3 mm in uninfected trees (Hrib and others 1983). MacKenzie (1987) estimated volume loss of 6%-13% due to lethal and sublethal infection over a 28-year ro¬ tation of radiata pine. Growth loss due to A. ostoyae in 80- to 100-year-old Douglas-fir was measured on trees stratified by the proportion of the root collar showing resinosis (Bloomberg and Morrison 1989). Growth dur¬ ing 5-year periods, expressed as a percentage of the stem volume at the start of each period, decreased sig¬ nificantly as resinosis increased due to colonization of the root system. In recently killed trees and in those with more than 50% basal circumference showing resinosis, growth began to decline 30 years previously. The volume increment of these trees during the last 5- year period was 10-50% less than that of healthy trees, depending on proportion of root system killed. Twenty- to 40-year-old Norway spruces with butt rot had one-sided root distributions because Armillaria had killed one or more primary roots through which it had entered the stem early in the life of the tree. A reaction zone from which bacteria could be isolated extended as far as 50 cm up the stem (Yde-Andersen 1958). Butt rot of older Norway spruce was recorded by Molin and Rennerfelt (1959). In Britain, butt rot of conifers is com¬ monly initiated when a small tap or sinker root is killed. The decay usually is limited to the lower 60 cm of the stem. Of species grown in Britain, Norway and Sitka spruces and western hemlock are most suscep¬ tible to butt rot while Douglas-fir, true firs, pines, and larches show considerable resistance (Gladman and Low 1963). Armillaria root disease may kill its hosts by girdling the stem at the root collar. Prior to death, diseased trees may be windthrown due to decay of structural roots (Gladman and Low 1963, Shaw and Toes 1977), or they may be attractive to bark beetles which kill all or part of the tree (Cobb 1989). Physioloqy of Symptom Development and Host Killing The physiological basis of symptom development and host mortality is little understood for Armillaria, but two hypotheses have been proposed. First, symptoms develop as a direct result of the fungus physically dis¬ rupting the host's vascular system and the host's re¬ sponses to it. Second, Armillaria species may produce metabolic toxins. The first hypothesis has been ac¬ cepted by many investigators due to the nature of symptoms induced by Armillaria, particularly in the fo¬ liage. In mature conifers, shoot growth declines and the amount and color of foliage change gradually over sev¬ eral to many years as Armillaria destroys the host's vas¬ cular tissue. This view is supported by the results of Kile and others (1982), who found that patterns of elec¬ trical resistance were similar in mechanically girdled trees and those killed by A. luteobubalina. However, no experimental studies are known of host physiological parameters relative to location or extent of root system infection. Several authors have postulated that symptoms are caused by a toxin produced by Armillaria. Orchard Diagnosis and Detection 67 trees affected by Armillaria appeared to exhibit symp¬ toms of toxicity, possibly due to effects of metabolic products of the fungus (Zeller 1926). He suggested that branches died from toxic products since only branches above diseased roots showed symptoms, and pruning an infected root did not result in branch death. This view is supported by results of Thornberry and Ray (1953) who obtained a dark brown protein-like pig¬ ment from a liquid culture of Armillaria. The fungus had been isolated from a wilting peach tree. The sub¬ stance induced wilting in tomato seedlings and peach twigs and penetrated 15-20 mm into vascular tissues. However, electrical resistance measurements around actively expanding lesions did not show that A. luteobubalina produces any systemic effects in eucalypts (Kile and others 1982). Further research is needed to clarify the physiology and biochemistry of killing of host tissues (see chapters 3 and 7). Understanding this process could lead to characterization of pathogenic species and suggest what makes a host resistant to disease development. Disease Diagnosis Woody plants express diverse symptoms which may be categorized, in approximate chronological order, as follows: reduction of shoot growth, changes in foliage characteristics, crown dieback, stress-induced repro¬ duction, basal stem indicators, and death. Generally, the nature of the symptoms and their rate of develop¬ ment relate to the position of attack and the rate of de¬ struction of the host root system. If the disease progresses rapidly or the host is small, not all symp¬ toms may be evident (Hartig 1874, Edgar and others 1976). Symptom development in conifers was more pronounced on vigorous hosts (Buckland 1953). Above-Ground Symptoms on Individual Plants Reduction of Shoot Growth On conifer seedlings and trees up to about 10 years old, Armillaria rarely reduces shoot growth prior to death because killing occurs within a few months to a year af¬ ter infection (Gibson 1960, Hartig 1874, Hintikka 1974). By contrast, the slower progress of the disease in older conifers causes a decline in shoot growth (fig. 5.9) which may be evident for many years (Molin and Rennerfelt 1959). In 80- to 100-year-old Douglas-fir, Bloomberg and Morrison (unpubl.) found terminal- shoot growth on diseased trees had declined for the previous 15-30 years. Actual time depended on the time since infection. Fruit trees affected by Armillaria root disease may have a stunted appearance due to a shortening of internodes (Barss 1913, Cooley 1943). Changes in Foliage Characteristics On conifers which are killed quickly, foliage turns red or brown as it dries. When the disease progresses slowly, as in older trees, foliage gradually becomes stunted, chlorotic, and sparse (fig. 5.9). These changes usually occur throughout the crown (Hartig 1874, Molin and Rennerfelt 1959, Morrison 1981, Williams and others 1989). Symptoms in the crowns of young Douglas-firs are frequently accompanied by prolific resin blisters on the stem and branches (Buckland 1953). Small hardwood trees frequently are killed so rapidly by A. tabescens that symptoms are not evident until the foliage withers and dies (Rhoads 1956) whereas the first indication of infection on larger trees is a thin crown with small leaves (Guillaumin 1977, Sokolov 1964). Trees later show gradual yellowing and defolia¬ tion followed by rapid wilting and dying of individual FIGURE 5.9 — A 12-year-old Douglas-fir showing reduced shoot growth (for 2 years), chlorotic foliage, and a stress- induced cone crop. (D.J. Morrison) 68 Diagnosis and Detection limbs above diseased roots (Barss 1913, Bliss 1944, Rhoads 1956). On apple trees, premature defoliation is sometimes an indicator of Armillaria infection (Marsh 1952); and on diseased stone fruit trees, leaves roll along the mid-rib and wilt (Cooley 1943). Attacked by 4. luteobubalina, eucalypt saplings up to 25 years old die suddenly (fig. 5.10), showing little deterioration of :rowns before death (Edgar and others 1976). On older saplings, leaves show gradual reddening followed by arowning and plant death (Pearce and others 1986). In pole-size to mature eucalypts, A. luteobubalina causes a general reduction in leaf density, drooping of foliage, ?picormic shoots along branches, and eventually a dead top (fig. 5.11). Large trees which could not com¬ partmentalize infections usually die 2-8 years after vis¬ ible crown deterioration appears (Edgar and others 1976, Pearce and others 1986). Crown Dieback In pole-size to mature eucalypts attacked by A. luteobubalina, dieback of fine twigs and branches may lead to a dead top (Edgar and others 1976). Cooley (1943) ob¬ served that limbs on apple trees ceased growth and died on the same side as the affected root. Frequently, the com¬ bined action of Armillaria root disease and other biotic or abiotic agents has been associated with crown dieback and eventual mortality of many forest species, such as those noted in chapter 7 and table 8.3. FIGURE 5.10 — A pole-stage mountain grey gum tree killed by A. luteobubalina. Little crown deterioration occurred prior to the sudden death of the tree. (G.A. Kile) FIGURE 5.11 — Messmate stringybark trees showing stages in crown decline caused by A. luteobubalina. (G.A. Kile) Stress-induced Reproduction Many woody plants respond to advanced infection by producing a seed crop, usually in the season before death. Thus, tung oil trees produce nuts which are smaller than normal (Rhoads 1956), orchard trees pro¬ duce poor, stunted fruit (Barss 1913), citrus trees de¬ velop an abnormally heavy bloom (Rhoads 1948), and conifers produce cones which are smaller but may be more numerous than normal (fig. 5.9) (Buckland 1953). Basal Stem Indicators Woody plants attacked by Armillaria frequently pro¬ duce exudates or develop cankers, cracks, or flutes at or just above the root collar. Genera of conifers which normally have resin canals (Pseudotsuga, Picea, Larix, and Pinus) or which form traumatic resin canals (Tsuga and Abies) may produce resin that exudes through fis¬ sures (fig. 5.12) in the bark of the root collar and lower bole (Buckland 1953, Gibson 1960, Hartig 1874, Hintikka 1974, Rykowski 1975). Usually, resin exuda- Diagnosis and Detection 69 FIGURE 5.12 — Copious basal resinosis on a radlata pine attacked by A novae-zelandiae or A. limonea. (C.G. Shaw III) tion is not evident above-ground until the fungus is near or has reached the root collar. Responding to ad¬ vanced A. tabescens attack, citrus trees occasionally (Rhoads 1948) and stone fruits commonly (Rhoads 1956) produce gum in the cambial region which may be so copious as to exude through cracks in the bark. Latex exudes from rubber trees at the root collar in the last stages of the disease (Riggenbach 1966). Exudation of kino through stem and root bark occurs on some ma¬ ture eucalypt trees infected by A. luteobubalina; and from stems of trees less than 20 years old, it may be abundant, permeating and blackening the adjacent soil (Edgar and others 1976, Kile 1981). Infections by Armillaria in 20- to 70-year-old Douglas- fir, white pine, and other conifers may be arrested after killing cambium at the root collar above a diseased root. Callusing occurs around the margin of the lesion. When fresh, lesions are resinous and have mycelial fans beneath the bark. Later, after the bark sloughs, the le¬ sions can still be recognized by their short length, broad triangular shape, and the impressions of mycelial fans on the scar face (Molnar and McMinn 1960). Conical basal scars on eucalypt stems (fig. 5.13) are frequently associated with A. luteobubalina infection (Kile 1981, Pearce and others 1986, Shearer and Tippett 1988). In citrus, basal lesions extend up to 35 cm above one or more diseased roots and may serve as entry points for other wood-rotting fungi (Rhoads 1948). The lesion at the base of some oil palms remains localized, dried, and apparently sealed off from the healthy tissue within; a mass of new roots forms above the canker (Wardlaw 1950). West African rubber trees infected with Armillaria or with Rigidoporus lignosus (Kl.) Imaz develop flutes at the stem base starting at the root col¬ lar near the point of infection (Riggenbach 1966). A diagnostic symptom of Armillaria root disease on woody plants such as tea, coffee, and cacao in tropical or subtropical regions is the conspicuous longitudinal cracks that appear at the root collar and quickly extend up the trunk, hence, the name "collar crack" (Dade 1927). The cracks are longer and more numerous on the side of the tree where infection occurred. Similar cracks were observed on the roots and lower stem of citrus at¬ tacked by A. tabescens (Rhoads 1948) and on the roots of several hardwood species (Sokolov 1964). In standing trees, heartwood decay (butt rot) does not produce external signs unless it is associated with a basal canker. In felled timber, butt rot caused by Armillaria may be recognized by characteristics of the decayed wood or confirmed by culturing. Where decay of structural roots is advanced in coniferous and broadleaved trees, they may be windthrown prior to death. This is particularly true where the tree is being sustained by adventitious roots. Symptom Development in Relation to Extent of Colonization The development of symptoms of Armillaria root dis¬ ease in foliage and at the stem base depends upon the rate and degree of invasion of the host root system. Thus, on young (Gibson 1960, Swift 1968) or small trees (Rhoads 1956) in which the root system is invaded rap¬ idly after infection, symptoms may appear just prior to death or only after the host is moribund. Death of ra- diata pine due to A. novae-zelandiae or A. limonea began 6 months after planting (MacKenzie and Shaw 1977). In 8- to 10-year-old plantations, an eastern white pine died 39 months after inoculation and a red pine in¬ fected by natural inoculum died 14 months after root examination showed it to be healthy (Patton and Riker 1959). On apricot trees, symptoms on aerial parts ap¬ peared only after the root collar was attacked or several large roots were killed (Guillaumin 1977); and on apple trees, girdling of the stem was complete 2-3 years after infection was first noted in one segment of the trunk (Marsh 1952). Diagnosis and Detection Invasion of the root system of old or large trees usually occurs slowly over many years. Growth-ring studies on conifers 80 to 110 years old suggest that recently dead and severely affected trees became infected up to 50 years previously (Molin and Rennerfelt 1959). Conse¬ quently, symptoms develop gradually after a portion of the root system is colonized. Bliss (1944) found that Armillaria root disease was well established in citrus roots before any symptoms appeared in the crown. The fungus must reach the root collar before exudation of resin, gum, or kino becomes visible. More than half the root system of grand firs (mean age 50 years) had been killed by Armillaria with no apparent decline in tree vigor (Maloy and Gross 1963). Sokolov (1964) observed that the color and thickness of the crown and the inci¬ dence of cracks and resin flow on the lower bole were related to the proportion of first-order roots infected. In 80- to 100-year-old Douglas-firs, height growth reduc¬ tion and the percentage of stem circumference showing basal resinosis were proportional to the amount of the root system colonized by A. ostoyae (Bloomberg and Morrison 1989). Crown symptoms on these trees were not obvious until one-half to three-quarters of the pri¬ mary roots had been invaded. Crown dieback in- FIGURE 5.13 — Basal lesion on mountain grey gum caused by 4. luteobubalina. (G.A. Kile) creased with increasing root collar infection in eucalypts attacked by A. luteobubalina (Edgar and oth¬ ers 1976, Kile 1981); the height of infection on stems was positively correlated with circumference infected (Kile 1981). Confirmation of Armillaria Occurrence Many symptoms described above are non-specific; that is, they may be induced by a number of biotic and abi¬ otic factors. To confirm Armillaria root disease, the root collar and lower bole of the tree must be examined for signs specific to the fungus. Those signs include myce¬ lial fans, rhizomorphs, basidiomes, and decay. Armillaria may also be confirmed by culturing from the host. Many of the signs are useful for identifying stumps and roots which are within disease centers or on cutover sites, and which may be inoculum sources for the next rotation. Mycelial Fans On plants showing symptoms of advanced infection and on those recently killed, creamy-white mycelial sheets up to 10 mm thick occur in the cambial zone of roots and the lower bole (Greig and Strouts 1983, Morrison 1981, Williams and others 1989). The mycelial sheets, commonly known as fans and occasionally re¬ ferred to as xylostroma, are the most useful diagnostic characteristic of Armillaria species in woody plants (figs. 5.14A,B). The mycelial fans of some Armillaria species are marked with perforations (fig. 5.15) 0.2-3 mm in diameter (Gibson and Corbett 1964, Kile and Old 1982, Rhoads 1945). In plants which have been dead for several years, mycelial fans usually can be found in roots below-ground but have disappeared from above-ground parts due to competing fungi or to unfavorable environmental conditions, such as desicca¬ tion. On conifers, impressions of fans in resin and bark may be present for several years after fans disappear (fig. 5.16). Several reports of Armillaria on African crops (Dade 1927) and on hosts of A. tabescens in Florida (Rhoads 1948) refer to frills of xylostroma, at first cream-colored then becoming dark brown with age, which protrude from the longitudinal fissures in the bark. The descrip¬ tion by Dade (1927) indicates that xylostroma sheets are extensions of subcortical mycelial fans which be¬ come melanized when exposed to air, an observation confirmed by Rhoads (1948). Rhizomorphs Rhizomorphs are initiated on the food base from the edges of mycelial fans, either subcortically when condi¬ tions such as loosening of bark prevents further growth Diagnosis and Detection 71 A FIGURE 5.14 — A: Mycelial fans of A. ostoyae in the cambial zone of an 8-year-old Douglas-fir. (D.J. Morrison). B: Mycelial fans of A luteobubalina on brown barrell eucalypt. Note rhizomorphs emerging from the fan margin where the bark was loosened. (G.A. Kile) FIGURE 5.15 — Perforated mycelial fans of A. luteobubalina developed in vitro in stem segments of silver wattle. (G.A. Kile) of the fan (fig. 5.14), or into soil when the fan reaches the bark-soil interface (Morrison 1972). For up to 1 cm from the growing tip, a rhizomorph is white; with in¬ creasing distance from the tip it becomes red-brown. brown, and finally black. A rhizomorph is hollow near the growing tip; however, within 2 cm, the hollow be¬ comes filled with randomly arranged fiber hyphae in a mucilaginous matrix (Redfern 1973, Schmid and Liese 1970). Rhizomorphs in soil and on the surface of roots are usually 1-3 mm in diameter (Morrison 1972, Pearce and others 1986, Redfern 1973). Occasionally, rhizo¬ morphs in soil, probably of A. gallica, are 5 mm in diameter (Redfern 1973). Rhizomorph structure is dis¬ cussed fully in chapter 3. In the north temperate zone (Greig and Strouts 1983, Wargo and Shaw 1985), New Zealand (Hood and Sandberg 1987), and at higher elevations in East Africa (Gibson 1960), India (Satyanarayana and others 1982) and Sri Lanka (Gadd 1930), rhizomorphs of ArmiUaria species grow freely through soil and on the surface of roots. The rate of growth and distance from the food base that they will grow varies greatly among species. Species with monopodially branched rhizomorphs, such as A. gallica, often produce extensive networks in 72 Diagnosis and Detection FIGURE 5.16 — Impressions of A. ostoyae mycelial fans on the inner bark of Douglas-fir. (D.J. Morrison) soil, whereas dichotomously branched species, notably A. mellea (Rishbeth 1982), appear to be restricted to within a few centimeters of the food base. For this rea¬ son, the usefulness of rhizomorphs as a diagnostic fea¬ ture is limited, particularly at the specific level. At low elevations in the tropics, rhizomorphs are not found in soil or on roots (Dade 1927, Gibson 1960, Rishbeth 1980, Swift 1968), although occasionally they grow up to 2 cm into soil and then die (Dade 1927). In Australia, rhizomorphs either were not observed in the field (Kile 1981, Shearer and Tippett 1988) or were found only on the surface of roots (Kile 1980b, Pearce and others 1986). Basidiomes Basidiomes occur in clusters arising from mycelial fans in the host or in small numbers from rhizomorphs on the host or in soil. Basidiomes facilitate surveys of dis¬ ease incidence (Pearce and others 1986) and identifica¬ tion of the Armillaria species (see chapter 1). Basidiom¬ es often occur on or near hosts lacking other signs and symptoms. In temperate regions, fruiting occurs from mid-summer to mid-winter, depending on latitude and weather. Precipitation and favorable temperatures are required to initiate fruiting and for basidiome develop¬ ment. Basidiomes which develop slowly due to cold or dry weather may have short, thickened stipes and small thick pilei; weather may also affect basidiome color (Kile and Watling 1981). In tropical regions, bas¬ idiome formation varies from rare in Sri Lanka (Gadd 1930) and East and Central Africa (Wallace 1935, Gibson 1960b, Swift 1972) to common in West Africa (Dade 1927, Riggenbach 1966), where it occurs almost exclusively in the wet season (Wardlaw 1950). Armillaria species cause a white rot of woody tissues as lignin and cellulose both decompose. The appearance of decayed wood varies somewhat among hosts. In co¬ nifers, wood with incipient decay is stained gray to brown, often with a water-soaked appearance. Later, decayed wood becomes yellow-brown and stringy (figs. 5.17 and 5.18) and is finally reduced to a very wet, stringy rot with pale yellow flecks (Greig and Strouts 1983, Williams and others 1989). Decayed wood of broadleaved hosts is watersoaked and white to yellow, becoming spongy and ultimately distinctly gelatinous (Greig and Strouts 1983, Rhoads 1956). Pseudosclerotial plates (zone lines) are common in woody tissues decayed by Armillaria species (Campbell 1934, Lopez-Real 1975, Greig and Strouts 1983, Podger and others 1978). These plates are composed of pig¬ mented bladder hyphae which are identical with the cells comprising the outer coat (rind) of mature rhizomorphs (Campbell 1934, Lopez-Real 1975). Wood decayed by some, but not all, Armillaria species is bi- oluminescent (Kile 1980b, Podger and others 1978). The biochemistry of bioluminescence is discussed in chapter 3. Isolation Technique and Appearance in Culture The presence of an Armillaria species in host tissue may be confirmed by culturing colonized bark or wood or subcortical mycelium on a medium such as potato dex¬ trose or malt extract agar. Molds or bacteria may be suppressed by acidifying the medium or amending it with a fungicide such as o-phenylphenol (Russell 1956) or benomyl (Hunt and Cobb 1971, Maloy 1974). Isola¬ tion of Armillaria from root tissues of dead and dying trees increased by 40% on malt agar amended with o- phenylphenol (Whitney and others 1978). The selective media developed by Kuhlman (1966) and Kuhlman and Hendrix (1962) for isolating H. annosum from wood and its spores from soil also is selective for Armillaria (Shaw 1981a). The fungus may be isolated from rhizomorphs by first washing short lengths in water then soaking them in 10% hypochlorite for 5 min (Rishbeth 1978b). Hood and Sandberg (1987) made iso¬ lations from rhizomorphs after dipping them in 95% ethanol, surface sterilizing in 10% hydrogen peroxide, and washing in distilled water. Nobles (1948) suggested that Armillaria cultures are rec¬ ognizable from macroscopic appearance alone, their red-brown crustose areas, rhizomorphs, and frequent luminosity of young, actively growing colonies being unique. Her description is based on four isolates, three of which were from conifers in British Columbia and Diagnosis and Detection 73 FIGURE 5.17 — Yellow stringy decay of Douglas-fir root wood caused by A. ostoyae. (D.J. Morrison) FIGURE 5.18 — Armillaria-caused butt rot of Norway spruce. (B.J.W. Greig) Washington. It is likely that the description is based on cultures of A. ostoyae. However, these features are char¬ acteristic of most, if not all, species of Annillaria. Differ- tiation of vegetative isolates of Armillaria is discussed : chapters 1 and 2. Biotic and Abiotic Conditions Causing Similar Symptoms Any agent or condition which affects the root system of a woody plant may cause some or all of the symptoms described above. In conifers, root diseases caused by H. annosum (Greig and Redfern 1974), Phellinus weirii (Murr.) Gilbn. (Thies 1984, Wallis 1976), Inonotus tomentosus (Fr.) Teng (Whitney 1978a) and Leptographium spp. (Wingfield and others 1988) may cause crown symptoms similar to those of Armillaria. On apple trees, winter injury to the roots or root collar or root suffocation due to flooding can induce symp¬ toms similar to Armillaria root disease (Cooley 1943). Stem girdling or root killing due to any cause induces foliage symptoms in citrus similar to those of Armillaria root disease (Rhoads 1948). Disease Detection Detecting Armillaria root disease in production forests, amenity woodlands, and agricultural plantations de¬ pends on observable symptoms in the crown and on the stem base plus signs of the fungus such as mycelial fans, rhizomorphs, and basidiomes on the host. Dis¬ eased trees occur as scattered individuals or in centers which reflect the distribution of the Armillaria species. Characteristics of disease centers are discussed in chap¬ ters 8, 9, and 10. Aerial photography and ground surveys conducted in¬ dependently or in combination have been used to de¬ tect root diseases, including those caused by Annillaria. Choice of survey method is influenced by the purpose of the survey. For example, the survey may intend to determine presence or absence of root disease, estimate wood volume in diseased trees, delineate distribution of disease, or provide input data for modeling pur¬ poses (see chapter 10). Aerial photographs (Kable 1974) and stem maps (Marsh 1952) also have been used to detect and record progress of Armillaria root disease in agricultural plantations. Using aerial photography permits large areas of forest to be inspected rapidly for visibly affected trees, for quantifying effects, and for providing a record of dis¬ ease occurrence. Some ground inspection is required to identify the pathogenic species involved and to verify the photographic assessment. The choices of image scale and film emulsion to be used are based on stand structure, ease of defining disease signature, and pur¬ pose of the imagery. While detection of disease centers and affected single trees may be accomplished at scales up to 1:10000 (Gregg and others 1978, Murtha 1972, Myers and others 1983, Williams 1973), larger scale im¬ agery, 1:1000-1:2000, may be necessary to provide rea- 74 Diagnosis and Detection onable accuracy in delineating areas affected. Gener- illy, relatively large scale imagery, 1:3000-1:6000, is nost often used for detecting and quantifying indi¬ vidual trees or centers (Gregg and others 1978, Myers nd others 1983, Wallis and Lee 1984, Williams 1973, Villiams and Leaphart 1978). Color and false color color infrared) emulsions are frequently used (Gregg nd others 1978, Heller and Bega 1973, Williams and ,eaphart 1978); black and white may also be effective fohnson and Wear 1975). n western North America, the signature of root disease enters on aerial photographs included openings in the orest canopy with dead or nearly dead standing trees in the margins, snags, and windthrown conifers, and ;enerally a shrub cover and some young trees in the ipening (Wallis and Lee 1984, Williams and Leaphart 978). Dead trees and crown decline characterized A. uteobubalina centers on photographs of jarrah forests in Vestern Australia (Shearer and Tippett 1988). Ground evaluations using various survey procedures re efficient if areas are small or if precise disease loca- ion and damage measurements are required. Survey lesign varies from regularly or randomly spaced ransects to systematically spaced variable and fixed- adius plots (Jacobi and others 1981). Pearce and others 1986) used random reconnaissance, transect and plot urveys to determine the occurrence of basidiomes and he incidence of infection in stumps, saplings, and rees. The ground survey method developed for P. veirii (Bloomberg and others 1980a,b) and modifica- ions for multiple-disease recording and stratification >y infection intensity (Bloomberg 1983) are applicable o surveys for Armillaria root disease. This transect ampling system involves randomly located sets grids) of lines to estimate the incidence, distribution, md area of root disease. Estimates of diseased area are lerived from length of transect intersecting root dis- ase centers and probability of occurrence. Random lo- ation of gridlines in a stand results in independent stimates for each grid, hence the variance of their neans can be estimated. rhe Bloomberg method is difficult to apply in logged, turned, or open stands with diffuse disease distribu- ion because locating infection boundaries can be diffi¬ cult. For that reason, Kellas and others (1987) used sys¬ tematically located transects with variable-sized plots around selected stumps to assess infection by A. luteobubalina in regeneration, regrowth, and overwood trees. Incidence and severity of Armillaria root disease can be assessed during inventory surveys (B. Geils, unpubl.). Ground survey data such as that frequently collected by the USDA Forest Service (1986) may be used to initialize a model of Armillaria root disease (see chapter 10), if augmented to include stumps in¬ fected with root disease (Stage and others 1990). Where survey information is required for large areas, multi-stage or double sampling designs incorporating aerial photography and ground evaluations can be em¬ ployed (Stewart and others 1982, Williams and Leaphart 1978, Wood 1983). Conclusions The infection process has been observed on hardwood and coniferous hosts. Post-infection disease develop¬ ment has been observed for a few host species but not throughout a rotation. The response to infection by a variety of host species has been recorded, primarily at the macroscopic level, but less is known of the interac¬ tions between hosts and Armillaria at the biochemical level. The effects of Armillaria root diseases on their hosts, growth loss, decay, and mortality, are known. Symptoms of Armillaria root diseases which are non¬ specific include reduction of shoot growth, changes in foliage characteristics, crown dieback, stress-induced reproduction, basal stem indicators, and death. Signs specific to Armillaria species are subcortical mycelial fans, rhizomorphs, and basidiomes. Cultures of Armillaria have distinctive characteristics. Ground and aerial methods for detecting Armillaria root diseases and ground procedures for determining disease area have been developed although work is needed to im¬ prove their utility. Understanding the biochemistry and physiology of the host-parasite interaction and studies of disease development during a rotation for representative combinations of host and Armillaria spe¬ cies remain the most urgent research needs relating to infection and disease development. Diagnosis and Detection 75 CHAPTER 6 Pathogenicity and Virulence Steve C. Gregory, John Rishbeth, and Charles G. Shaw III T he terms pathogenicity and virulence both refer to an ability to cause disease. That "Armillaria niellea" can cause disease has been known for over a century, but its propensity to do so has been a matter of controversy. Rhizomor- phs commonly surround tree roots without infecting them, yet Armillaria may cause extensive mortality elsewhere in the same area. Such observations were interpreted by some early authors as indicating that trees in affected areas were weakened or predisposed to infection in some way (Day 1927b, 1929). Others, for example Piper and Fletcher (1903) and Childs and Zeller (1929), proposed that there were several forms of the pathogen that differed in virulence. According to the former view, Armillaria was a second¬ ary pathogen capable of attacking only trees with low¬ ered resistance. Thus, Day (1929) concluded that "all the evidence goes to show that it is always secondary to some other factor acting as a primary cause of dis¬ ease." Boyce (1961) stated that the fungus "does not attack thrifty trees" and Gremmen (1976) expressed the view that "control of A. mellea in forestry ... has no ef¬ fect since the fungus is not the primary cause of die- back." Contrary to these assertions, however, there are early accounts of Armillaria disease (for example Hen¬ drickson 1925, Zeller 1926) that describe attacks by a fungus with every appearance of a "virulent primary pathogen," as it was termed by Patton and Riker (1959). Dade (1927) similarly described the behavior of '‘Armillaria mellea" in tropical West Africa, and Brooks (1928) regarded it as "perhaps the most dangerous subterranean parasite of trees, bushes and certain her¬ baceous plants." Many contradictions regarding the pathogenic behav¬ ior of " Armillaria mellea" can now be understood as having arisen from observations made on different Armillaria species. They can differ markedly in patho¬ genicity yet closely resemble each other in the appear¬ ance of their basidiomes, rhizomorphs, and mycelial sheets. The extremely low pathogenicity of some spe¬ cies may partly explain the dismissive attitude some earlier authors held toward Armillaria as a pathogen. Pathogenicity, Virulence, and Disease Distinguishing between pathogenicity and virulence is especially important when so many species and so many different hosts are involved. "Pathogenicity" means the quality or characteristic of being able to cause disease as applied to a genus or species (British Federation of Plant Pathologists 1973). "Virulence" means the observed infective capacity of individual entities of a pathogenic species (British Federation of Plant Pathologists 1973). Pathogenicity of an Armillaria species was first estab¬ lished in an inoculation experiment by Hartig (1874) though his method fell short of satisfying Koch's postu¬ lates, which are now generally accepted as the require¬ ments for proving pathogenicity (British Federation of Plant Pathologists 1973). An extensive world literature on Armillaria now contains enough data from inocula¬ tion experiments to leave no doubt that several patho¬ gens occur in the genus. Some physiological attributes of the fungus that may be associated with high or low virulence are discussed in chapter 3, but the genetic basis of virulence in Armill¬ aria is largely unknown. Two studies have shown that haploid isolates derived from single basidiospores ma\ display high virulence, in some cases as high as the parent isolate (Raabe 1972, Shaw and Loopstra 1988). The wider genetic significance of this finding and its relevance to field behavior remain to be investigated. Reaves and others (1988) suggested that the occurrence of virus-like particles in the cytoplasm of some Armill¬ aria isolates might be associated with high virulence, but little evidence supports this hypothesis. 76 Pathogenicity and Virulence Saprophytic and Parasitic Behavior Arrnillaria species have both saprophytic and parasitic phases in their life cycle, but distinguishing the two may be difficult in an activity such as colonization of a moribund stump. By causing root- and butt-rot in standing trees, Arrnillaria species can also be classified as perthophytes because they utilize dead tissues in living hosts (British Federation of Plant Pathologists 1973). Most of the methods of capturing resources for saprophytic or perthophytic exploitation that have been outlined in chapter 4 depend on the fungus' abilities as a parasite even though the tissues involved might be of extremely low vigor, as in stumps and dying trees. Pathologists and mycologists now recognize that Arrnillaria species differ markedly in pathogenicity. Highly pathogenic species survive saprophytically in the hosts they kill through their parasitic activities, whereas weak pathogens probably exist for the most part as saprophytes or possibly perthophytes (Korhonen 1978, Rishbeth 1985a, Wargo and Shaw 1985). This diversity poses the question whether weakly pathogenic species are better able than highly patho¬ genic species to colonize moribund tissues and compete with saprophytic micro-organisms. Little information is available on this subject, and it is clearly an area that merits further research. Rishbeth's (1985a,b) experiments with excised root and stem material demonstrated that, in some circumstances at least, the weak pathogen A. gallica is no more capable than the highly pathogenic A. mellea of colonizing woody material with residual host resistance and may even have an inferior ability to penetrate intact bark on such material. The same studies suggest that these two species may differ little in their ability to colonize com¬ pletely dead material, and both may possess consider¬ able competitive saprophytic ability ( sensu Garrett 1956a, 1970). Arrnillaria ostoyae, another highly pathogenic species, did not perform as well as A. mellea and A. gallica in Rishbeth's (1985a,b) tests with excised material. In west¬ ern North America A. ostoyae is considered incapable of colonizing stumps that were not already infected as living trees (Filip 1989a). Although it is one of the as¬ sumptions underlying recent models of disease devel¬ opment (see chapter 10; McNamee and others 1989), the reasons for this apparent inability are not clear. It may reflect the species' limited capacity for spreading by rhizomorphs as much as any deficiency as a sapro¬ phytic competitor. An important attribute of weakly pathogenic species is an ability to act as facultative parasites on stressed or sickly hosts (Kile 1980b, Rishbeth 1985a, Wargo and Shaw 1985). However, many observations suggest that highly pathogenic species are also capable of invading weakened hosts (Davidson and Rishbeth 1988, Dumas 1988, Gregory 1989, Guillaumin and others 1989a, Rish¬ beth 1985a). In nature, it is probable that the weakly pathogenic species more often do so (Kile 1980b, Kile and Watling 1983, Rishbeth 1982). Quite possibly, some of the less pathogenic Arrnillaria species have evolved strategies, such as rhizomorph behavior, that confer advantage of position in exploit¬ ative situations (Gregory 1985, Rishbeth 1985a, Wargo 1984b, see chapter 4). Indeed, the paucity of data per¬ mits more general speculation that the undoubted suc¬ cess of such species owes less to any greater ability to penetrate and invade weakened or dead tissues than to an ecology that affords them the maximum opportu¬ nity to encounter such material. This is more fully dis¬ cussed in other chapters, but it is relevant to note here that such considerations necessitate great caution in interpreting observations and experiments on patho¬ genic behavior. Conditions For Disease Implicit in the definition of pathogenicity is the qualifi¬ cation that measurement should be made under speci¬ fied conditions. Among the most important elements that may influence the expression of pathogenicity are the host, the external environment, and the nutrition of the pathogen. Pathogen nutrition is contained in the concept of inoculum potential which was elaborated by Garrett (1956a, 1970). The ability of a pathogen, what¬ ever its inherent virulence, to cause disease is strongly influenced by the energy available to it at the host sur¬ face. The subject of inoculum potential is discussed in chapter 4. Host resistance is an important constraint on disease, and many studies have shown that susceptibility to Arrnillaria disease differs among host species. European forest hardwoods have been shown to possess consid¬ erably more resistance than native or exotic conifers (Redfern 1978, Rishbeth 1984), results that are in accord with most field observations. However, some conifers are notably resistant (Guillaumin and Pierson 1978) and some hardwood genera, Prunus and Citrus, for example, are notoriously susceptible (Guillaumin and Pierson 1978, Raabe 1967, Wilbur and others 1972). Differences in susceptibility of woody species within individual genera have frequently been demonstrated (Benjamin and Newhook 1984b, Guillaumin and others 1989b, Proffer and others 1988); and Azevedo (1970-71) found that two forms of the same species (Japanese redcedar) also differed. Pathogenicity and Virulence 77 Host resistance is not only a genetic attribute but also a result of circumstances. Notwithstanding the historic controversy over the role of host predisposition in Ar¬ millaria pathogenesis, factors associated with low host resistance will favor disease. Good circumstantial evi¬ dence from several parts of the world indicates young trees are more prone to infection than older trees of the same species (Gibson 1960, Ono 1970, Pearce and oth¬ ers 1986, Redfern 1978), and many pathologists believe stress imposed by ill-health, injury, or unsuitable growing conditions can increase susceptibility (see chapter 7). The best known limitations imposed by the external environment on the activities of pathogenic Armillaria species are the effects of soil on rhizomorph growth and production. The complicated relationships be¬ tween rhizomorphs and disease are discussed briefly in the following sections and more fully in chapter 4. Decay and Disease The commonly accepted definitions of disease refer to deviation from normal functioning of physiological processes (British Federation of Plant Pathologists 1973). It is therefore arguable whether butt rot, which involves the decay of largely non-living interior wood in living trees, constitutes disease. We will accept it as such since living cells in the wood are likely to be af¬ fected to some degree in many cases. The ability of Armillaria species to cause decay in standing trees is therefore an expression of pathogenicity though it ap¬ pears not to have been investigated experimentally. Most experiments assess virulence entirely by the effects of the pathogen's development in the phloem and cambium. In practice, root killing and root decay are often not clearly separable since one closely follows the other. Nevertheless, these processes involve the capacity to invade and exploit two quite different tissues, and the decay-causing ability of an isolate is not necessarily related to its capability as an agent of tree mortality. Decay has been little studied in Armillaria, but field observations in Europe (Gregory 1989, Korhonen 1978, Rishbeth 1982) suggest that species with limited ability to kill trees are associated with butt rot at least as often as highly pathogenic species. Host Specialization Many Armillaria species have a wide host range, both among the genera which occur naturally in their habi¬ tat and among exotics. For example, the Australian species A. luteobubalina not only attacks many native and shrub species in many genera but is also pathogenic to some North American conifers (see chapter 8; Morrison 1989). Such behavior does not preclude the existence of adaptive relationships be¬ tween particular pathogens and particular hosts ("host specialization" or "host preference"), though few have been clearly demonstrated in Armillaria. In Europe, the area from which most data are available, A. ostoyae appears to be better adapted to coniferous hosts and A. mellea to hardwoods (Guillaumin and others 1985; Guillaumin and Lung 1985; Guillaumin and others 1989a; Rishbeth 1985a; Siepmann 1985). However, dis¬ tinguishing the effects of host specialization from those of site history and pathogen ecology is often difficult. Both may limit the opportunities for contact between the fungus and some potential hosts. Assessing Pathogenicity and Virulence The ability to cause disease can be estimated from di¬ rect measurement of the amount of disease actually caused in inoculation trials, observation of field behav¬ ior, or an assay of some feature thought to be associ¬ ated with the pathogen's ability to cause disease. All three approaches have been attempted with Armillaria, but the first two have undoubtedly been the most useful. As already discussed, the intrinsic ability of an Armill¬ aria species to cause disease may be modified by cir¬ cumstances and environment. Hence, inoculation trials must be conducted under specified conditions, choice of which is exceptionally difficult with tree-root patho¬ gens, such as Armillaria, that have a wide host range and that can attack trees of virtually any age. More¬ over, the infection of such a massive and well-pro¬ tected structure as a woody root requires a specialized pathogen ( sensu Garrett 1970) for which the method of infection, and particularly the necessary inoculum po¬ tential, may be difficult to achieve artificially. For many Armillaria species, the chief means of infection is the rhizomorph, a specialized structure that may develop only under certain conditions and the efficacy of which is governed partly by the energy supplied to the infec¬ tive tip (Garrett 1956b). Choice of Host for Inoculation Trials Most investigators have selected trees or shrubs for pathogenicity trials. However, some have attempted to avoid the considerable difficulties of experimentation with intact woody hosts by using parts of plants or plant organs which may possess much less host resis¬ tance than a tree but might retain enough to repel iso¬ lates of low virulence. Large vegetable roots and tubers have proved valuable subjects for the study of infection biology. Garrett (1956b), Thomas (1934), and van Vloten (1936) used 78 Pathogenicity and Virulence potato tubers to demonstrate apparent differences in virulence between Armillaria isolates. Gregory (1984, 1985) and Rishbeth (1984) also attempted to use potato tubers to test virulence, comparing the results obtained with them to those obtained by using the same isolates on young trees. Although Gregory (1984,1985) found some correspondence, infection of the tubers generally occurred too readily for it to be pursued as a useful discriminatory method. The dangers of using material with low host resistance for determining the virulence of Armillaria isolates may be increased when the "host" is an excised root or stem. The ability to colonize moribund material may be of equal evolutionary advantage, and hence as well developed, in pathogens of low virulence as in those of high virulence. As discussed by Rishbeth (1985a,b), there is compelling evidence that Armillaria species of low pathogenicity can successfully colonize such mat¬ erial both in nature and in the laboratory. Indeed, the commonly used method of preparing inocula develop¬ ed by Redfern (1970,1975) ciepends on this very ability. Rishbeth (1984) compared the ability of several isolates to colonize excised stems and roots. His results did not encourage the use of this ability as a measure of viru¬ lence since isolates of A. gallica generally performed better than those of A. ostoyae, a reversal of the normal ranking for pathogenicity. Among workers who have used trees or shrubs for pathogenicity tests, many have chosen to include more than one species because of known or suspected differ¬ ences in susceptibility among potential hosts (Benjamin and Newhook 1984b, Guillaumin and Lung 1985, Guil- laumin and Pierson 1978, Kile 1980b, Morrison 1982b, Mugala and others 1989, Proffer and others 1988, Raabe 1967, Rishbeth 1985b, Shaw and Loopstra 1988). Other investigators have confined themselves to a host in which Armillaria is a current economic problem (Leach 1937, Mallett and Hiratsuka 1988, Ono 1970, Podger and others 1978, Wilbur and others 1972). The type, age, and method of cultivating experimental subjects have differed greatly, but four plant types have been commonly used: very young seedlings grown under laboratory conditions, potted plants, plants in field plots, and established trees. Several attempts have been made to use seedlings un¬ der sterile or near-sterile conditions for laboratory in¬ fection studies (Christensen 1938, Irvine and McNabb 1962, Rayner 1930, Rishbeth 1984, Zollfrank and Hock 1987). In these experiments, infection either hardly occurred at all (Christensen 1938, Rayner 1930, Rishbeth 1984) or was achieved only by growing the seedlings in a culture medium permeated by the fun¬ gus (Irvine and McNabb 1962, Zollfrank and Hock 1987). The symptoms reported in some cases do not resemble those that occur in the field (Rayner 1930, Zollfrank and Hock 1987). Laboratory methods inevita¬ bly limit host size and the type of inoculum that can be used, so results must be considered as bearing little relationship to pathogenesis in vivo. Inoculating young trees in containers (figs. 6.1, 6.2) has, by contrast, provided much valuable information on the infection biology and pathogenicity of Armillaria. In North America, this method contributed to several important studies of '‘Armillaria mellea" (Bliss 1946; Patton and Riker 1959; Raabe 1955,1967,1972; Shaw 1977; Thomas 1934), and it formed the basis of several recent investigations into the pathogenicity of the cur¬ rently recognized North American species (Mallett and Hiratsuka 1988, Morrison 1989, Mugala and others 1989, Proffer and others 1988, Shaw and Loopstra 1988) . European, Asian, and Australasian studies have also made extensive use of container plants (Gregory 1985; Guillaumin and Lung 1985; Guillaumin and Rykowski 1980; Kile 1980b, 1981; Ono 1970; Pearce and others 1986; Podger and others 1978; Redfern 1978; Shaw and others 1980, 1981; Siepmann and Leibiger 1989) . Several workers (Morrison 1989; Ono 1970; Redfern 1975,1978; Pearce and others 1986; Proffer and others 1988) have used several plants per container with each container being treated as a plot. Experimental field plots established in open ground have also been used effectively in Armillaria research. Most experimental data on virulence of European iso¬ lates derive from the field plot inoculations of Rishbeth Figure 6.1 — Inoculation of a ponderosa pine seedling with a branch segment of red alder containing A. ostoyae (see Shaw 1975, 1977). The jar contains inoculum segments on which A ostoyae mycelium is visible as white tufts. (G. M Filip) Pathogenicity and Virulence 79 Figure 6.2 — Two treatments from Redfern's (1975) trial, photographed 18 months after inoculation with European isolates of A. gallica (S3) and A. mellea (S4) in root segments of planetree. Each container originally held 25 young Sitka spruce. 4. mellea (S4) killed all but a few plants in this replicate (treatment total of 61 %), whereas /A. gallica (S3) killed none (less than 5% over the whole experiment). (D. B. Redfern) (1982, 1984,1985a,b) who primarily used 2-year-old Scots pine but also worked with other conifers and a range of hardwood trees and shrubs. Guillaumin and Pierson (1978) used 4- to 5-year-old specimens of peach, Persian walnut, downy oak, and silver fir in field trials in France. In the United States, Wilbur and others (1972) used field plots of peach. One of the few inoculation trials to have been reported for an African Armillnria isolate was conducted in a field plot of com¬ mon tea seedlings by Leach (1937). Relatively few inoculations of established plantation or forest trees have been reported though the hosts for the earliest recorded inoculation were 8-year-old pines in Germany (Hartig 1874). One of the first demonstrations that "Armillaria mellea" exhibited differences in viru¬ lence was achieved by inoculating young plantation pines in the United States (Patton and Riker 1959). Also in the United States, there has been a history of field inoculations in research on A. tabescens (Plakidas 1941, Rhoads 1956, Weaver 1974). The pathogenicity of two other species has been proven by field inoculation. Kile (1981) inoculated young eucalypts with A. luteobubalina in Australia, and Dadant (1963a) inoculated field- grown albizia with A. fuscipes in Madagascar. Large woodland trees have been inoculated in several other studies in which the objective was investigation of host-parasite relationships rather than straightforward testing of pathogenicity (Davidson and Rishbeth 1988, Redfern 1978, Wargo and Houston 1974, Whitney and others 1989b). Inoculating forest or plantation trees could yield data more relevant to field experience than any other method discussed in this section. However, the practi¬ cal difficulties involved are often formidable. Using containers offers ease of handling, flexibility of experi¬ mental design, and greater freedom in environmental control, but conditions in containers, even those as large as Ono (1970) and Redfern (1975,1978) used, can be quite artificial, particularly the rooting environment. Any stress imposed by such conditions could lower host resistance and might thereby obscure differences in virulence between isolates. As noted elsewhere in this chapter, some species of Armillaria with limited ability as primary pathogens can nevertheless act as effective secondary pathogens on weakened trees. Con¬ ditions in containers, such as extremes of soil moisture, also may adversely affect the fungus (Guillaumin and Leprince 1979). Growing conditions in field plots are clearly more natural than those in containers though trees are not necessarily stress-free. Morrison (1982b) mentioned drought stress as a possible factor contributing to high infection in plots established on a sandy soil. Con¬ versely, one cannot assume that artificial or unnatural conditions are always detrimental to the host. Well- tended plants in pots or field plots may be less prone to stress, and hence potentially more resistant, than trees of the same age in some natural situations. Container plants are usually young and are therefore likely to be less resistant to infection and killing than older trees, an obvious drawback to applying results to the field. In most pot trials, experimental plants have been seedlings, cuttings, or transplants 1-4 years old at inoculation. Exceptionally, seedlings only a few weeks old have been used (Entry and others 1986) but results in such cases are likely to have little relevance to field behavior. Field plots offer greater opportunity for us¬ ing older plants, though in many such studies age at inoculation has been 5 years or less (Guillaumin and Pierson 1978; Morrison 1982b; Ono 1970; Rishbeth 1982, 1984; Rykowski 1984). Rishbeth (unpubl.) used a range of isolates and Armill¬ aria species to inoculate, in parallel trials, 1-year-old plants in pots, 2-year-old plants in field plots, and. 7- year-old plantation trees of Corsican pine. Although no true comparison was possible, the data suggest that isolates of low virulence could receive higher ranking from the results of trials with young potted plants than would be justified by other methods, including field observation. Results presented by Proffer and others (1988) are also of interest in this connection. They found uniformly high mortality in cherry (Primus) 80 Pathogenicity ami Virulence seedlings inoculated with one of three Armillaria spe¬ cies including A. gallica, which is normally regarded as an extremely weak pathogen. Quite possibly the isolate used was of exceptionally high virulence, but more likely, the methodology gave a spuriously high result. The hosts were 1-year-old seedlings to which inocula were attached at the time of planting. As the trial ran for only 1 year, both stress of transplantation and the young age of the plants might have increased suscepti¬ bility. The large amount of inoculum used per plant was another possible factor identified by the authors. Choice of Inoculum Garrett's (1956a) development of the concept of inocu¬ lum potential was founded on the recognition that failure to achieve experimental infection with root pathogens was often the result of using unsuitable inocula. As discussed in chapter 4, the inoculum poten¬ tial for Armillaria pathogenesis in vivo is almost exclu¬ sively derived from woody substrates. Accordingly, although successful inoculations of young trees have been achieved with other material, the main experi¬ mental contributions to our knowledge of Armillaria pathogenicity and virulence have been based on the use of woody inocula. Some workers have used naturally infected roots (Kile 1980b, 1981; Leach 1937; Ono 1970; Proffer and others 1988; Rhoads 1938), but these are of limited value for comparative work because of the uncertainty that uni¬ form colonization has been achieved by a single isolate. Most investigators, including some of the earliest to conduct successful inoculation experiments, have used sterilized wood pieces inoculated with pure cultures of the isolates under investigation (Bliss 1946, Guillaumin 1977, Patton and Riker 1959, Podger and others 1978, Raabe 1955, Rishbeth 1984, Shaw 1977, Thomas 1934, van Vloten 1936, Wilbur and others 1972). Some have used inocula prepared in this way to infect live stem or root pieces which have then been used to inoculate the experimental plants (Gregory 1985; Redfern 1970,1975, 1978; Rishbeth 1972b, 1982; Siepmann and Leibiger 1989). This two-stage method has proved advanta¬ geous with some isolates that do not readily produce rhizomorphs from sterilized wood (Redfern 1970). Both methods are time-consuming because inocula take many weeks to become completely colonized, the stage at which they are usually used (Podger and others 1978, Redfern 1975, Shaw 1977). Wilbur and others (1972) incubated inocula for as long as 20 months be¬ fore use. The consequences of using inocula too early have been noted by Benjamin and Newhook (1984b) who found that incompletely colonized inocula did not produce rhizomorphs and that the colonization rate varied greatly among the several types of wood that they tried. Rhizomorph production can be an impor¬ tant factor in achieving artificial infection, as will be discussed below, and it may be influenced directly by the food base used (Azevedo 1970-71; Morrison 1972; Redfern 1970; Rishbeth 1972b; Rykowski 1981c, 1984). Redfern (1975) demonstrated that food base type can affect the amount of experimentally induced disease independently of how it affects the number of rhizomorphs. Choice of wood species for inocula is therefore potentially important for experimentation though the criteria used have rarely been stated. Sev¬ eral authors have used standard hardwood inocula for a range of hosts (Guillaumin and Lung 1985, Guillaumin and Pierson 1978, Raabe 1967, Rishbeth 1984, Shaw 1977, Shaw and others 1981, Siepmann and Leibiger 1989); Pearce and others (1986) used two dif¬ ferent types for each host. Other workers matched in¬ oculum to host (Dadant 1963a, Ono 1970, Podger and others 1978, Proffer and others 1988), or used unrelated species that are a common source of infection in nature (Leach 1937), or material that can be conveniently col¬ lected (Mallett and Hiratsuka 1988). The popularity of hardwood inocula even for coniferous hosts may well reflect the widespread view that hardwoods offer a superior food base for Armillaria species (see chapter 4; Redfern 1970, 1975). The relative merits of using root, branch, or stem wood for inocula have received little attention although the origin could conceivably affect the fungus' pathogenic behavior. Several workers have used root segments (Dadant 1963a, Ono 1970, Patton and Riker 1959, Redfern 1975, Weaver 1974, Wilbur and others 1972), presumably reflecting the most common inoculum source in nature, but many others have achieved worthwhile results with stem or branch material (Gre¬ gory 1985, Guillaumin and Pierson 1978, Kile 1981, Morrison 1982b, Raabe 1967, Rishbeth 1982, Rykowski 1984, Shaw 1977). The size of inocula and their positioning relative to the host have been little discussed despite Garrett's (1956b) early demonstration that both factors affect the ability of rhizomorphs to cause infection. Harrington and others (1989) and Patton and Riker (1959) attributed disappointing results in their field inoculations to under-sized inocula. Size influenced infection in Azevedo's (1970-71) and Rykowski's (1981c, 1984) ex¬ periments with young trees, but the latter still achieved infection of 3-year-old pines with inocula less than 5 cm 3 . Gregory (1985) and Guillaumin and Pierson (1978) conducted successful pathogenicity trials with com¬ paratively small inocula (1.5-2 cm diam x 4-5 cm long) used singly and placed close to the collar or major roots of the host. Other workers have generally used larger inoculum segments and several have used more than one per host. Redfern (1975) used five large segments Pathogenicity and Virulence 81 (2.5-5.5 cm diam x 10 cm long) in each tub (30 cm diam) of 25 small conifers, whereas Davidson and Rishbeth (1988) used similarly sized inoculum segments singly to attempt inoculation of 32-year-old oak trees. Leach (1937) used a massive amount of inoculum to establish infection: large pieces of infected root were placed in a layer through which the roots of tea seedlings were allowed to grow. More recently Proffer and others (1988) used extremely large inocula relative to the size of the host: three stem segments 1.2 cm diam x 12-13 cm long were attached to the collar (approx 1 cm diam) of each 1-year-old experimental plant. As mentioned previously, the experiment gave unusually high levels of disease, an outcome which may have been partly due to the high inoculum potential resulting from the inoculation method. As well as helping to increase inoculum potential, plac¬ ing inocula close to the host may help to prevent dis¬ ease escape. Rishbeth (1984), although working in an area and with species in which infection by rhizomorphs is probably the norm, considered it im¬ portant to place inocula in contact with the host to al¬ low the opportunity for non-rhizomorphic infection by isolates which are poor rhizomorph producers. In stud¬ ies of species such as A. tabescens and A. fuscipes that normally infect only through root contacts, inocula are necessarily placed in contact with the host (Dadant 1963a, Plakidas 1941, Rhoads 1938, Rishbeth 1985b, Weaver 1974). In some experiments with A. tabescens and A. fuscipes artificial wounds have been made at the point of inocu¬ lation (Dadant 1963a, Plakidas 1941, Weaver 1974). Wound inoculation has not commonly been employed with other species though Whitney and others (1989b) reported that such inoculations with A. ostoyae on fir roots were more successful than non-wound inoculations. Rhizomorphs and Measurement of Disease in Inoculation Trials Assessing virulence in trials has commonly been based on one or more of the following: amount of root infec¬ tion, amount of mortality, or rapidity of infection or mortality. Such relatively straightforward measure¬ ments are, however, often complicated by the need to consider the role of rhizomorphs as extensions of the experimental inoculum. Serious ArmiUaria diseases occur in several regions of the world where rhizomorph growth is restricted or absent (Dadant 1963a, Dade 1927, Kile 1981, Morrison 1981, Podger and others 1978, Rhoads 1956, Rishbeth 1980). Although non-rhizomorphic infection occurred commonly in Kile's (1981) inoculation trials with A. luteobubalina and Dadant's (1963a) with A. fuscipes, it has proved difficult to achieve experimentally with A. tabescens, the other economically important species known to infect in this way naturally (Rhoads 1956, Weaver 1974). Non-rhizomorphic infections by temper¬ ate species have occasionally been observed in inocula¬ tion trials (Rishbeth unpubl., Shaw 1977, Whitney and others 1989b) but some attempts to induce them delib¬ erately have failed (Redfern 1978). Most ArmiUaria experimentation has involved species in which rhizomorphs have been assumed to be the normal, or only, means of infection; most inoculation experiments have included a final assessment of the presence or absence of rhizomorphs. Among these studies are several reports of isolates which do not produce rhizomorphs readily, or at all, under experi¬ mental conditions (Gregory 1985; Mallett and Hiratsuka 1988; Rishbeth 1984; Rykowski 1981c, 1984; van Vloten 1936). Such isolates may eventually pro¬ duce rhizomorphs given time (Patton and Riker 1959, Gregory unpubl.) or may be induced to do so by alter¬ ing the method of inoculum production (Redfern 1970, Rishbeth 1968) or inoculum size (Benjamin and Newhook 1984b). Rhizomorph production, and hence disease, may also be strongly influenced by soil condi¬ tions (see chapter 4). Consequently, it may be difficult to decide whether lack of rhizomorphs, which is usu¬ ally associated with lack of infection, reflects genuine field behavior or defective technique. Interpreting results can be difficult in experiments where inocula in some replications produce rhizomorphs while those in others do not. Gregory (1985) treated such replicates as missing values, and Morrison's (1982b) scoring system also excluded repli¬ cates in which no rhizomorph contacted the host. How¬ ever, some authors have included these data among non-infected categories, accepting the risk that this might distort results of trials with species that are poor rhizomorph producers. Some of the problems associated with rhizomorph behavior are represented in the data of Mallett and Hiratsuka (1988), who found low disease levels and no rhizomorphs in trials with Canadian isolates of A. ostoyae. Since other evidence (discussed below) sug¬ gests that this species is a serious pathogen in both North America and Europe, the few infections achieved probably resulted not from low intrinsic pathogenicity but rather from the species' inability to produce rhizomorphs under the experimental condi¬ tions. European isolates of the same species have been characterized by Guillaumin and others (1985) and Gregory (1985) as poor producers of rhizomorphs in experiments. 82 Pathogenicity and Virulence Rhizomorph production may have a bearing on the optimum duration of Armillaria inoculation trials, a subject which has been briefly discussed by some au¬ thors (Benjamin and Newhook 1984b, Gregory 1985, Mallett and Hiratsuka 1988, Patton and Riker 1959), but which merits further attention. Several studies indicate that certain isolates take longer than others to cause visible, above-ground signs of infection (Gregory 1985, Raabe 1967, Redfern 1975, Rishbeth 1984, Wilbur and others 1972). The three isolates used by Wilbur and others (1972) differed little in virulence assessed simply as the proportion of experimental plants killed at the end of a 3-year trial. They would have been judged to differ markedly from each other, to the extent of one being almost non-virulent, had the final assessment been made after 18 months. Yet, this timespan equals or exceeds that chosen by many workers. In an unpub¬ lished trial using methods similar to those of Redfern (1975), Gregory found that 2 years after inoculation A. mellea had killed twice as many young conifers as A. ostoyae; however, after 3 years the position was re¬ versed and was maintained until the trial ended 5 years after inoculation. If a relatively slow rate of disease development reflects a relatively poorer ability of rhizomorphs contacting a host to initiate infection, then it may be a legitimate expression of lower virulence as some authors have proposed (Raabe 1967, Rishbeth 1984). If, by contrast, experimental manipulation adversely affects rhizomorph production and subsequently causes slow disease development, then the use of rate in compara¬ tive assessments is questionable. Guillaumin and oth¬ ers (1985) have noted that European species differ in the time taken to produce rhizomorphs under experi¬ mental conditions. They cite A. ostoyae as being espe¬ cially tardy, an observation that coincides with unpublished data of Gregory and Rishbeth. Most investigators who have studied pathogenicity in Armillaria have measured the amount of disease simply by the proportion of host plants killed or infected dur¬ ing the experiment. Several authors have used lesion size for scoring the severity of non-fatal infections (Gre¬ gory 1985, Guillaumin and Pierson 1978, Morrison 1982b, Rishbeth 1982). Assessments of dead or symp¬ tomatic plants have nearly always been visual and involved destructive examination. The main exception to the latter is a study by Zollfrank and Hock (1987), who conducted their experiments under aseptic condi¬ tions and used immunofluorescence to detect hyphae in seedling tissues. Field Observation The century-old descriptions of Armillaria disease by Robert Hartig (1874,1894) reveal the field experience of a remarkable observer and stand comparison with modern accounts. From this beginning, field observa¬ tions have been a major source of information about Armillaria disease, but they have also fueled much con¬ troversy over the role of Armillaria as a pathogen. Armillaria diseases are probably almost as difficult to observe critically in the field as they are to investigate by experiment. Worthwhile field observations require a comprehensive knowledge of forest pathology and of Armillaria biology as well as meticulous site investiga¬ tion. Regrettably, some studies assume that the situa¬ tions from which basidiomes have been collected fully circumscribe the ecology and pathogenic behavior of the fungus. With our present ability to identify separate species of Armillaria, field observation has contributed significant information about pathogenicity. Despite Rishbeth's extensive experimental work, an appreciable propor¬ tion of our knowledge of pathogenicity in the Euro¬ pean species derives from field observations (Gregory 1989; Guillaumin and others 1985; Guillaumin and Berthelay 1981; Korhonen 1978; Rishbeth 1982, 1984, 1985b). Field observations, notably those of Morrison and others (1985a), also constitute a major source of published data on North American species. In New Zealand and Australian studies, inoculation trials have complemented extensive field observations (Kile 1980b, 1981; Kile and Watling 1983; Pearce and others 1986; Podger and others 1978; Shaw and others 1981). Indirect Methods of Assessing Virulence Attempts to assess virulence indirectly have had only limited success. The idea of a direct relationship be¬ tween virulence and host may be traced back to the observations Childs and Zeller (1929) made on what appeared to be a virulent "oak strain" of the pathogen and a non-virulent "fir strain." They were careful to acknowledge the danger of extrapolating their observa¬ tions to other regions, but the idea of a link between host and virulence has persisted. However, despite having been investigated experimentally a number of times, no such connection has been demonstrated (Guillaumin and Pierson 1978, Raabe 1967, van Vloten 1936). Possible relationships between virulence and the capac¬ ity to produce rhizomorphs have also received consid¬ erable attention. The apparent reliance on rhizomorphs for infection was taken by van Vloten (1936) to indicate that isolates which appeared to lack them were de facto non-virulent. Rykowski (1981c, 1984) observed good agreement between infection and rhizomorph produc¬ tion in his numerous experiments and used the relative abundance of rhizomorph growing tips as an index of "infection threat" in his three isolates, all of which Pathogenicity and Virulence 83 belonged to A. ostoyae. Some other studies involving single isolates or several isolates of the same species suggested a positive relationship between infection and rhizomorph production (Azevedo 1970-71, Guillaumin and others 1989a, Shaw 1977), but studies involving several isolates of widely different virulence have gen¬ erally failed to demonstrate such a relationship (Guillaumin and Pierson 1978, Raabe 1967, Rishbeth 1984). Conversely, some evidence indicates a negative correlation of rhizomorph production to pathogenicity among European species (Gregory 1985, Red fern 1975, Rishbeth 1985b). Morrison (1972,1982b) and Redfern (1975) suggested an association between dichotomous branching of rhizomorphs and high virulence. The same authors also noted that highly virulent isolates tended to pos¬ sess fragile rhizomorphs. We now know that Morrison's (1982b) three branching types represented three different species (Morrison 1989) and that Redfern's (1975) four isolates were also from four spe¬ cies (Gregory 1985). Later studies (Guillaumin and others 1985, Morrison 1989, Rishbeth 1982) have con¬ firmed that branching habit and fragility of rhizomorphs are species characteristics. Morrison's (1989) data, drawn from 15 species, showed that a di¬ chotomous branching habit (fig. 4.1) more often than not accompanied high pathogenicity but the associa¬ tion was not invariable. Three of the eight dichoto- mously branching species which he tested were of low pathogenicity. It may be unrealistic to seek universal relationships between growth patterns and pathogenic¬ ity among species that have evolved to survive in such widely different forest and soil conditions as have the various Armillaria species. A few attempts have been made to assay virulence in Armillaria by in vitro characters. The most notable were based on the work of Wargo (1981 d) that indicated a link between gallic acid metabolism and virulence in certain North American isolates. Shaw (1984,1985) tested this hypothesis extensively on a collection of 72 isolates drawn from three continents. He found that although the ability to tolerate gallic acid varied among isolates, differences could not be utilized consistently as markers for virulence. Rishbeth (1986) reached a similar conclusion. Differences in Pathogenicity and Virulence Although taxonomists have for decades postulated the xistence of several morphological species of Armillaria V' chapter 1), the recognition by pathologists of dis- v, i iogens has been comparatively recent. Two ceptions were provided by A. tabescens, which was accepted as a pathogen in its own right in the southern United States in the 1940's, and A.fuscipes, which Dadant (1963a) demonstrated to be a root patho¬ gen of woody plants in Madagascar. Otherwise, before the late 1970's forest pathologists generally referred attacks of Armillaria disease to a single but variable taxon with worldwide distribution, "Armillaria mellea." Some older data on the pathogenicity of "Armillaria mellea" have been reinterpreted relative to current taxa, but much information from before 1970 is of limited value. Modern studies of pathogenicity and virulence have concentrated largely upon North American, Euro¬ pean, and Australasian isolates. Outside these regions, pathogenic species of Armillaria undoubtedly exist (see chapter 9), but little is known about the variation among them. European and North American Species Although forming a rather artificial grouping, these species are considered together because at least three, including the major pathogens A. mellea and A. ostoyae, appear to be common to both continents. Evidence from inoculation trials identifies A. mellea as probably the most pathogenic species in this group. In Europe, isolates of this species have not only consis¬ tently been ranked highest in comparative studies but have also been demonstrated to cause disease in genera normally regarded as highly resistant to Armillaria (Davidson and Rishbeth 1988; Gregory 1985; Guillaumin and Pierson 1978*; Morrison 1982b*; Redfern 1975*; Rishbeth 1982,1984). Three Canadian trials have included European isolates of A. mellea alongside North American isolates of other species, and in all cases the former have proved the most viru¬ lent (Mallett and Hiratsuka 1988; Morrison 1989, and pers. comm.; Mugala and others 1989). However, the results of inoculation experiments done by Guillaumin and Lung (1985) suggest that A. mellea may be less pathogenic than A. ostoyae to some conifers, an out¬ come which the authors interpreted as evidence of host specialization. Field observations in Europe indicate that A. mellea is the most pathogenic species on ornamental trees, or¬ chard crops, and vines (Guillaumin and Berthelav 1981; Guillaumin and others 1985; Intini 1988; Rishbeth 1982, 1985a). Even though it often kills ornamental conifers, and some isolates are extremely virulent toward young conifers in experiments, it is not widely associated with disease in forest or plantation conifers. In the United "Isolates in Redfern (1975) were identified by Gregory (1985); those in Guillaumin and Pierson (1978) were identified by Guillaumin and Berthelav; those in Shaw (1977) were identified by Shaw (1984) and those in Morrison (1982b) were identified by Morrison (1989). 84 Pathogenicity and Virulence States, Proffer and others (1987,1988) found that A. mellea was associated with root disease of cherry in Michigan, but few other observations on North Ameri¬ can isolates involve this species. In inoculation trials. North American and European isolates of A. ostoyae have generally been moderately or highly virulent towards young conifers (Gregory 1985; Guillaumin and Lung 1985; Morrison 1982b, 1989; Redfern 1975; Rishbeth 1982,1984,1985b; Shaw 1977; Siepmann and Leibiger 1989). Under experimental conditions, the species appears to be only weakly pathogenic to European forest hardwoods (Lung- Escarmant and Taris 1989, Rishbeth 1984). Rishbeth's (1984) data suggest that A. ostoyae could be classed with A. gallica as virtually non-pathogenic to common ash and silver birch although the isolates of A. ostoyae used were highly virulent to Scots pine in the same trial. Proffer and others (1988) found Michigan isolates of A. ostoyae to be highly virulent to Primus species, but in¬ terpreting their results requires caution because of the equally high disease levels achieved with A. gallica isolates in the same experiments. Possible reasons for this have been discussed earlier in this chapter. Isolates of A. ostoyae showing low virulence towards conifers have been reported in Europe (Rishbeth 1984), and recently, Mallett and Hiratsuka (1988) demon¬ strated apparently uniform low virulence toward young lodgepole pines in a range of Canadian isolates. As suggested earlier, such results may reflect the poor ability of some isolates to produce rhizomorphs under experimental conditions rather than innate low viru¬ lence. Indeed, A. ostoyae may be consistently under¬ rated in inoculation studies for this reason. Field observations in North America (Bloomberg and Morrison 1989, Dumas 1988, Harrington and others 1989, Mallett and Hiratsuka 1988, Morrison and others 1985a), Fenno-Scandia (Korhonen 1978, Piri and others 1990), and Europe (Gregory 1989, Guillaumin and Berthelay 1981, Guillaumin and others 1985, Intini 1988, Rishbeth 1985a, Siepmann 1985) indicate that A. ostoyae is a major forest pathogen of conifers in those regions. Several of these accounts show the species can kill trees of all ages and can also cause butt rot in older crops. So consistently has A. ostoyae been associated with disease in conifers that it is commonly assumed to be the probable pathogen whenever serious Armillaria disease is encountered in North American or European coniferous forests (Filip 1989a, Hadfield and others 1986, Hansen and Goheen 1989, Rizzo and Harrington 1988a, Whitney 1988b). Despite the low pathogenicity towards hardwoods indicated by inoculation experiments, field observa¬ tions suggest that A. ostoyae is capable of attacking broadleaved trees and shrubs growing within diseased conifer stands (Guillaumin and others 1985, Morrison and others 1985a, Rishbeth 1985a). Harrington and others (1989) recorded it as a cause of death of birch and maple in the northeastern United States, and the observations made in Canada by Dumas (1988) suggest that it may have a rather wide host range among hard¬ woods there, at least as a secondary pathogen. Armillaria gallica has been widely categorized as a weak pathogen by both field observations and inoculations in Europe and North America (Gregory 1985,1989; Guillaumin and Berthelay 1981; Guillaumin and Pierson 1978; Guillaumin and others 1985; Morrison 1989; Morrison and others 1985a; Redfern 1975; Rishbeth 1982,1984; Shaw 1977, 1984; Siepmann and Leibiger 1989). Some isolates have been designated as virtually non-virulent (Rishbeth 1982, Shaw 1984), yet in few trials has this species completely failed to cause disease. In some cases (Guillaumin and Pierson 1978, Proffer and others 1988), it has caused appreciable damage, albeit to highly susceptible species. As already discussed, the level of damage achieved in trials with young, and possibly stressed, experimental plants may be artificially high. However, since newly planted crop or ornamental trees are also often young and stressed, it might therefore be unwise to dismiss comparatively weak pathogens such as A. gallica as harmless. More¬ over, the ability of A. gallica to act as a secondary agent of mortality in large trees and to cause root- and butt- rot in live trees (Gregory 1985, Rishbeth 1982) implies a far from negligible capacity to cause disease. These remarks apply to most of the other species categorized below as weak pathogens. Armillaria cepistipes is regarded in Europe as an ana¬ logue of A. gallica: a weak pathogen virtually indistin¬ guishable from A. gallica in behavior and appearance (Guillaumin and others 1985). Few inoculation trials have been reported for this species, but those of Redfern (1975) and Morrison (1989) both indicated low virulence in the isolates tested. Rishbeth has unpub¬ lished data from the trials reported in 1985b, table 3, that also demonstrate low virulence. The species is nevertheless associated with butt rot of conifers in Fin¬ land and Scotland (Gregory 1989, Korhonen 1978, Piri and others 1990). Of the six North American biological species (NABS) not clearly identified with European species (A. gemina, A. calvescens, A. sinapina, NABS IX, NABS X, and NABS XI), A. sinapina (NABS V) has probably received most attention because it is relatively common in some im¬ portant forest areas (Mallett and Hiratsuka 1988, Morrison and others 1985a, Shaw and Loopstra 1988). The inoculation experiments with young trees in con¬ tainers carried out by Morrison (1989), Mugala and Pathogenicity and Virulence 85 others (1989), and Shaw and Loopstra (1988) suggest that the species is of low pathogenicity towards some North American conifers. However, in another trial, Mallett and Hiratsuka (1988) found more infection caused in potted lodgepole pine by Canadian isolates of A. sinapina than by A. ostoyae. Although, as noted above, the latter may have been seriously underesti¬ mated in this experiment, the data for A. sinapina are nonetheless anomalous, the more so as Mugala and others (1989), using similar methods, reported low virulence towards the same host by one of the same isolates. Field observations in Canada support the view that A. sinapina is a weak pathogen (Dumas 1988, Morrison and others 1985a). NABS IX also appears to have limited pathogenicity towards young conifers according to field observations and inoculation trials in British Columbia and Alaska (Morrison and others 1985a, Morrison 1989, Shaw and Loopstra 1988). Shaw and Loopstra (1988) found that haploid isolates of this species and A. sinapina caused significantly more disease than the parent isolates. The observations of Morrison and others (1985a) in British Columbia placed the other northwestern spe¬ cies, NABS XI, in the same category as A. gallica, A. sinapina, and NABS IX. All are weak pathogens charac¬ teristic of suppressed or overmature hardwoods. The results of Morrison's (1989) subsequent inoculation trial with young Douglas-fir in pots supported this view since all four species received the same very low rating. Armillaria gemina (NABS II) and A. calvescens (NABS III) were also included in Morrison's (1989) trial. Both were accorded the same low rating as NABS IX, NABS XI, A. sinapina, and A. gallica. Despite this, field observations on A. calvescens by Proffer and others (1987) in Michi¬ gan and by Harrington and others (1989) in New Hampshire associate it with root rot and mortality of hardwoods. In the case of A. gemina, Berube and Dessureault (1989) have stated that it is "identical to A. ostoyae in terms of . . . pathogenicity," but this view is based on extremely limited field observations. Little is known about the pathogenicity of NABS X, although McDonald (1990) suggests, again from limited observa¬ tions, that it may be moderately pathogenic. The northern European species A. borealis, which has not been recorded in North America, is generally re¬ garded as a rather weak pathogen (Guillaumin and others 1985, Korhonen 1978), though observations from Britain suggest that some genotypes might be virulent to young conifers (Gregory 1989). Korhonen (1978) identified A. borealis as an important cause of butt rot of Norway spruce in Finland, and it has been associated with similar damage in Germany and Britain (Gregory 1989, Siepmann 1985). Only two inoculation trials have been reported. Both utilized young potted conifers; and both suggested that A. borealis is a rather weak patho¬ gen, intermediate between A. mellea and A. ostoyae on one hand and A. gallica and A. cepistipes on the other (Morrison 1989, Siepmann and Leibiger 1989). Although A. tabescens has been cited as causing root disease in trees in several parts of the world, consider¬ able doubt now exists that a single species is involved (see chapter 1). Most information is available from the southern United States, where it is known as a serious pathogen of ornamental trees and commercial crops (Rhoads 1956, Sinclair and others 1987). The fungus can attack a wide range of woody species in a variety of genera but, according to Rhoads (1956), exotics are much more susceptible than native trees and shrubs. Rhoads (1956) also reported that damage caused by A. tabescens was particularly prevalent on drought-prone sites, and Weaver (1974) suggested that disease in peach only followed infection of previously killed or injured roots. Other reports associate A. tabescens with disease in stressed trees or trees primarily attacked by other agents (Filer and McCracken 1969, Ross and Marx 1972, Sinclair and others 1987). A fungus referred to as A. tabescens has also been re¬ corded in southern Europe as a root pathogen on sev¬ eral hosts including citrus on Corsica (Laville and Vogel 1984), eucalypts in southwestern France (Lung- Escarmant and others 1985a), and cork oak in Portugal (Azevedo 1976). Further north, European isolates of A. tabescens appear to be non-virulent in the sense of caus¬ ing root mortality, though field observations have linked the species with decay of live trees in Britain (Rishbeth 1984,1985b). The apparently southern distri¬ bution of diseases attributed to A. tabescens in both Europe and the United States is noteworthy because observations in China (Chang and others 1982) also associate severe root infection by A. tabescens with high soil temperature. Australasian Species Three Australasian species are regarded as serious pathogens on the evidence of field observation and inoculation trials: A. luteobubalina, A. novae-zelandiae , and A. limonea. Isolates of all three were represented in Morrison's (1989) trial which tested a range of Euro¬ pean, North American, and Australasian species against 2-year-old Douglas-fir seedlings in containers. His results suggested that the three Australasian patho¬ gens may be ranked with A. mellea and A. ostoyae. How¬ ever, the amounts of disease recorded in most Australasian trials have been low by comparison to European or North American results with A. mellea and A. ostoyae. The contrast is particularly noteworthy in 86 Pathogenicity and Virulence similar tests on radiata pine conducted by Shaw (1977) in the United States with A. gallica and A. ostoyae and by Shaw and others (1980,1981) in New Zealand with A. novae-zelandiae and A. limonea). Field observations in New Zealand by MacKenzie and Shaw (1977) and Shaw and Calderon (1977) attributed disease in radiata pine crops to two native Armillaria species, A. novae-zelandiae and A. limonea , with the for¬ mer appearing to be the more serious pathogen. Inocu¬ lation trials with young radiata pine in containers (Shaw and others 1980, 1981) demonstrated that both species were moderately pathogenic although some isolates of each had low virulence. Benjamin and New- hook (1984b) undertook trials with the same two spe¬ cies and found them highly pathogenic toward radiata pine, but in tests with eucalypts, A. limonea seemed to be less pathogenic than A. novae-zelandiae. Armillaria novae-zelandiae also occurs in Australia where Kile and Watling (1983) recorded it as a secondary pathogen of native trees and a frequent cause of decay in myrtle beech. More recently, it has been cited by Kile and Watling (1988) as causing localized losses in young crops of exotic conifers, in which it is linked with A. fumosa and A. pallidula. Little else is known about either of these species though an isolate of A. fumosa was in¬ cluded in Morrison's (1989) trial in which it proved virtually non-virulent. The chief Australian pathogen is undoubtedly A. luteobubalina. Field observations (Kile 1981, Kile and others 1983, Pearce and others 1986, Podger and others 1978, Shearer and Tippett 1988) have repeatedly dem¬ onstrated that it is a major primary pathogen in native sclerophyll forests where it kills eucalypts and a wide range of understory trees and shrubs. Infection can occur on eucalypts of all ages, resulting in crown die- back or mortality of large overstory trees as well as serious losses among seedlings and saplings. The fun¬ gus also attacks a wide range of species in vineyards, orchards, and ornamental plantings (Kile and Watling 1988). Armillaria hinnulea by contrast was found to be weakly pathogenic in inoculation experiments with both native species and North American conifers (Kile 1980b, Morrison 1989). Morrison's (1989) data indicate that this species is similar to the European A. borealis in its ability to infect young Douglas-fir in containers. Field observations have characterized A. hinnulea as a weak pathogen capable of causing localized root lesions and decay in resistant hosts. It is nevertheless an effective secondary pathogen, and in this capacity, it is of some economic importance in Tasmania through association with "regrowth dieback," a decline of eucalypts of which the primary cause is unknown (Kile 1980b, Kile and Watling 1983). Non-Australasian Tropical and Subtropical Species Dadant (1963a) demonstrated experimentally that the morphological species he knew as A. elegans was pathogenic to field-grown albizia sp. His detailed ob¬ servations and numerous isolations leave little doubt that the fungus he studied is a serious pathogen of coffee bushes and shade trees in Madagascar. Blaha (1978) associated the same fungus with damage to a similar range of hosts in Cameroon. The fungus is now known to occur widely in Africa and to be conspecific with A. fuscipes (see chapter 1), which was described by Petch (1923) as a root pathogen of acacia and probably also of tea bushes in Sri Lanka. Most of the numerous accounts of Armillaria diseases in tropical and subtropical crops (see chapter 9) cannot now be validly attributed to morphological or biologi¬ cal species. However, the recent work by Mohammed and others (1989) with African isolates suggests that other pathogenic species in addition to A. fuscipes occur on that continent. Ironically, one of these appears to be at least partially interfertile with A. mellea —the name associated by default with disease in Africa since the early years of this century. Conclusions The genus Armillaria contains several virulent patho¬ gens and other species that have evolved as successful secondary or facultative pathogens. Failure to appreci¬ ate this variation within the genus probably accounts for much of the controversy that has arisen in the past over the pathogenic status of Armillaria. Without doubt some species are primary pathogens, though the amount of disease caused by even the most pathogenic taxa may be conditional upon the nature of the host and the environment of the fungus. Most species ap¬ pear to have a wide host range, but some species are apparently adapted to particular groups of hosts or site conditions or both. There is strong evidence that viru¬ lence differs among isolates of some species. Experimentation with Armillaria poses formidable problems, and the interpretation of data from experi¬ ments and field observations is rarely straightforward. Nevertheless, our understanding has advanced re¬ markably rapidly in the past 20 years, though many aspects of pathogenicity merit further investigation. Despite the advances, relatively little is known about several North American biological species and even less about tropical and subtropical species. Pathogenicity and Virulence 87 CHAPTER 7 Host Stress and Susceptibility Philip M. Wargo and Thomas C. Harrington rmillaria root disease has historically been considered a disease of weakened trees. Early observers indicated that Armillaria was secondary to some other factor that predisposed trees to attack (Day 1927a, 1928,1929; Falck 1918,1923; Muller 1921; Nechleba 1915,1927; Thomas 1934). Although not always the case, predispo¬ sition is considered common with Armillaria root dis¬ ease, and seems to be more important in this disease than in the other woody root diseases of forest, shade, and orchard trees. As with all diseases, susceptibility to Armillaria root disease depends on interactions among host, pathogen, and the environment. The importance of predisposing stresses and their impact on host vigor (the environ¬ mental component) must be considered in the context of the host and the pathogen. Armillaria has an extremely broad host range (Raabe 1962a), but these hosts vary in their susceptibility. Furthermore, many species of Armillaria are now recognized and these vary greatly in their pathogenicity (see chapter 6). Some are primary pathogens capable of killing vigor¬ ous hosts while others colonize only severely stressed individuals. Stresses generally predispose trees to Armillaria root disease by reducing host vigor and, thus, compromis¬ ing host defenses. Host defense mechanisms are ad¬ dressed in chapters 4 and 5, but a brief review will set the stage for our discussion of stress and predisposi¬ tion. Chronic and acute stresses and how they might affect resistance are covered in general, and specific examples of abiotic and biotic stress agents known to predispose trees to Armillaria are given. Lastly, we discuss forest management of Armillaria root disease relative to stress-induced susceptibility. Stress Concepts and Host-Pathogen Interaction Variation Among Armillaria Species, Host, and Site Confusion about Armillaria taxonomy has hampered our understanding of stress effects on disease develop¬ ment. Unfortunately, very little research on stress-in¬ duced susceptibility has been conducted with known species of Armillaria. Where species of Armillaria have been identified, evidence suggests that root disease caused by A. mellea, A. ostoyae, or A. gallica is more likely to occur in a stressed host (Davidson and Rishbeth 1988). Obviously, variation in pathogenicity among the Armillaria species (see chapter 6) has an important bear¬ ing on the requirement for a predisposing stress in disease development. Armillaria gallica only attacks stressed trees (Davidson and Rishbeth 1988) whereas A. mellea and A. ostoyae can infect and kill apparently vigorous trees. Stress may also broaden the host range of some Armillaria species. For example, A. ostoyae at¬ tacks primarily conifers but will also attack oaks when they are stressed (Davidson and Rishbeth 1988). Predisposing stresses may be more important for dis¬ ease development in relatively resistant species than in the more susceptible species. In general, hardwoods are considered more resistant to Armillaria root disease than coniferous species in northern temperate forests (Redfern 1978, Rishbeth 1972a). As discussed later, predisposing factors have been more often noted in Armillaria root disease on hardwoods than on conifers. However, Armillaria may be equally aggressive on healthy hardwoods, and this observation may reflect 88 Host Stress and Susceptibility the limited distribution of A. mellea, the species most capable of colonizing apparently healthy hardwoods (Davidson and Rishbeth 1988, Rishbeth 1982). Also, research on root and butt rots in hardwoods has been limited, and the disease may be more prevalent on hardwoods than commonly realized (Nordin 1954, Shigo and Tippett 1981). Very limited information is available on resistance among hardwood species, but work on rootstocks of horticultural species shows that resistance varies both among and within species. Thomas and others (1948) reported that pear and walnut were quite resistant to Armillaria, but apricot and prune were susceptible. Variation in root stock resistance among several Primus species was also reported in France (Guillaumin and Pierson 1983). Both studies demonstrated that peach and apricot root stocks were more susceptible to Armillaria than plum root stocks. Recent work by Guillaumin and others (1989b) verified that this rela¬ tionship exists for A. mellea sensu stricto. The resistance of plum species was a dominant trait, and resistance to infection and colonization was maintained in some plum x peach hybrids. Armillaria root disease; occurs on many coniferous species (Raabe 1962a), but resistance varies consider¬ ably among and within species. In an English forest where Scots pine and Norway spruce were growing together, large patches of pine were killed while spruce were unaffected (Rishbeth 1972a). Inoculation studies on small trees, comparing resistance between conifers and hardwoods, showed that large differences existed among tree species in the percent of trees infected by Armillaria, and in the ratio of killed trees to surviving- infected trees; the hardwood species were generally the most resistant (Redfern 1978). Morquer and Touvet (1972b) also noted variation in resistance among conifer species, but no species tested was immune to infection. Differences in resistance clearly occur within and among host species, but much of this observed differ¬ ence may be related more to tree vigor than to genetic resistance. The importance of tree vigor in Armillaria root disease and the interplay of vigor and resistance make ranking of species susceptibility difficult, even with inoculation data (see chapter 6). Likewise, unless clonal material is available, identifying the importance of stresses and tree vigor is difficult. Site factors and host adaptation play an important role in host vigor and susceptibility to Armillaria root dis¬ ease. McDonald and others (1987a) found that the inci¬ dence of pathogenic Armillaria was low in habitat series of high productivity, unless the site was disturbed. In habitat series of low productivity, Armillaria was patho¬ genic in both disturbed and pristine sites. Disturbance was associated with increased disease incidence, but the association was weaker in highly productive sites where adaptive tolerances of the tree species were not exceeded. They suggested that Armillaria root disease was a problem on conifers in sites affected by human activities (including fire suppression), insects, or dis¬ eases, and in pristine sites where tree species were not adapted physiologically to their environment. While little experimental work has been done to test this hypothesis, observations on where Armillaria is or has been a problem in forest stands tend to support it. For example, in the Northwestern United States Armillaria problems often occur in off-site plantations (Hadfield and others 1986, U.S. Dept. Agric. 1983) or transition forests that have been perpetuated by fire and disturbed recently by logging activity and fire control (Shaw and others 1976a). Problems with exotic species can also be related to maladaptation. Although these species may grow very well in new regions, they may not be well adapted to the climatic extremes in their new habitat. Consider, for example, radiata pine in high rainfall areas in New Zealand (Hawkins and Sweet 1989a,b). The factors important to site adaptation and tolerance of climatic extremes, including such physiological processes and conditions as net photo¬ synthesis, cold and drought tolerance, and genetic vari¬ ability, are also related to resistance to Armillaria root disease. Host Vigor and Predisposition The term "vigor" has been used to describe the overall robustness of a tree as indicated by its relative growth and absence of signs and symptoms of disease. Vigor is determined by a tree's physiological performance within a particular environment, and this performance depends upon the tree's genetic capacity. Genetic variation gives a range of physiological performances and therefore a range of physiological conditions or tree vigors under a given set of environmental condi¬ tions. Crown position (dominant, intermediate, or sup¬ pressed) and crown condition (good, fair, or poor) are commonly used to classify tree vigor. These are good indices of a tree's past relative growth and general vigor. However, they indicate little about a tree's cur¬ rent health and its vulnerability to the effects of stress (Wargo 1978a,b,c). When stressed by defoliation, for example, trees in all of the above vigor categories may be attacked and killed by Armillaria (Wargo 1977), indi- Host Stress and Susceptibility 89 eating that within these general vigor categories there are gradations of tree health. Herein, host vigor refers to the tree's current health and vulnerability. Yarwood (1976) defines predispostion as "... the ten¬ dency of treatments and conditions acting before in¬ oculation or before the introduction of the incitant, to affect susceptibility to biotic and abiotic pathogens." In the strict sense of this definition, trees are not always predisposed to infection by Armillaria since the patho¬ gen may have already infected the roots prior to the stress. Many observations, especially in the Armillaria - hardwood relationship, suggest that for some combina¬ tions of hosts and Armillaria species the fungus rarely infects and colonizes an unstressed tree despite epi¬ phytic pathogen growth on root surfaces (see chapter 8). Yarwood's broader definition of predisposition also includes changes that induce greater resistance to dis¬ ease; however, only examples of increased susceptibil¬ ity are emphasized in this chapter. Predisposition to disease may play a much larger role in pathogenesis of forest-tree species than in other plant types because of their longevity. During the lifespan of a tree, it may be exposed to numerous stress-inducing episodes ranging from mild to acute and from short-term to chronic. Also, stresses that were inconsequential during a tree's early years can have devastating effects on the tree later. As trees increase in size and completely occupy their sites, their ability to maintain adequate moisture, nutrients, and energy levels approaches the physical limitations of the root and shoot systems; stresses can then cause consider¬ ably more damage. Resistance to pathogenic organisms is the rule rather than the exception in forest trees. "If this were not so, they [trees] would have ceased to exist," (Shain 1968); or at least they would not live as long as they do. Al¬ though all trees have some capacity to resist infection, this resistance requires substantial energy. This meta¬ bolic energy is necessary to maintain or synthesize structural or chemical defenses that influence growth of pathogens at the surface of the plant or internally (Wood 1967). Production of physical and chemical barriers depletes the host's energy reserves, and trees of less than optimal vigor may not have the energy reserves required to resist infection and are therefore predisposed to disease. Conversely, host species with little genetic resistance will succumb if the pathogen is present, regardless of their energy reserves. Stresses and Resistance to Armillaria The term "stress" has been used to describe any envi¬ ronmental "actor that can have potentially unfavorable influences on living organisms. Levitt (1972) defines "biological stress" as "any environmental factor ca¬ pable of inducing a potentially injurious strain in living organisms" and "biological strain" as any change pro¬ duced by the stress. The strain may be physical, such as the reduction of water flow through the transpiration stream in trees, or it may be chemical, such as a shift in carbohydrate metabolism. Chronic and acute stresses may disturb plants by alter¬ ing resource allocation or by interfering with sink- source relationships (Waring and Patrick 1975). Stresses may interfere with the resistance response by reducing the energy reserves available for reaction (McLaughlin and Shriner 1980). Acute stresses may also temporarily impede metabolism at the infection site, and thus compromise the resistance response. The effects of a particular stress depend on severity, dura¬ tion, season, frequency of occurrence, and the condi¬ tion of the tree when it is stressed (Wargo 1978a,b). Starch content has been used as an indicator of physi¬ ological performance and the effects of stress (Wargo 1978c). The susceptibility of stressed trees that are low or depleted in starch content probably relates, in part, to the reduced energy available for defense reactions (McLaughlin and Shriner 1980). For example, many oak trees are colonized by Armillaria after defoliation by the gypsy moth, but not all trees are infected, and not all infected trees are colonized to the same extent (Wargo 1977). Mortality of oak and sugar maple after defoliation was related to carbohydrate production and storage (Wargo 1981b,c,e; Wargo and Houston 1974). Trees with low or depleted starch when defoliation occurred were more likely to be colonized by Armillaria and to die after stress from defoliation (fig. 7.1). Starch content at the time of stress was related to how long a tree survived and how many defoliations it could tolerate. Barriers and Energy Reserves Preformed physical barriers such as outer bark play an important role in protecting roots from invasion by pathogens (see chapters 4 and 5). Outer bark may offer less protection from Armillaria than from those root- rotting fungi that cannot penetrate without wounds. Existing evidence does not suggest that predisposing stresses enhance susceptibility by allowing penetration through intact outer bark. However, some stress agents may cause bark injury and provide infection courts for Armillaria. Wind-induced root movements and break¬ age (Harrington 1986, Hintikka 1972, Rizzo and Harrington 1988b), rock abrasions (Stone 1977), and insect feeding provide infection courts for Armillaria and other root pathogens (Redmond 1957, Whitney 90 Host Stress ami Susceptibility B FIGURE 7.1 —Armillaria and energy reserves in roots of sugar maple. A: Sections of roots from defoliated (left) and non- defoliated (right) trees inoculated with A. gallica and incubated for four weeks; B: Starch reserves in roots from defoliated (left) and nondefoliated (right) trees in the fall after defoliation. Starch grains have been stained with l 2 KI and appear purple- black in the tissue. (P. Wargo) 1961). Wounding and root breakage also stress trees since the tree must expend energy to close the wound, prevent infection, and replace damaged roots. Wounds may not be so important in removing the bar¬ rier of the dead outer bark as they are in removing the living, responsive, inner bark. Once the outer bark is penetrated, the pathogen encounters living tissues where physiological factors, such as lytic enzymes or toxic secondary metabolites, may limit hyphal penetra¬ tion of the inner bark. The limitation of Armillaria hyphae developing within healthy host plant tissues has been described for the mycotrophic association between the fungus and achlorophyllous orchids (Hamada 1940, Kusano 1911, see chapter 8). In this relationship, lysis of the hyphae and reinfection by the fungus occur seasonally. The mechanism of hyphal lysis is unknown, but it could result from digestion by host enzymes. Chitinase and B- 1,3-glucanase, enzymes that can dissolve the hyphal wall of Armillaria, are present in the inner bark and sap of forest tree species (Wargo 1975), and they constitute a potential mechanism to limit the growth of Armillaria calvescens hyphae in resistant bark tissue (Wargo 1975, 1976, and unpubl.). The activities of these enzymes are reduced by stress from defoliation (Wargo 1976). An important component of the resistant reaction of the inner bark is the formation of wound periderms (Biggs and others 1984, Rykowski 1975, Thomas 1934). Some general observations indicate that stressed trees cannot produce periderms rapidly or fail to form wound peri¬ derms in response to Armillaria (Rykowski 1975). Even if they are formed, under some circumstances Armillaria has the ability to penetrate such suberized periderms (Rykowski 1975), probably by enzymatic degradation (Swift 1965, Zimmermann and Seenuiller 1984). Conversion of extant energy reserves into secondary compounds in response to wounding or invasion of inner bark or sapwood may benefit the host by forming compounds that are directly toxic to the pathogen, that are unavailable for pathogen metabolism, or that protect more complex carbohydrates from fungal extracellular enzymes (Worrall anti Harrington 1988b). Gums, resins, phenolic compounds, and other metabolites may be produced in higher concentrations in response to wounding or invasion by pathogens than in unaltered sapwood (Hepting and Blaisdell 1936, Shain 1967). Oleoresins in the inner bark and sapwood of conifers are potentially inhibitory to the fungus and are secreted in response to infection and colonization by Armillaria. Volatile components of oleoresin from Scots pine re¬ duced the growth of Armillaria on agar by half (Rishbeth 1972a), and fewer rhizomorphs of A. ostoyae developed from resinous rootwood of Corsican pine than from non-resinous rootwood (Rishbeth 1985b). Roots of stressed conifers do not produce as much resin as healthy trees, and root tissues are colonized by fungi more rapidly than are roots of unstressed trees (Gibbs 1967,1968; James and others 1980a,b; Rykowski 1975). In spite of the emphasis on the role of the fungus as a phloem colonizer, Armillaria is capable of colonizing the inner wood of roots and stems without killing phloem tissues. This typical root- and butt-rot colonization may occur in relatively vigorous trees capable of resisting phloem colonization, and may proceed for decades without host mortality (Shigo and Tippett 1981, Tippett and Shigo 1981). Two general sapwood responses are known (see chapter 5). First, sapwood tissues may be converted to non¬ living, reaction zone tissues that resist pathogen coloni- Host Stress and Susceptibility 91 zation (Shain 1967). Inhibitory, secondary compounds similar to those in the inner bark are also found in the reaction-zone tissues of the sapwood. As discussed in connection with the inner bark, these compounds re¬ quire substantial energy reserves, and in stressed trees may not be produced in sufficient quantity or soon enough to stop Armillaria colonization. Second, whether or not the pathogen becomes estab¬ lished in the reaction zone, another impediment to pathogen development, the barrier zone, may be formed. The cambium may respond by producing a unique layer of xylem that resists penetration by the pathogen and tends to restrict it to those growth rings of xylem formed prior to injury (Hepting and Blaisdell 1936, Shigo and Larson 1969). Barrier zones of this sort, formed in response to infection and colonization by Armillaria, have been observed in roots of both conifers and hardwoods (Shigo and Tippett 1981, Tippett and Shigo 1981). Although evidence is limited, sapwood and cambium of less vigorous trees may form less inhibitory reaction zones and weaker barrier zones than the sapwood and cambium of healthy trees (Armstrong and others 1981, Shearer and Tippett 1988, Shigo and Hillis 1973). In such cases, Armillaria may be slowed but not stopped from developing in the sapwood, and continued devel¬ opment reduces the amount of sapwood available for water transport, increases the energy expended in re¬ sistance responses, and may allow penetration from the sapwood into the cambium and inner bark. Pathogen Nutrition Stress also affects resistance indirectly by nutritionally enhancing Armillaria growth. Predisposition of defoli¬ ated sugar maple to Armillaria occurs in part through changes in the carbohydrate and amino nitrogen com¬ pounds induced by defoliation (Wargo 1972). Severe defoliation triggers hydrolysis of starch and results in large increases in reducing sugars in the cam- bial zone and neighboring tissues (Parker 1970, Parker and Houston 1971, Wargo 1972, Wargo and others 1972). Qualitative and quantitative changes in amino nitrogen also occur (Parker and Patton 1975, Wargo 1972) and, combined with increases in glucose, signifi¬ cantly stimulate the growth of Armillaria calvescens in vitro (Wargo 1972,1981a, and unpubl.) (fig. 7.2). Hy¬ drolysis of starch to glucose would certainly be more beneficial (nutritionally) to Armillaria than would con¬ version of starch to secondary metabolites, as would occur in the production of reaction zone tissues in healthy trees. Stresses, such as excess soil moisture and defoliation, may also increase the ethanol in root tissues (Wargo unpubl.). Ethanol is a potent growth stimulant for Armillaria (Weinhold 1963) and its presence in root tissue could affect susceptibility to the fungus. The host may directly produce ethanol in response to stress (Coutts and Armstrong 1976, Crawford and Baines 1977); ethanol may be produced by associated microor¬ ganisms and promote the growth of Armillaria (Pentland 1967); or under anaerobic conditions, Armillaria may produce its own ethanol (Tarry 1969). Chemical changes in roots of stressed trees apparently allow the fungus to metabolize phenols and probably other compounds that would normally inhibit it (Wargo 1980a, 1981d, 1983b, 1984a,b). Glucose, ethanol, and nitrogen levels and nitrogen source affect the abil¬ ity of the fungus to oxidize phenols in vitro. Oxidation and polymerization of phenols by Armillaria can re¬ move those that are inhibitory or that precipitate extra¬ cellular fungal enzymes. Also, phenol metabolism affects melanin formation by Armillaria (Bell and Wheeler 1986, Malama and others 1975, Worrall and others 1986) and could provide rhizomorphs and pen¬ etrating hyphae greater protection against enzymatic lysis from host-produced enzymes (Bloomfield and Alexander 1967). All of these host-pathogen biochemi¬ cal interactions are discussed more fully in chapter 3. DATE OF DEFOLIATION FIGURE 7.2 — Reducing sugar concentrations (% dry wt) in ex¬ tracts from roots of defoliated and nondefoliated sugar maple, and fungal dry weight of A calvescens after 3 weeks' growth on the extracts. Unlike letters above the bars indicate a signifi¬ cant difference at P=0.01. (Modified from Wargo 1972) 92 Host Stress ami Susceptibility Stress Agents and Armillaria Root Disease General Trees are exposed to stress throughout their lives. Stresses such as drought, waterlogging, frost damage, some pollution events, insect defoliation, other tree diseases (especially foliar diseases), and short-term coppice cutting may be considered acute (short dura¬ tion, high intensity). Other stresses may be considered chronic in that the tree may be exposed over its life time to low but relatively constant levels of the stress. Air pollutants, soil nutrient deficiencies, and long-term moisture deficiencies are examples of chronic stress. Shade-intolerant trees in forest understory can also be considered chronically stressed from reduced light. Acute stresses may affect the metabolism of the entire tree, and Armillaria may rapidly colonize the entire root system or the root collar region of such trees and kill them quickly. Colonization of the roots of defoliated oak and sugar maple exemplifies this relationship (Wargo 1977, Wargo and Houston 1974). When acute stresses affect only a portion of the tree, Armillaria inva¬ sion may be partial and sometimes progressive, caus¬ ing the tree to die slowly over several years. The relationship of Armillaria and beech bark disease dem¬ onstrates this interaction. Armillaria usually colonizes only those roots of American beech that are associated with the portion of the stem killed by Nectria coccinea var. faginata Lohman, Watson and Ayers, a canker- causing fungus (Wargo 1983a). The timing of the stress event is also very important (Wargo 1978b). Stresses that occur early in the growing season and then abate have less of an effect than mid¬ season stresses because the trees have more of the growing season in which to recover. Likewise, stresses occurring late in the growing season may cause less harm because most of the growth and energy produc¬ tion by the tree has already occurred. The effects of any stress, no matter when it occurs, ultimately depend on its duration within and across growing seasons. Stresses may also interact. Defoliation by phytopha¬ gous insects, especially those associated with oaks, have historically been linked to drought (Falck 1918, 1923; Houston 1981a,b, 1984; Nechleba 1915). These two stress factors working in concert affect tree health, resulting in widespread mortality, much of it associ¬ ated with Armillaria. Defoliation can also exacerbate Armillaria root disease on beech affected by beech bark disease. On defoliated trees, Armillaria spreads from existing lesions on roots associated with the stem can¬ ker into adjacent roots and root collar tissues, resulting occasionally in rapid mortality (Houston 1974a, Wargo 1983a). Abiotic Stress Factors Light Predisposition to Armillaria root disease from inad¬ equate light has been observed in natural forests and plantations, and it has been demonstrated experimen¬ tally. Armillaria commonly attacks suppressed under¬ story trees, upon which it acts as an ecosystem scavenger (Davidson and Rishbeth 1988, Pearce and others 1986, Rishbeth 1983). While these trees may be more susceptible to Armillaria attack because of genetic makeup, they are also affected by the reduced sunlight, which reduces the amount of energy available for de¬ fense against pathogens. Susceptibility, therefore, would be influenced by the shade tolerance of the tree species. Redfern (1978) demonstrated the effects of insufficient light on susceptibility of trees to Armillaria in both plantation and greenhouse studies. Dominant and sup¬ pressed Scots pine in a 19-year-old plantation were inoculated and examined after 9 months. Similar num¬ bers of dominant (12/15) and suppressed (13/15) trees were infected; however, the severity of infection, as measured by length of root invaded, was greater in the suppressed trees than in the dominant trees. Two sup¬ pressed trees were killed. Inoculation studies with known Armillaria species on subdominant trees and suppressed trees growing in reduced light showed that suppressed English oak and Scots pine were infected by A. mellea, A. ostoyae, and A. gallica but not by A. tabescens (Davidson and Rishbeth 1988). Only A. mellea colonized the healthier subdomi¬ nant oaks, and only A. ostoyae colonized the subdomi¬ nant pine. In one greenhouse study, Armillaria killed significantly more seedlings of Japanese larch growing under an 8- hr daylength than those growing under a 16-hr daylength for 20 weeks (Redfern 1978). In a second greenhouse study, seedlings of grand fir, western hem¬ lock, and English oak were inoculated and grown un¬ der shade (70% light reduction) and compared with seedlings grown in full sunlight (Redfern 1978). Light did not affect susceptibility of western hemlock, which is a shade-tolerant species; 60% of seedlings were killed in each treatment. Shade, however, increased the sus¬ ceptibility of the less-tolerant species, with 44% and 76% of the fir and 2% and 22% of the oak seedlings killed in full sunlight and shade treatments, respec¬ tively. Greenhouse studies with A. ostoyae on western Host Stress and Susceptibility 93 white pine also showed that very young seedlings (3- week-old) were more susceptible to infection if grown under reduced-light conditions (Entry and others 1986). Temperature Both high and low temperature extremes can stress trees and render them susceptible to opportunistic organisms. The effects of high temperatures, however, are com¬ monly associated with drought, and distinguishing their individual effects can be difficult. In his report on envi¬ ronment and Armillaria root disease. Day (1929) indi¬ cated that the fungus attacked trees affected by sun scorch, drought, and defoliation. Sun scorch on leaves is caused by high temperatures associated with dry condi¬ tions and can significantly damage trees. Hole (1927a,b) found that drought and sun scorch on the foliage and sunscald on the smooth bark of morinda spruce in India significantly injured the crowns and predisposed the root systems to Armillaria colonization. Mortality was greatest on the hot, western and southern slopes and least on the cool, northern sites. Elevated soil temperature, attributed to a slightly warmer summer climate and opening of the canopy by extensive logging, was proposed as a major factor in birch dieback in eastern Canada and Maine (Redmond 1955). Experimentally elevating the soil temperature by 2°C increased rootlet mortality from 6% to 60%. Trees in stands suffering "birch dieback" were characterized as having progressively greater rootlet mortality as crown vigor decreased. These trees were frequently colonized by Armillaria, but the fungus was not considered the primary cause of this decline (Hansbrough and others 1950, Spaulding and MacAloney 1931). Bliss (1946) found that the greatest resistance to infection and colonization by Armillaria occurred at soil tempera¬ tures that were most favorable for root growth. Viru¬ lence was greatest at lower soil temperatures (10-18°C) on host species with a high soil temperature range for optimum root growth (17-31°C), such as peach, apricot, and geranium. On host species with a low soil tempera¬ ture range for optimum growth (10-17°C), such as sweet orange, sour orange, orange and rose, virulence was greatest at higher soil temperatures (15-25°C). Stress from freezing damage and subsequent coloniza¬ tion by Armillaria is documented for snowbrush. Severe crown kill of this evergreen shrub occurred in 1963 in Montana during a winter of light snow and after a sud¬ den temperature drop from above freezing to -14°C to -20°C (Stickney 1965). A subsequent survey of snow¬ brush dieback in the Northwestern United States showed that Armillaria was associated with dead and dying clumps of this shrub (Tarry and Shaw 1966). Per- . ps iiie freeze-damage predisposed the shrub to Armillaria. Subsequent work on this dieback (Tarry 1969) showed that 77% of the declining snowbrush stumps were infected by Armillaria. Results of inocula¬ tions in healthy plants were poor; less than 5% of 108 inoculation attempts resulted in infections, suggesting that colonization depended primarily on predisposing stress. Infection and colonization of peach trees by Armillaria and other secondary organisms also were attributed (Poole 1933) to sudden exposures to low temperatures (-12°C to -9°C). These were extremes for peach or¬ chards in the Carolinas (United States), and tree mor¬ tality ranged from 10% to 100%. Damage from late spring frosts also predisposes trees to Armillaria. In North Carolina, late spring frosts were associated (Beal 1926) with the death of numerous white oaks. Later work indicated that much of this mortality was associated with Armillaria and bark in¬ sects (Hursh and Haasis 1931). Armillaria also infected chestnut trees (probably American chestnut) twice defoliated by late spring frosts (Long 1914). Trees can also be stressed from events associated with but not directly related to low temperatures. Severe deterioration of an 80-year-old stand of red oak after a severe ice storm was attributed to Armillaria which colonized trees weakened by ice damage to their crowns (Dance and Lynn 1963). Hintikka (1974) sug¬ gested that Scots pine in plantations were predisposed to Armillaria infection by heavy snows that severely bent the saplings. However, Armillaria damage was severe in these snow-damaged plantations, perhaps due to increased wounding of the roots that lifted when the trees were bent rather than from direct reduc¬ tion in tree vigor. Moisture Drought is probably the most common stress affecting trees, and at some time during most years trees experi¬ ence either short- or long-term reductions in soil mois¬ ture. In their reviews on the relationship of Armillaria with widespread dying-off of forest stands in Europe, Twarowski and Twarowska (1959) and Nechleba (191a) indicate that attack of both conifer and hardwoods by Armillaria has been associated with drought since the late 1800's. Parasitism by Armillaria on true fir species was reported to increase during dry seasons, while wet seasons favored its saprophytic role (Nechleba 1927). Muller (1921) observed that droughts in the 1890's and early 1900's preceded Armillaria- caused deaths of many firs in Germany. Nechleba (1915) suggested that drought was the major factor in predisposing conifers to Armillaria and that the fungus "... under normal con¬ ditions of moisture and temperature, is a pronounced 94 Host Stress and Susceptibility and blameless saprophyte." Falck (1918,1923) and Hen (1914) observed that drought was also involved in predisposing oaks to Armillaria. These early observa¬ tions of stress-induced susceptibility to Armillaria led to the widely held view of the fungus as a secondary pathogen on forest trees. Later reports also confirm the relationship of drought and Armillaria. Biraghi (1949) observed that infection of fir was enhanced during prolonged drought; however, mechanical injury also played a role. In East Africa, radiata pine were killed by Armillaria after an extended drought (Anon. 1952). In the United States, drought and subsequent Armillaria infection have been reported for western white pine (Ehrlich 1939), eastern hemlock (Secrest and others 1941), and balsam fir (Livingston and others 1982). Oak decline and mortality in the United States have been frequently associated with drought conditions. Drought, in combination with defoliation from late spring frosts, followed by attack of the stressed trees by Armillaria, resulted in large-scale mortality in white, black, red, and scarlet oaks (Hursh and Haasis 1931). Staley (1965) also concluded that drought and defolia¬ tion from insects and frost damage predisposed scarlet oak to Armillaria. Similar relationships of drought, de¬ foliation, and mortality of oak associated with Armillaria root disease were observed in Europe (Falck 1918, Hen 1914, Georgevitch 1926b). The European situation was further complicated by powdery mildew fungi that caused additional defoliation. Drought also predisposes other hardwoods to Armillaria. The severe drought in the late 1950's through the mid 1960's in the Eastern United States was considered a predisposing factor in sugar maple decline. Armillaria afflicted 46% of symptomatic sugar maple trees in New York State in the early 1960's (Hibben 1964). Drought is also the most likely initiator of regrowth dieback of eucalypts in Tasmania where A. hinnulea and A. novae-zelandiae are important secondary pathogens (Kile 1980b, Kile and Watling 1983). In a review paper on forest declines, Houston (1987) listed seven dieback and decline diseases, their epi¬ sodic occurrence in North America since the early 1900's, and their associated stress factors and second¬ ary organisms. Drought was listed as a stress factor in five of the seven diseases discussed; root-rot fungi, predominantly Armillaria, were involved in most of the declines. Other associations of drought, forest decline diseases, and Armillaria appear in table 8.3. Root-system development may play some role in the predisposition effects of drought. Observations of Armillaria root disease on Scots pine indicated that root systems of healthy trees were deeper and better developed than those of infected trees. Susceptibility to drought and subsequent infection by Armillaria were favored in trees with a shallow, poorly developed root system (Ritter and Pontor 1969). Shallow roots and prolonged drought stress (7 years) were also associated with the decadence of eastern hemlock in Wisconsin (Secrest and others 1941). Declining trees were colo¬ nized by Armillaria, and root systems of some living trees with "normal" green crowns were also com¬ pletely colonized by the fungus. Excess moisture may be as stressful to trees as drought in regards to Armillaria root disease. However, the majority of such reports concern hardwood species. Excess soil moisture can cause physiological drought by interfering with water uptake in oxygen-deprived roots. Also, anaerobic conditions in the roots promote the production of ethanol, which can stimulate aggres¬ sive Armillaria growth (see chapter 3). An early report on Armillaria root disease in the United States (Long 1914) indicated that Armillaria attack on various oak species and chestnut was greater and more severe on sites where the soil was wet sea¬ sonally- Wet summers also were observed to predis¬ pose chestnut species to Armillaria infection in Germany and Austria (Bazzigher 1956). Native oaks in California were apparently infected with but not usually killed by Armillaria unless they were irrigated during the summer (Raabe 1966a). Whether irrigation resulted in excess soil moisture that stressed the trees or provided a better environment for more aggressive growth of Armillaria was not deter¬ mined. Dade (1927) observed that high humidity pro¬ moted infection in cocoa. High rainfall years and poor soil drainage were also linked to infection of rubber trees in Nigeria (Fox 1964). Decline of ohia has occurred periodically in Hawaii since 1875 and has been associated with poor soil drainage which, as the trees age, eventually predis¬ poses them to Armillaria and other agents (Hodges and others 1986). In Japan, Armillaria on larch was related to low host vigor as indicated by annual growth incre¬ ments, but incidence of infection depended mainly on the amount and duration of excess soil moisture (Kawada and others 1962). Disease was especially se¬ vere where larch were growing on soils with a high or perched water table. Nutrients and Other Soil Factors Armillaria root disease generally occurs more fre¬ quently and severely on nutrient-deficient soils or on Host Stress and Susceptibility 95 soils with poor physical and chemical characteristics for host growth. Armillaria- caused mortality in tea plantations growing in nutrient-deficient soils was considerably greater than in areas where soil fertility was adequate for growth (Butler 1928). In a 32-year-old plantation of eastern white pine in New York, damage by Armillaria was associated with low soil nutrients (Silverborg and Gilbertson 1961). Ono (1965,1970) re¬ ported that Armillaria caused severe losses in Japanese larch plantations on both upper slopes and lowlands. In both areas, he attributed disease severity to physical and chemical soil characteristics unfavorable for larch. Some evidence suggests that predisposition by nutrient deficiency depends on which tree species grows where a particular nutrient is low. Reduced nitrogen and phosphorus levels were linked to rapid development of Armillaria root disease in conifer plantations in New¬ foundland (Singh 1970). Calcium deficiency was re¬ lated to increased Armillaria damage in walnut plantations (Marchal and Foex 1931). Low soil nitrogen and low soil pH were associated with Armillaria- caused decay in Douglas-fir, while low soil calcium and phos¬ phorus and high soil potassium were associated with Armillaria- caused decay in grand fir (Shields and Hobbs 1979). Armillaria root disease has been related to extractable aluminum concentrations in soils from sites surveyed for root disease. Browning and Edmunds (1985) found that incidence of A. ostoyae on coastal Douglas-fir in the Northwestern United States was generally higher on sites where aluminum levels in the soil were low. Labo¬ ratory studies did not conclusively confirm this rela¬ tionship (Browning 1987). Aluminum inhibited fungus growth but only at high concentrations in buffered media (200 ug/g and above). Fungal growth in coastal soil extracts decreased as extractable aluminum mea¬ sured in these soils increased, but the correlation was not significant. Inoculated seedlings growing in soils from sites with high and low disease incidence also failed to associate disease incidence with extractable aluminum (Browning 1987). Relationships between nutrients and susceptibility to Armillaria have been demonstrated experimentally. Rate, incidence, and severity of infection of seedlings of Norway spruce, black spruce, Sitka spruce, and Scots pine were greater when they were grown in forest soil with low nutrient levels and low pH (Singh 1983). Three-week-old seedlings of western white pine grown under reduced light and nutrient deficiencies were also infected more frequently and more severely than seed¬ lings grown under adequate light and nutrient supply t itry and others 1986). With adequate light, more 10% F Eklund & Wennmark (1925) Sweden 5.5% F Bjorkman and others (1964) P. tremuloides Michx. Minnesota M Schmitz & Jackson (1927) Ontario 27% F Basham (1958) see also Black (1951) Alberta 0.9% DV Thomas and others (1960) Ontario 9.2% F Basham & Morawski (1964) Colorado 0.5% DV Hinds and Wengert (1977) Quebec M Laflamme & Lortie (1973) P. trichocarpa Torr. & Gray British Columbia 3.1% DV Thomas & Podmore (1953) Quercus spp. Eastern USA M Hepting & Hedgcock (1937) Eastern USA 10% F Roth & Sleeth (1939) Tilia americana L. Ontario 54.5% F (small sample) Basham & Morawski (1964) * Covers records from both natural forests and plantations (Chapter 9), although the latter are restricted to European records for Ptcea abies, P- sitchensis, Thuja plicata, Tsuga heterophylla, Larix decidua M = minor if recorded as such or effects not quantified but appear to be so on the evidence presented and in the sense of causing little loss of merchantable timber volume. % Frequency = incidence of infection in trees assessed or percentage of identified infections. % DV = percentage of the total decay volume recorded attributed to Armillaria species. Natural Forests 109 Few observations in natural forests document the spe¬ cific site and stand factors which might affect the inci¬ dence and severity of Armillaria butt rot. This reflects the relatively minor contribution of Armillaria to butt rot losses, being treated usually as incidental to those of more destructive stem-rot organisms. Birch (1937) considered overstocking may contribute to the high incidence of butt rot in silver beech pole stands in New Zealand. Basham and others (1953) noted that decay caused by Armillaria was more frequent in stands on poorly drained sites than in stands on relatively well drained slopes with more hardwoods in the stands. Most butt rots are believed to develop via root infec¬ tions. Basham (1958) suggested that in quaking aspen wind stress and frost heave could facilitate the entry of butt rot organisms, with occasional entry through basal wounds. Nordin (1954) found that frost cracks could provide entry points in sugar maple. Armillaria butt rot may also occur in the same tree with other decay organisms such as Heterobasidion annosum (Fr.) Bref. (Kallio and Norokorpi 1972, Molin and Rennerfelt 1959), or Armillaria infection may allow host entry for other decay organisms such as Phaeolus schiveinitzii (Barrett 1970, Barrett and Greig 1984, see chapter 5). The general effects of butt rot on host growth rates and longevity are poorly understood (Wagener and Davidson 1954) though information on the relative susceptibility of some species to Armillaria butt rot has been obtained under plantation conditions (Gladman and Low 1963, Greig 1962). However, these findings may need to be interpreted in relation to the Armillaria species involved. Lethal Primary Disease Armillaria kills trees in natural coniferous and hardwood forests in different spatial and temporal patterns and with ecological and economic effects of varying signifi¬ cance. As noted earlier, it is part of the continuum of primary disease effects (fig. 8.3). Four such primary dis¬ ease syndromes may be recognized around the world: (1) Armillaria root disease in boreal forests and western North American coastal conifer forests; (2) ring disease of mountain pine in France; (3) root rot of mixed-species conifer forests in western North America; and (4) root rot of dry sclerophyll eucalypt forests in Australia. An addi¬ tional and historically interesting report of possible pri¬ mary parasitism, but for which no further information is available, is that of Geschwind (1920),who observed mor¬ tality of conifers when common beech was selectively logged from mixed forest in Bosnia and Herzegovina (Yugoslavia). ;rs ; three diseases involve mainly coniferous spe- md a common Armillaria species, A. ostoyae, al¬ though other Armillaria species may be pathogenic in boreal and mixed-species coniferous forests. On a qualitative basis the last three appear to be the most significant diseases for their impact on stand structure and progressive disease development. Other common features of these latter three diseases include their oc¬ currence in relatively drier environments, the discon¬ tinuous distribution of the pathogens within the affected forests, the apparent greater pathogenicity of the causal Armillaria species compared with the same species or different species in wetter forest environ¬ ments, and the apparent intensification of disease de¬ velopment following harvesting operations. Root Rot in Boreal Forests and Western North American Coastal Coniferous Forests Mortality of single or small groups of seedlings or sap¬ lings occurs early in stand development of naturally regenerated, moist coniferous forests in North America and northern Europe-Scandinavia (Baranyay and Stevenson 1964; Bourchier 1954; Buckland 1953; Hintikka 1974; Mallet and Hiratsuka 1985; Morrison 1981; Whitney 1978b, 1988a; Whitney and Myren 1978; Whitney and others 1974). Mortality typically com¬ mences soon after stand establishment, reaches a maxi¬ mum at age 10-20 years, and then decreases in frequency, possibly as the food base declines and host tolerance increases. Effects on overall stocking are usu¬ ally minimal, although the disease pattern may vary in some regions with limited mortality occurring through the rotation. Trees may survive with root and butt in¬ fections (Morrison and others 1985a, Whitney 1988a, Whitney and Myren 1978, Whitney and others 1974). Disease expression may sometimes be associated with stress (Buckland 1953, Baranyay and Stevenson 1964). A number of Armillaria species occur in affected forests, although most disease appears coincident with the presence of A. ostoyae (Dumas 1988, Mallet 1989, Morrison and others 1985a, Whitney 1988a). Beyond the natural range of this species or where its distribu¬ tion is limited, disease is less prominent. Thus in south¬ eastern Alaska forests, where less pathogenic species such as A. sinapina may be widely distributed, little killing is evident in regeneration stands (Shaw 1981b, Shaw and Loopstra 1988). Armillaria borealis and A. cepistipes cause minor mortality and butt rot in Finnish forests (Korhonen 1978, Piri and others 1990). Ring Disease of Mountain Pine In relatively undisturbed 120- to 150-year-old moun¬ tain pine forests at 1,600-2,200 m in the eastern Pyrenees, mortality from A. ostoyae is extensive and chronic (Durrieu and others 1981,1985; Durrieu and Chaumeton 1988). Killing may be diffuse but most 110 Natural Forests characteristically occurs in scattered but clearly delim¬ ited rings with a marginal zone of dying and dead trees (fig. 8.4). Ring diameter may reach more than 120 m and may expand 1 m per year. Historical ring develop¬ ment, followed on aerial photographs taken over a 36- year period (Durrieu and others 1981), indicates some rings show intermittent development while others cease expanding and gradually disappear. Following stand opening, mountain pine begins regenerating and is only moderately susceptible to A. ostoyae. A succes- sional sequence occurs in the understory/ground flora until the forest returns to a pre-disease form. The origin of the rings, the means of pathogen spread within them, and the factors controlling their initiation are poorly understood. The affected forests occur on light-textured, shallow soils, often on steep slopes; rainfall is relatively low (600-750 mm per annum), and bark beetles may act as a stress agent (Torossian 1984). However, the long-standing and strongly patterned nature of the disease and the infection and killing of provence broom, an understory species, supports A. ostoyae as the primary disease cause. Durrieu and oth¬ ers (1985) suggested the fungus is part of the forest's natural ecology, leading to the regeneration of the dominant tree species. While the disease is most severe in the Cerdagne region of France, it also extends west¬ wards into drier transitional forests and may also occur in other parts of the range of mountain pine (Brang 1988). Armillaria Root Disease in Mixed Coniferous Forests of Western North America Lethal primary disease affects hundreds of thousands of hectares of natural coniferous forests in western North America. The primary documented areas of forest management concern, where Armillaria root disease occurs most extensively and severely, are east¬ ern Oregon and Washington, northern Idaho, western Montana, and the southern interior of British Colum¬ bia. The disease is also recognized in the central and southern Rocky Mountains (Wood 1983). In these drier, interior forests, continuing mortality in all age classes is common in many stands; understocked stands may result from multiple disease centers. Armillaria root disease has been known for many decades in these forests (Ehrlich 1939; Hubert 1918,1931, 1950,1953) but has received minimal attention until the mid 1970's largely because the overall impact was not appreciated. Smith (1984) estimated the average annual volume losses to five major root diseases [Phellinus zoeirii (Murr.) Gilb., H. annosum, Armillaria spp., Phaeolus schiveinitzii, and Ophiostoma wageneri (Goheen: Cobb) Harrington] throughout the western United States to be 6.7 million m 3 , or 18% of the total annual mortality. While the proportion of this loss due to Armillaria root disease cannot be determined, local severity has been evaluated. Shaw and others (1976a) found volume loss to Armillaria in a ponderosa pine stand in south-central Washington to have increased from 9 m 3 per ha in 1957 to 24 m 3 per ha in 1971. In a mixed-conifer stand in southern Oregon, Filip (1977) found 7% of trees com¬ prising 32% of the standing volume were infected with or killed by Armillaria. Filip and Goheen (1982,1984) found annual mortality of more than 3 m 3 per ha in other situations. In Montana, a root disease patch in a Douglas-fir stand contained 82% less timber volume per 0.4 ha than the adjacent healthy stand (Byler unpubl.). In British Columbia's interior cedar-hemlock zone, annual timber losses caused by Armillaria root B FIGURE 8.4—Development of ring disease in mountain pine for¬ est, Pyrenees, France. A. Photographed in 1942. B. Same area photographed in 1962. (G. Durrieu) Natural Forests lU disease were estimated to be 105,000 rrP (Taylor 1986). Volume growth of Douglas-fir infected by A. ostoyae in four stands in southeastern British Columbia decreased significantly as disease severity, measured by basal resinosus, increased from infected stem bases (Bloomberg and Morrison 1989). Armillaria was also recognized as a major cause of stand damage in other ground and aerial surveys which have recorded incidence and area of root disease centers (Williams and Leaphart 1978). James and others (1984) estimated active root disease centers, mainly attributed to Armillaria and Phellinus weirii, occupied almost 32,000 ha (about 1% of the total commercial forest land) of seven national forests in the northern Rocky Mountains. A detailed study of one of those forests, the Lolo, found 123,255 ha (18.8% of the total forest) were diseased, of which 8,011 ha (1.2%) were unstocked patches (Byler and others 1990). Besides timber loss and the creation of unproductive areas through chronic infection, particularly where susceptible hosts are climax (McDonald and others 1987a), Armillaria root disease may change species composition, create hazardous trees in recreation for¬ ests, and affect the choice of silvicultural system. Armillaria ostoyae (NABS I 1 ) (Morrison and others 1985a, Wargo and Shaw 1985) and possibly NABS X (McDonald unpubl.) are pathogenic on conifers in these interior west¬ ern forests, although A. ostoyae is considered the most widespread and aggressive. Additional taxonomic or biological species known to be present in western North America are A. sinapina (NABS V), A. gallica (NABS VII), NABS XI (A. cepistipes?), and NABS IX (Anderson and Ullrich 1979, Morrison and others 1985a, Shaw and Loopstra 1988, Wargo and Shaw 1985). The latter two species have been collected infrequently. Some of these species may act as secondary pathogens. Where they occur, Armillaria species have a complex interaction with about two dozen conifer species. Data on mortality rates resulting from root disease caused by Armillaria in different community types and geo¬ graphic areas are lacking, although observations indi¬ cate Douglas-fir and true fir are the most susceptible (Hagle and Goheen 1988, Morrison 1981). Exceptions to this occur in south-central Washington where pon- derosa pine is most susceptible and Douglas-fir ap¬ pears tolerant, and possibly in some other areas where Engelmann spruce (McDonald and others 1987b) and western hemlock appear very susceptible (Morrison 1981). Root disease may also afflict hardwood shrubs (Adams 1974; McDonald and others 1987a,b; Morrison N \BS (N. t merican Biological Species) as described fully in chapters 1 and 2. 1981; Shaw 1975; Tarry and Shaw 1966; Williams and Marsden 1982). In individual stands, mortality often begins within a few years of regeneration and may continue through¬ out the rotation, particularly in Douglas-fir/true fir forests. For other species, such as western redcedar, mountain hemlock, western larch, western white pine, ponderosa pine, and lodgepole pine, damage tends to diminish with stand age beyond 20-30 years. Disease occurrence varies from individual infected trees (fig. 8.5) to patches (fig 8.6) of tens of hectares (Byler unpubl., Filip 1977, James and others 1984, Smith 1984, Wargo and Shaw 1985). Patches typically contain coni¬ fer regeneration, brush, or grass and have a perimeter of dead and dying trees. Rate of disease spread in a ponderosa pine stand was 1-2 m per annum (Shaw and Roth 1976), but in a Douglas-fir stand less than 0.25 m per annum (Byler unpubl.). Typical infection foci are usually occupied by 1-3 Armillaria genotypes (Adams 1974, McDonald and Martin 1988, Shaw and Roth 1976). The dynamics of disease within infection centers and across rotations in these mixed-conifer forests is discussed further in chapter 10. Armillaria frequently causes damage concomitant with other root-rot pathogens of mixed-conifer forests, par¬ ticularly with Phellinus weirii (Filip and Goheen 1984, Goheen and Filip 1980, James and others 1984, Miller and Partridge 1973. Williams and Leaphart 1978), but also with O. wageneri (Cobb and others 1974) and H. annosum (F. Cobb pers. comm.). Armillaria may be active on the same site or in the same root system as other pathogens. Hansen and Goheen (1989) attributed these associations to chance and to primary-secondary relationships, but the roles have not been adequately defined. Armillaria infection and other root diseases predispose some conifers to bark beetle infestation (Hertert and oth¬ ers 1975, Hinds and others 1984, James and Goheen 1981, Kulhavy and others 1984, Lane and Goheen 1979, Lessard and others 1985, Partridge and Miller 1972, Tkacz and Schmitz 1986). Armillaria root disease may be an impor¬ tant factor in the survival of endemic populations of some bark beetle species. Hinds and others (1984), Lessard and others (1985), and Tkacz and Schmitz (1986) associated such populations of the mountain pine beetle (Dendroctonus ponderosae Hopkins) with Armillaria infec¬ tion in ponderosa and lodgepole pine stands in South Dakota and Utah, respectively. Interaction between bark beetles and Armillaria root disease is considered further in chapter 10. Western North America is marked by complex land- forms and specific associations of plant communities. Large variation in elevation, aspect, slope, altitude, and 112 Natural Forests A B FIGURE 8. 5— Armillaria root disease killing individual trees near infected stumps in a mixed-species conifer forest in western North American. A: Ponderosa pines; B. Grand fir. FIGURE 8.6—Armillaria root disease center in virgin coniferous forest in western North America. The lowermost center covers nearly 8 ha (20 acres). soil type has produced an elaborate mixture of forest ecosystems with widely differing levels of vulnerability to Armillaria root disease. Root disease centers have been associated with particular forest habitat types (Byler and others 1986,1990; McDonald 1990; McDonald and others 1987a,b; Williams and Marsden 1982). Armillaria root disease probably played an important role in forest succession and the determination of stand composition and structure on many mixed-conifer forest sites prior to European settlement (Byler 1984, Byler and others 1990, Hagle and Goheen 1988, Haig and others 1941, Shaw and Roth 1976, Wargo and Shaw 1985). Armillaria ostoyae, for example, accelerates suc¬ cession in interior British Columbian forests especially on wetter sites. There, the pioneer species (usually Douglas-fir or lodgepole pine) is killed and the open¬ ings fill with shade-tolerant western hemlock or west¬ ern redcedar after Douglas-fir, or subalpine fir after lodgepole pine. These species are not markedly less susceptible to A. ostoyae but appear to be more tolerant, more frequently restricting infection to root lesions and butt rot (Morrison 1981). Williams and Marsden (1982) suggested a similar role for Armillaria and Phellinus zveirii in the succession on northern Idaho sites where western hemlock was climax. Disease is also evident in other forests undisturbed by human activity (Haig and others 1941, Wargo and Shaw 1985). Natural Forests 113 Armillaria Root Disease in Dry Sclerophyll Eucalypt Forests As a primary pathogen, A. luteobubalina affects many eucalypt and understory species in dry sclerophyll mixed-species eucalypt forests in central Victoria, and in karri and jarrah forests in southwestern Western Australia (Kile 1981, Kile and others 1983, Pearce and others 1986, Shearer and Tippett 1988). The affected forests occur between 300 m and 1,200 m altitude on soils of variable fertility, and receive annual rainfall of 700-1,200 mm. Most have a long history of logging. Hosts in these forests include at least 81 eucalypt, understory, and ground species (table 8.1). The evidence for the primary pathogenicity of A. luteobubalina includes the constant association of the fungus with disease, a pattern of contagion consistent with that for an organism dependent on a woody food base, a correlation between infection and symptom development in large trees, and pathogenicity of the fungus in pot and field inoculations of some host tree species (Kile 1981, Pearce and others 1986, Shearer and Tippett 1988). In Victorian forests, diseased trees tend to occur in roughly circular foci although the pattern of disease development is often obscured by multiple infection, cutting, and burning (fig. 8.7). Within patches, which may range from a few trees to 1 ha or more, the disease usually shows progressive outward expansion, with more recently dead and dying trees towards the mar¬ gin and older dead and often wind-thrown trees to¬ wards the center. The chronic nature of infection is apparent by the death of eucalypt or understory regen¬ eration that was established following death or re¬ moval of the previous strata. Typically, A. luteobubalina or other Armillaria species are not found in healthy forest surrounding diseased areas. Similar disease de¬ velopment occurs in jarrah forest. In karri forest, the disease is most active in young stands; with increasing stand age, mortality is restricted to suppressed or sub¬ dominant trees although larger trees may be infected (Pearce and others 1986). Young infected trees often die suddenly with a major proportion of their foliage intact. In contrast, large, mature trees generally show progressive crown die- back before eventual death. Some trees develop basal cankers from infection, which limit fungal spread and promote host survival (Kile 1981). The fungus forms few rhizomorphs in forest soils, and underground spread between hosts occurs via root contacts at an average rate of 1-1.5 m per annum (Kile 1983b). i >: thousand hectares of Australian eucalypt forest ■ rlously affected by the disease (Edgar and others FIGURE 8.7 — Aerial view of Armillaria luteobubalina root dis¬ ease center in dry sclerophyll eucalypt forest, Victoria, Australia 1976, Shearer and Tippett 1988). Edgar and others (1976) estimated mature stands with moderate to se¬ vere disease had respective sawlog increments of about one-half and two-thirds that of an average healthy stand, with growth losses of 0.3-2.0 m 3 per ha per year depending on site and disease severity. Besides these losses, scattered and small patch mortality is evident in regrowth stands. In 30-year-old regrowth messmate stringybark, with 51-75% of ground-level stem circum¬ ference infected by A. luteobubalina, average monthly girth increment was only 41% of that of healthy trees (Kile and others 1982). The wide distribution of A. luteobubalina in southern Australia and its intimate association with eucalypt forest communities indicate that it is indigenous. While Kile (1983a) reported evidence of its pathogenic activity in unlogged eucalypt forest, the greatest incidence and severity of disease has been observed in selectively logged forests (Edgar and others 1976). Strong relation¬ ships exist between infected stumps and disease inci¬ dence (Edgar and others 1976, Kellas and others 1987, Kile 1981, Pearce and others 1986). Though disease is endemic, logging apparently alters the balance be¬ tween host and pathogen toward more severe local epidemics. Armillaria Species as Secondary Pathogens Biotic or abiotic stress of natural forests or individual trees (see chapter 7) within them may transform indig¬ enous Armillaria species into vigorous secondary pathogens. This phenomenon is most notable in forests where, prior to stress onset, disease is restricted to epi- Natural Forests phytic associations, root lesions, and butt rot. This sec¬ ondary role has been recognized since early this cen¬ tury (Nechleba 1915; see also reviews by Day 1929 and Twarowski and Twarowska 1959) and has often domi¬ nated views of the pathogenic behavior of Armillaria species (Day 1929, Gremmen 1976). Virtually all historical reports of secondary pathogen¬ esis refer to A. mellea, but many other species also act in this manner. The identity and relative importance of species of different pathogenicity in broadscale second¬ ary diseases such as those shown in table 8.3 therefore require reappraisal. Although forest diebacks and declines are episodic diseases of varying etiology, all share a causal complex that begins when tissues of healthy trees are altered or predisposed by stress and culminates when those tis¬ sues are invaded and killed by facultative parasites (Houston 1973, 1982,1984,1987). Because infections by weakly pathogenic organisms are unsuccessful or re¬ stricted in the absence of stress, and because in the absence of these organisms trees usually recover with the abatement of stress, organisms of secondary action such as Armillaria species are an integral component of the disease syndromes. This does not imply, however, that stress alone cannot kill trees (Houston 1987). Stress factors include insect damage, primary patho¬ gens, drought, waterlogging, fire, temperature ex¬ tremes, air pollution, or silvicultural treatments. These stresses may be either protracted or relatively ephem¬ eral, and they may occur months or even years prior to eventual tree mortality. Not all stresses enhance patho¬ genic activity, however, and some air pollutants prob¬ ably have an adverse effect on the fungus itself, restricting its ability to take advantage of weakened hosts (Singh and Sidhu 1989). The prominent role of Armillaria species in diseases such as those shown in table 8.3 results from their ex¬ tensive natural distribution in the stress-affected forests and their primary infection of or epiphytic presence on many root systems prior to the advent of stress. The fungus is thereby able to take advantage of changed circumstances to spread quickly from existing infec¬ tions or establish new ones. For example, regarding dieback of regrowth messmate stringybark and moun¬ tain ash in Tasmania, A. hinnulea and/or A. novae- zelandiae usually infected a large proportion of each tree's root system at the time of death. Excavations, however, indicated that in healthy forest at least 70% of trees had minor root infections or epiphytic rhizomorphs (Kile 1980b, Kile and Watling 1983). In this and many other diebacks and declines, Armillaria is probably responsible for the ultimate death of many trees. Unlike lethal primary disease caused by Armillaria species, where dead and dying trees are usually clus¬ tered in expanding foci, the pattern of mortality in dieback and decline diseases is typically more variable, ranging from essentially random to more site or topo¬ graphically related patterns. Armillaria infection is less readily associated with identifiable food bases. The distribution of different Armillaria species may explain variations in infection and subsequent patterns of mor¬ tality, because species of different pathogenicity may invade root systems at different stages of host debilita¬ tion (Guillaumin and others 1989a). The susceptibility of individual trees or stands to infection will be miti¬ gated by site and soil factors and tree vigor. Experimental studies have demonstrated the increased susceptibility of various tree species to infection when stressed by defoliation, suppression, reduced light intensity, adverse soil moisture conditions, or nutrient supply (Davidson and Rishbeth 1988, Entry and others 1986, Ono 1970, Redfern 1978, Wahlstrom and Unestam 1989, Wargo 1972, Wargo and Houston 1974). In¬ creased susceptibility is related to biochemical changes in the host induced by stress, which lowers host resis¬ tance and stimulates development of the fungus. Indi¬ vidual stress factors and their effects on pathogenesis by Armillaria are fully discussed in chapter 7. Dispersal and Distribution The spatial development of Armillaria populations in natural forests ranges from the discontinuous distribu¬ tion of discrete genotypes of one species, to a mosaic of genotypes to which one or more species may contrib¬ ute. Through the infection of living hosts, stumps, and roots and the proliferation of rhizomorphs, the latter situation can be equated to a continuous distribution, although even in multi-species populations individual species may have restricted occurrences. Discontinuous distributions appear more typical of temperate, Medi¬ terranean, and tropical forests while continuous distri¬ butions are more evident, although not omni-present, in boreal and cool temperate forests. However, better quantification of spatial distributions are required in relation to both Armillaria species and forest type. In boreal and temperate forests, genotypes of different Armillaria species have been identified and mapped using alleles of the incompatibility (mating) genes as genetic markers or by intraspecific pairings of diploid forest isolates (see chapter 2). Dispersal and distribution occur via basidiospore infec¬ tions that create new infection foci and vegetative growth that expands the local distributions of particu¬ lar genotypes. Local expansion may proceed by Natural Forests 115 TABLE 8.3 — Examples of diebacks and declines in natural forests in which Armillaria species were recognized as important secondary pathogens.* Disease and primary host Major initiating stress Location Time frame Reference Alaska yellow-cedar d\eback(Chaemaecyparis nootkatensis Unknown Southeast Alaska early 1900s to present Frear (1982) Shaw and others (1985) Hennon and others (1990) Dieback and mortality of coniferous species a,c Eastern Canada late 1960s- early 1980s Hudak and Singh (1970) Hudak and Wells (1974) Raske and Sutton (1986) Birch dieback (Betula alleghanensis) a,c,f,g Northeastern North America mid 1930s- late 1950s Spaulding and MacAloney (1931) Oak declines (Quercus species) a,c,d 1) Europe regional occurrences during this century Baumgarten (1912) Hen (1914) Falck (1918) Georgevitch (1926b) Day (1927a) Stalina (1954) Petrescu (1974) Guillaumin and others (1983) a,c,d,g 2) Midwest and eastern USA Macaire (1984) see reviews by Staley (1965) and Houston (1987) Wargo (1977) Ohia decline (Meterosideros polymorpha) b,e Hawaii mid 1950s- early 1970s Laemmlen and Bega (1974) Hodges and others (1986) Pole blight (Pinus monticola) a Western USA British Columbia 1930s-1950s Hubert (1950, 1953) Leaphart and others (1957) Regrowth dieback a,c (Eucalyptus regnans, E. obligua) Tasmania regional occurrences early 1960s-present Kile (1980b) Sugar maple declines (Acer saccharum) a,c,f Eastern North America regional occurrences 1950s-present Houston and Kuntz (1964) Wargo and Houston (1974) * The derivation of the table arrange¬ ment and some data from Houston (1987) is acknowledged. Stress factors a water deficit/high temperature b poor drainage/water excess c defoliation by insects d defoliation or damage by fungi e nutrient imbalance f logging disturbance g low temperature damage rhizomorphs or mycelial growth through and between contacting root systems. The role of basidiospores as inoculum and the involvement of rhizomorphs in spread and infection are considered fully in chapter 4, and comment here is restricted to points particularly relevant to these processes in natural forests. Basidiospores igh the potential epidemiological importance of lore infection has long been recognized (Boyce 1919, Rishbeth 1964), the evidence for its u natural forests remains circumstantial based on the detection of multiple genotypes and unique combinations of mating alleles (Kile 1983b, Korhonen 1978). As with many other macromycetes, spore production by Armillaria basidiomes may be prolific and the period for potential basidiospore infection relatively long. Rishbeth (1970) recorded deposition rates of up to 1,000 viable basidiospores per dm : per min. close to basidiomes. Basidiospores have also been trapped from the air on screens and on freshly cut wood (Hood and Sandberg 1987, Molin and Rennerfelt 1959, Rishbeth 1970, Swift 1972). Basidiome production of some 116 Natural Forests Armillaria species may extend over several months (Fedorov and Bobko 1989, Kile and Watling 1981, Pearce and others 1986, Rishbeth 1970, Shubin 1976). Shaw (1981a) found basidiospores could remain viable on the outer bark of conifers over an Alaskan winter. Environmental factors, particularly moisture, host, and Armiliaria species, may influence spread and infection by basidiospores (Rishbeth 1970). In the relatively dry inte¬ rior forests of western North America, few genotypes of A. ostoyae, and the large areas occupied by some of them, suggest limited opportunities for basidiospore infection (Shaw and Roth 1976). Similarly, in dry sclerophyll euca- lypt forest in Victoria, where 36 genotypes were found in 24 ha, Kile (1983b) estimated the rate of basidiospore infection for A. luteobubalina could average less than one per year. In wet sclerophyll eucalypt forest in Tasmania, 46 genotypes of A. hinnulea were isolated in 1.1 ha, sug¬ gesting relatively frequent basidiospore infection in this forest type (Kile 1986). A situation comparable to the latter probably exists in deciduous forests in the North¬ eastern United States, Finnish coniferous forests, and New Zealand hardwood-podocarp forests where mois¬ ture conditions are favorable for frequent and abundant basidiome production (Anderson and Ullrich 1979, Hood and Sandberg 1987, Korhonen 1978). Because of our poor knowledge of basidiospore infection courts and the potentially large number of factors which may influence the incidence of basidiospore infection, further experimental studies of the process are needed. Rhizomorphs and Root Contacts Both rhizomorphs and root contacts are important for spread and infection in natural forests. The actual contri¬ bution of rhizomorphs to these two processes probably depehds on the forest environment and the characteris¬ tics of the rhizomorphs of the particular Armillaria spe¬ cies (see chapters 4 and 6), but they are not obligatory for either spread or infection. Infection via root contact has received little attention in the past, but its efficiency in some forests suggests it could contribute to spread even in situations where rhizomorphs are present. Armillaria luteobubalina spreads almost exclusively by this means in Australian eucalypt forests even though it produces rhizomorphs on agar and in pot culture (Kile 1981, Morrison 1989, Pearce and others 1986, Podger and others 1978, Shearer and Tippett 1988). Rhizomorphs also are formed rarely in many tropical forests (Butler 1928; Dade 1927; Swift 1968, 1972). The lack of rhizomorphs in native forest soils has been attributed to unfavorable physical environments for their initiation and growth (Pearce and Malajczuk 1990a, Rishbeth 1968) or to inhibitory compounds in the soil (Olembo 1972, Swift 1968). Pearce and Malajczuk (1990a) showed limited rhizomorph development of A. luteobubalina in karri forest soils in Western Australia related to unsuitable combinations of soil temperature and soil moisture levels for rhizomorph growth during much of the year. Whether this explanation is adequate in other forests where rhizomorphs are absent remains to be determined. Spatial Distributions Over time, dispersal processes contribute to varied patterns of genotype and species distribution. These patterns appear to form a continuum from the simple to the complex depending on the frequency of new infections and the number of Armillaria species. In co¬ niferous forests in western North America and dry sclerophyll eucalypt forests in Victoria, relatively few genotypes of A. ostoyae and A. luteobubalina, respec¬ tively, may develop large, discrete, clones (2-3 ha or more) with individual genotypes occurring hundreds of meters apart (Adams 1974, Anderson and others 1979, Hood and Morrison 1984, Kile 1983b, Shaw and Roth 1976) (fig. 8.8). The size of some clones and the discontinuous distribution of some genotypes when compared with rates of spread is taken as evidence that the original infections may have begun decades or even several centuries previously (Kile 1983b, Shaw and Roth 1976). In contrast, A. borealis, A. ostoyae, A. gallica, A. cepistipes, A. mellea, A. hinnulea, A. limonea, and A. novae-zelandiae form relatively small clones (approx. 50- 100 m maximum distance between isolates of the same genotype) in a variety of coniferous and moist temper¬ ate hardwood forests. A relatively frequent number of genotypes occurs per unit area, often in close proximity or intermingling and possibly with clones of more than one species (Hood and Sandberg 1987, Kile 1986, Korhonen 1978, McDonald and Martin 1988, Rishbeth 1978b, Thompson 1984, Ullrich and Anderson 1978). Once established in the forest, any given genotype may persist for decades or centuries, occupying successive woody substrates as a result of parasitic and sapro¬ phytic activity. Multiple genotypes form a type of sub¬ terranean mosaic, but the physical and biochemical interaction between intra- or inter-specific genotypes has received limited investigation (Mohammed and Guillaumin 1989). These kinds of interactions could be mediated by extracellular structures such as those ob¬ served in Postia placenta (Fr.) M. Lars, et Lomb. (Green and others 1989), although they have not been ob¬ served in Armillaria. Studies of dispersal and distribution emphasize the dynamic nature of the interaction between Armillaria species and natural forest ecosystems. Kile (1983b) Natural Forests 117 FIGURE 8.8 — Occurrence of seven genotypes of Armillaria luteobubalina in a eucalypt forest in the 33 ha Victoria Mill Scenic Reserve, Mount Cole State Forest, Victoria, Australia. (From Kile 1983b, courtesy Australian Journal of Botany) considered that the major factors influencing clonal development were likely to be: the number and loca¬ tion of existing and new infections, the pathogenicity of the individual genotypes, their longevity in individual food bases, the stand and tree characteristics including host resistance, and perturbations within the forest such as fire and logging. Combinations of such factors over long periods could account for the limited size of some clones, the extensive or dispersed distribution of others, the presence of multiple genotypes in the same area, contraction or expansion of clone size, differing disease intensity, and predictably the loss of some genotypes from the forest. Pathogenicity, Environment, Host Resistance, and Primary Disease Expression As in other plant diseases, the expression of Armillaria root disease is influenced by species pathogenicity, host resistance, and environmental factors. How these factors interact relative to primary disease in natural forests is poorly understood at the present time. A general observation can be made from the disease reports considered in this chapter: while non-lethal primary disease may be common in relatively wetter boreal and temperate forests (which may be translated into lethal secondary infection by stress), the most seri¬ ous primary disease occurs in relatively drier Mediter¬ ranean or continental forests. As there is no evidence bat recent introductions of pathogenic species are ■risible for disease, the current situation presum¬ ably reflects the results of long-term coevolution of hosts and pathogens. One explanation for a difference of this nature in dis¬ ease epidemiology encompasses the pathogenicity of the indigenous Armillaria species and the environmen¬ tal and biological factors which control the population of the fungus. The large food base which may develop in wet forests in mild climates is seemingly balanced bv the weak pathogenicity of the resident Armillaria spe¬ cies. In harsher forest environments, stress may have selected species or genotypes of greater pathogenicity which can effectively maintain themselves in the forest community from a more limited food base. In the former forests, weak pathogenicity, wide distribution, and long survival in inoculum are the elements of mu¬ tual coexistence (a feature of K-selected organisms). In the latter forests, greater pathogenicity, discontinuous distributions, and shorter survival in inoculum achieve the same end. In neither situation is the survival of the host species threatened. When stress leads to secondary Armillaria infection, not all trees are killed and such events typically lead to the establishment of regenera¬ tion. These concepts, represented in fig. 8.9, can be illus¬ trated by Australian examples. In Tasmania's wet sclerophyll eucalypt forest, A. hinnulea / A. novae- zelandiae are almost ubiquitous on root systems as epi¬ phytic rhizomorphs or root lesions (Kile 1980b, Kile and Watling 1983). Logging or wildfire creates a vast food base and results in a high incidence of massive root and stump infection but with virtually no mortal¬ ity among the regenerating stand of eucalypts and other species. Disease is restricted to the same form as that in the pre-existing stand (Kile 1980b). In Victoria's dry sclerophyll forest, on the other hand, eucalypts and other species are killed by A. luteobubalina even in the absence of artificial disturbance (Kile 1981,1983b). However, food bases are fewer and the fungus gener¬ ally only survives in stumps for 15-25 years (Kile 1981). The relative pathogenicity of different Armillaria spe¬ cies on the same host under controlled conditions can be ranked (Morrison 1989). Variable disease expression on the same host supports the view that the differences described for eucalypt forests are to a significant de¬ gree caused by interspecific differences in pathogenic¬ ity rather than by host or short-term environmental effects. Messmate stringybark is a common host in both forest types, yet is only killed where A. luteobubalina occurs (Kile 1980b, 1981). Direct pathogenicity com¬ parisons of the three Armillaria species present in the forests have not been made on messmate stringybark, but those tests undertaken with the individual species support the view that A. luteobubalina is inherently more pathogenic than either A. hinnulea or A. novae- 118 Natural Forests DRY FORESTS Limited inoculum and saprophytic - survival Restricted distribution Root and stem lesions Primary pathogenicity (low host resistance) / -/-► Tree death / / / Increased inoculum Increased disease WET FORESTS showing a strong tendency to decrease as stand pro¬ ductivity increased (fig. 8.9). They further suggested that disease incidence is also greatest in host popula¬ tions in transition zones between climax species. These are seen as being less adapted to the site and therefore more vulnerable through lower host resistance. Soil factors may also be influential (Shields and Hobbs 1979, Williams and Marsden 1982). Intraspecific varia¬ tion in host resistance has not been investigated rela¬ tive to primary disease but could help explain regional differences in intraspecific pathogenicity. Presently, there is little basis on which to judge the merits of these different hypotheses, but they lend themselves to cre¬ ative experimentation. Abundant inoculum Long saprophytic survival Wide distribution Weak pathogenicity (high host resistance) Epiphytic associations, root lesions, stem cankers butt rot Tree death Progressive infection ■ Stress (abiotic or biotic including senescence) Increased inoculum may increase disease but does not cause lethal disease Possible feedback mechanisms such as that proposed for mountain hemlock stands infected by Phellinus weirii may also operate to influence host resistance during stand development. Waring and others (1987) suggested pathogen-induced disturbance may increase nutrient and light availability following death of ma¬ ture stands, and increase resistance of young trees against infection. Although Shaw and others (1988) challenged this assumption and suggested vigorously growing young trees would have more roots and thus an increased probability of infection, such controversy does not deny the possible existence of such effects. FIGURE 8.9—Conceptual models of the pathogenic behavior of Armillaria species in terms of two general forest environments. zelcuidiae (Kile 1980b, 1981; Morrison 1989; Podger and others 1978). Intra-regional differences in the epidemiology of dis¬ ease caused by the same Armillaria species on the same or related hosts have been observed. In western North America, A. ostoyae is an aggressive pathogen on inte¬ rior forest species but usually only causes minor dis¬ ease on many of the same hosts in wet coastal forests (Morrison 1981, Morrison and others 1985a, Wargo and Shaw 1985). A similar situation may occur with A. luteobubalina in Western Australian forests. Shearer and Tippett (1988) noted that host mortality following in¬ fection was greater in the intermediate and low rainfall zones of the eastern jarrah forest than in the higher rainfall zones to the west. The fungus also appears to be less damaging in the wetter karri forest in the same region (Pearce and others 1986). For western North America, Morrison and others (1985a) considered these differences might result from intraspecific variation in pathogenicity between coastal and interior isolates of A. ostoyae. McDonald and others (1987a,b) proposed that the difference in pathogenic behavior is linked to site productivity, host adaptation, or stress, with the incidence of pathogenic behavior Disease expression in natural forests is clearly a com¬ plex phenomenon. Establishing the significance of host, pathogen, and environmental moderation in disease expression is a major challenge for future research. These interactions and consequent disease expression are in turn modified by forest management practices. Forest Management and Disease Forest harvesting and other disturbances cause fluc¬ tuations in inoculum levels in natural forests (Kile 1980b). In forests where the major Armillaria disease effects are non-lethal (root lesions, minor root rot, and butt rot), these fluctuations appear to have little effect on disease epidemiology from crop to crop unless man¬ agement severely stresses residual trees. Where Armillaria species are aggressive primary pathogens, such as in the situations already described, manage¬ ment practices such as logging and control of fire se¬ verity and frequency may significantly affect disease expression. These may occur directly through impact on inoculum levels or interactions with species compo¬ sition and stand structure. Much of the available infor¬ mation is observational, however, and little experimental study has been done on the effect of man¬ agement practice on disease levels. Selective logging in old-growth or mature-regrowth stands may intensify disease development in the re- Natural Forests 119 sidual stand. Damage is dependent on stand age and species and is usually attributed to increased inoculum, but physiological stress from exposure in retained trees could also be of some importance in increased tree susceptibility (see chapter 7). Logging old-growth pon- derosa pine in southern Washington State led to strik¬ ing disease development in young trees, with zones of dead and dying seedlings and saplings surrounding the infected old-growth stumps and eventually leading to the creation of an open area (Shaw 1975, Shaw and others 1976a). Harvesting, particularly selective log¬ ging, has also led to inoculum buildup on many species of conifer stumps in other western North American forests (Byler and others 1990, Filip and Goheen 1982, Hagle and Goheen 1988). Severe Armillaria root dis¬ ease in dry sclerophyll eucalypt forest in Victoria, Aus¬ tralia was associated with repeated (approximately 10-year) selection cutting of the larger trees (Edgar and others 1976). Subsequently, Kellas and others (1987) showed that cutting intensity per se did not affect dis¬ ease incidence, but that frequency of cutting within infected forests is probably the critical factor promoting disease development. Regular creation of stumps in¬ creased inoculum levels and the probability of residual trees being in close proximity to inoculum, thereby altering the balance in favor of the pathogen. Unless A. luteobubalina can access and infect stumps within 3-4 years of cutting, it is excluded from colonization by other microorganisms (Kile 1981). Clearfelling in dis¬ ease patches could therefore be a better management practice by reducing the number of stumps infected and the disease level in the subsequent crop. Partial cutting practices may make stands more suscep¬ tible to disease through changes in species composi¬ tion. Selective logging in mixed-conifer forests in western North America, particularly those where spe¬ cies of pine and larch predominate, can favor regenera¬ tion of the more root-disease-susceptible Douglas-fir and true fir (Byler and others 1990, Filip and Goheen 1982, Hadfield 1984, Hagle and Goheen 1988, Shaw and others 1976a). Such changes may not always be adverse, however. Shelterwood cutting in dry sclerophyll eucalypt forest favors regeneration by broad-leaved and narrow-leaved peppermint as op¬ posed to messmate stringybark, the commercially pre¬ ferred species (Kellas and others 1987). The former species have a similar susceptibility to A. luteobubalina. It may be feasible to space or commercially thin young, even-aged regrowth stands without increasing levels of Armillaria root disease if small stumps do not provide a sufficient food base to establish new disease foci, or tree vigor is enhanced sufficiently to resist root disease. Filip and others (1989) and Johnson and Thompson < 1975) found no adverse effects on stocking 20 years thinning in a young ponderosa pine stand, and although Koenigs (1969) found thinning in an 80-year- old released understory stand of western redcedar increased root rot, disease was also apparently influ¬ enced by other stress factors. Precommercial thinning is generally not recommended in Armi/laria-infected Douglas-fir regeneration (Morrison 1981). Further ex¬ perimental studies on effects of spacing and thinning in relation to species composition appear necessary, how¬ ever, before firm conclusions can be made. Fire management can influence the susceptibility of forest stands to disease. Fire may directly affect Armillaria activity in forests through destruction of inoculum or indirectly through stress effects on the fungal mycelium which lead to natural biocontrol (Reaves and others 1990). Although a reduction in dis¬ ease has not been demonstrated, fire frequency and intensity may also be a major determinant of the sus¬ ceptibility of stands to disease through its influence on tree vigor, species composition, and stand structure. No known studies quantify the effect of fire on inocu¬ lum quantity and viability, although Hood and Sandberg (1989) reported reduced isolation success from rhizomorphs on a clearcut native forest site after burning. General observations suggest a significant direct effect on inoculum levels is only achieved by high-intensity fire which burns or chars stumps and major buttress and lateral roots (Kile 1980b, 1981). Even then it is likely a significant proportion of below¬ ground inoculum will escape direct effects. Munnecke and others (1-976) showed that heating se¬ verely weakens the vitality of Armillaria mycelium, rendering it susceptible to parasitism by Trichoderma viride Pers.:Fr. and other soil-inhabiting fungi. Similar effects may operate to reduce inoculum in burned for¬ ests. Reaves and others (1990) found isolates of Trichoderma species from burned and non-burned soils beneath a ponderosa pine forest in Oregon were an¬ tagonistic to A. ostoyae, reducing colony growth and rhizomorph formation in culture. Isolates from burned soils were more antagonistic than those from non- burned soils as fire favored the growth of more antago¬ nistic Trichoderma species. In the same situation, ash leachates inhibited growth of A. ostoyae in vitro while having a positive effect on Trichoderma (Reaves and others 1984, 1990). While appropriate use of fire may be effective in elevating populations of Trichoderma that are antagonistic to Armillaria, the mechanism, extent, and persistence of such effects need clarification. In forest types in which burning has been a determi¬ nant of species composition and stand characteristics, fire suppression or exclusion may interact with silvicul¬ tural management to promote Armillaria root disease by allowing regeneration of species which are more 120 Natural Forests susceptible to Armillaria (Byler 1984, Byler and others 1990, Filip and Goheen 1982, Hagle and Goheen 1988, Shaw and others 1976a). Fire control associated with selective logging in drier coniferous forests in western North America has favored regeneration and over¬ stocking of Douglas-fir and true fir in stands formerly composed predominantly of ponderosa and white pine and western larch, species apparently less susceptible or more tolerant to root disease. A need exists for more careful consideration of long-term ecological effects induced by various stand treatments on Armillaria root disease. A better understanding of such effects could lead to refinement of silvicultural methods. While not a direct management effect, introduction of white pine blister rust ( Cronartium ribicoln J.C. Fisch.) into the northern Rocky Mountains of western North America has likely enhanced Armillaria- caused mortal¬ ity (Byler and others 1990). The rust epidemic emulated a partial cut by killing large numbers of pole-sized and larger western white pines in many individual stands. It also modified succession in new stands by reducing or eliminating western white pine regeneration. A dra¬ matic shift in species composition of many stands from the tolerant western white pine to susceptible Douglas- fir and grand fir was one result. The rapid killing of larger trees probably contributed to inoculum buildup as well. A related problem was the application of many western white pine salvage cuts due solely to the threat of C. ribicola. Apart from direct management effects on quantity of inoculum or species composition, practices which se¬ verely stress plants may also increase disease. Armillaria species are thus a potential hazard for inten¬ sively managed coppice forests. Incidence and extent of root rot increased with shortened rotation in quaking aspen and bigtooth aspen sucker stands in Ontario and Wisconsin, and sucker numbers declined as a greater proportion of stumps were invaded by the fungus in successive rotations (Stanosz and Patton 1987a,b; Shell and Berry 1986). Chronic low-level primary disease in the natural forest was transformed into more progres¬ sive secondary infection of physiologically stressed stumps and root systems by this type of management system. Conclusions Armillaria species are remarkably successful compo¬ nents of many natural forests. A large proportion of tree and shrub species of different strata, particularly in boreal and temperate forests, may be susceptible to Armillaria infection. Besides a pathogenic role, mem¬ bers of the genus contribute significantly to decomposi¬ tion and mineral cycling as well as playing minor roles as mycoparasites and mycotrophic associates with some achlorophyllous plants. Armillaria in natural forests is endemic, evidence of disease is often obscure, and it may often have minimal effect on host health and growth. Flowever, a con¬ tinuum of disease effects from non-lethal to lethal in¬ volves Armillaria species as primary or secondary pathogens. Epidemic disease involving Armillaria in either role may result when the balance between patho¬ genicity and host resistance is altered by stress or dis¬ turbance. Many forests may be utilized without aggravating the endemic disease level. In others, dis¬ turbance such as logging may lead to a major imbal¬ ance between host and pathogen. Inappropriate management in some regions has created a heritage of root disease problems. Our understanding of the ecology and dynamic behav¬ ior of Armillaria in natural forests has developed sig¬ nificantly in recent years through better knowledge of species identity, pathogenicity, ecology, and a clear recognition of primary and secondary modes of behav¬ ior. In one case at least, it is now feasible to integrate this knowledge through a computer simulation model to better understand root disease dynamics and their response to forest management treatments (see chapter 10). Significant research needs remain. Specifically, future investigations should examine inoculum buildup, quantifying pathogen spatial distributions and dynamics, the relative importance of host, patho¬ gen, and environment on disease expression, and the ecological effects of disease. Natural Forests 121 CHAPTER 9 Armillaria in Planted Hosts Ian A. Hood, Derek B. Redfern, and Glen A. Kile I n chapter 8, Armillaria was examined in its natu¬ ral forest environment, modified or not by the activities of man. We now look at the disease in the various artificial habitats created when select¬ ed hosts are cultivated either for commercial produc¬ tion or as ornamentals for their aesthetic appeal. The distinction between natural and artificial environments is not always a sharp one, as when seedlings are plant¬ ed beneath a natural overstory or when new planta¬ tions are infiltrated by seed regeneration from nearby stands. Indeed, it is sometimes difficult to decide if a plant growing within (or even outside) its natural range has been planted or naturally seeded. Even so, the planted host generally occurs in a very different setting from its natural counterpart which is adapted to its own particular ecosystem. We might therefore ex¬ pect Armillaria to be frequent and widespread in many species of cultivated plants (Day 1929, Garrett 1956a, Mallet and others 1985). To demonstrate just how widespread attack by Armill¬ aria really is, a broad overview of the disease's geo¬ graphic distribution begins the chapter. Sixty years of records are used to assess the global importance of Armillaria on cultivated plants. An account of the de¬ velopment and impact of Armillaria in plantations then follows, and the chapter is concluded by discussing how various management procedures may affect dis¬ ease development. Distribution and importance The literature on Armillaria in planted crops and orna¬ mentals is vast. A large selection of reports has been collated and summarized in tables 9.1 (p. 140) and 9.2 (p. 142), and in fig. 9.1. Together, these present a broad picture of the disease in various host groups through¬ out the world (for host species lists see, for instance, Spaulding 1961, Raabe 1962a, Hansbrough 1964, Browne 1968). The number of references in these tables . ’ : ates the importance of the disease for particular n different countries, with some qualifications: may reflect the regional activity of plant pa¬ thologists besides that of the disease, and may be bi¬ ased toward hosts of greater economic importance. The tables indicate only the presence of the disease, not its severity, which is often expressed only in qualitative terms. Despite these drawbacks, the figure and tables summarize where Armillaria occurs in planted hosts throughout the world. Several trends may be detected in tables 9.1 and 9.2, and fig. 9.1. For example, Armillaria is not listed on any major cereals (wheat, rice, maize, oats, barley, rye) or certain other cultivated food crops (peas, beans, groundnut) which are normally cultivated on arable land but rarely, if ever, planted on former forest sites (see later). Records appear weighted in favor of cash- crop plants present in commercial plantations rather than non-commercial hosts used in subsistence crop¬ ping or shifting agriculture on cleared forest land. Records for ornamental trees and shrubs also seem under-represented and somewhat fragmentary. Proba¬ bly many occurrences of the disease in amenity plant¬ ings go unreported or are listed only in unpublished records of state agricultural agencies and experiment stations. Figure 9.1 clearly shows that Armillaria is widespread in planted hosts throughout temperate regions and in much of the tropics. Detailed information on disease occurrence in particular regions is normally unavail¬ able but occasionally does exist. For instance, figure 9.2 shows the distribution of 574 Armillaria-caused deaths of mainly planted hosts in southern Britain during the past 25 years. Although the frequency of records is biased toward the collection center, and is influenced by the uneven distribution of parks or gardens and former woodland sites, results indicate a widespread occurrence of disease within the survey region. No doubt intensive long-term surveys would present a similar picture where conditions favor the disease else¬ where in the world (Hood 1989). Lack of detailed information on disease incidence pre¬ vents precise comparisons between regions. However, 122 Planted Hosts Host Type » Conifer 9 Broadleaf (forest, shade, ornamental) E Economic host (except forest). FIGURE 9.1 — Distribution of recorded Armillaria attacks in planted hosts, by country (and region). the literature suggests that the disease is most common in areas with a moist environment and a moderate tem¬ perature range (Bliss 1946; Browne 1968; Bunting 1924; Fox 1964; Gibson 1961,1975,1979; Ivory 1987; Jie 1982; Jorge 1977; Kile 1980a; Longenecker and others 1975; Manka 1980; Mohammed and others 1989; Rivera 1940; Rudd-Jones 1950; Rykowski 1980; Sokolov 1971; Tarry 1969). Precipitation and temperature may be the prima¬ ry factors governing both the altitudinal and latitudinal distributions of Armillaria. Thus, in many tropical areas the disease is known only in plantations established at higher elevations where the climate is cooler and wetter (Arentz and Simpson 1989, Barnard and Beveridge 1957, Bernard 1926, Brazilian Inst. Forestry Dev. 1976, Fox 1970, Gill 1963, Ivory 1975, Raabe and Trujillo 1963, Rayner 1959, Satyanarayana and others 1982, van der Goot 1937). This contrasts with certain other tropical root disease fungi which are found mainly in planta¬ tions growing at low and mid elevations where the climate is hotter ( Phellinus noxius (Corner) Cunning¬ ham, Rigidoporus lignosus (Klotzsh) Imazeki, Fox 1970). In the temperate zones, Armillaria attacks plantations established at low and mid altitudes, but not those planted at higher elevations where it is too cold (John¬ son 1976, Rahm 1956, Singh and Khan 1979, Twarowski and Twarowska 1959). In the same way, the disease may occur less often at higher latitudes where the cli¬ mate is colder. Armillaria also appears to be absent from certain inland regions with extreme continental climates, as in parts of the Soviet Union (Sokolov 1964). Europe and the Soviet Union Europe has a tradition of plantation forestry dating back at least 3 centuries. Armillaria is widespread in forest plantations on this continent, and few countries, if any, lack disease records (tables 9.1, 9.2a,b; Day 1929; fig. 9.1). Reports are particularly numerous from France (see also Boullard 1961), Germany, Poland, and the United Kingdom. Armillaria is widespread in plant¬ ed forests in the Soviet Union, (Fedorov and Bobko 1989, Fedorov and Poleschuk 1981, Fedorov and Smol- jak 1989, Sokolov 1964) and has also been found on eucalypt species in Spain, Portugal, and Cyprus (Gib¬ son 1975, Ivory 1987). Reports are more numerous for conifer than hardwood stands (tables 9.1, 9.2a). Much fundamental research has been undertaken in European plantations since Robert Hartig first estab¬ lished the relationship between Armillaria and disease Planted Hosts 123 FIGURE 9.2 — Distribution of deaths in planted hosts due to Armillaria in southern Britain, 1962-1986. Dot diameter indicates number of records (respectively, over 50, 20-50, IQ- 20, under 10, in decreasing order of size; data courtesy Pathology Branch Advisory Service, UK Forestry Commission). in Germany in 1873 (Hartig 1873b, 1874; Nechleba 1915), and this has contributed greatly to our knowl¬ edge of the disease's nature and development world¬ wide. Until recently, most disease records in Europe were attributed to A. mellea (sensu Into), and sometimes A. tabescens. We now know (see chapters 1, 2) that at¬ tack in European plantations is caused predominantly by A. mellea (sensu stricto) and A. ostoyae (Korhonen 1978, reviewed Roll-Hansen 1985). This knowledge has encouraged research to define the ecological roles and behavior of the European Armillaria species (Guillau- min and Berthelay 1981; Guillaumin and Lung 1985; Guillaumin and others 1984, 1985, 1989a; Holdenrieder 1986; Rishbeth 1982,1983, 1985a,b, 1987; Siepmann 1985). Recent work shows that at least in western Eu¬ rope vigorous trees in pure conifer plantations are at¬ tacked mainly by A. ostoyae. Most infection in planted hardwoods (forest and ornamental trees, table 9.2a) is due to A. mellea; in stressed hosts, several less patho¬ genic species are sometimes responsible (e.g., A. gallica; Clancy and Lacey 1986; Davidson and Rishbeth 1988; Durrieu and others 1985; Guyon and others 1985; Intini 1989a,b; Laville and Vogel 1984; Lung-Escarmant and Taris 1985,1989). More severe attacks in conifers usual¬ ly occur on former natural hardwood forest sites (often oak) rather than pure conifer stands (Peace 1962, Red- fern 1975, Rishbeth 1982, Uscuplic 1980), so that the precise manner in which A. ostoyae invades and devel¬ ops in new conifer plantations requires elucidation (Guillaumin and others 1989a). Guillaumin and Lung (1985) indicated, however, that A. ostoyae can grow and uce rhizomorphs quite successfully on hardwood 124 species even though it is a parasite primarily on conifers (see also Davidson and Rishbeth 1988, Gregory 1989, Redfern 1975, Rishbeth 1982). Although Armillaria is considered Europe's most impor¬ tant root disease after Heterobasidion annosum (Fr.) Bref., its impact on forest plantations is comparatively minor in this region (Peace 1962). It creates canopy gaps and lessens the returns from early thinnings; but in general, the effects of early mortality are probably over-estimat¬ ed due to the rather spectacular appearance of the dis¬ ease in young stands (Pegler and Gibson 1972). Under certain stand or site conditions, damage can become more severe on some hosts. For instance, in Poland Ar- millaria has been responsible for serious losses of Nor¬ way spruce and Scots pine (Manka 1980,1981; Twarowski and Twarowska 1959; table 9.1). Armillaria kills ornamental trees and shrubs planted in gardens, parks, and reserves (table 9.2b; Ingelstrom 1938) and reports are particularly numerous from Great Britain (Boughey 1938, Gibbs and Greig 1990, Greig and Strouts 1983, Peace 1962, Rishbeth 1983, Schilling 1989). The records of mortality shown in fig. 9.2 were predom¬ inantly from amenity plantings and were mainly on hosts in the families Pinaceae (31% of records), Rosaceae (21%>), Fagaceae (13%), Oleaceae (12%), and Salicaceae ( 10 %). Food production is far older than plantation forestry, and losses from Armillaria must have occurred histori¬ cally wherever people cultivated plants. Today, the disease commonly attacks many European crops, partic¬ ularly pip fruit ( Citrus , Mediterranean countries), pome fruit (Mains, Pyrus), stone fruit ( Primus , Guillaumin 1977, Guillaumin and Pierson 1983, fig. 9.3), walnuts, and grapes (Guillaumin 1986b, see table 9.2b and Guil¬ laumin 1982, Guillaumin and others 1982, fig. 9.4a). The FIGURE 9.3 — Mortality gap in almond orchard (on peach rootstock) caused by A. mellea with trees dying at the margin; Aigaliers, southern France. (D. Barrett) Planted Hosts B FIGURE 9.4 — A: Mortality gap in grape vineyard, caused by A. mellea near Bordeaux, France. (J.-J. Guillaumin). B: Group of trees killed by Armillaria sp. in 1 5-year-old plantation of slash pine; Usa, Tanzania. (M.H. Ivory) disease has been reported less frequently in cane fruits (Marsh 1952), cork oak, fig, and flower crops (Guillau¬ min and others 1982), hazelnut, hops, kiwifruit, mul¬ berry, olive, strawberry, various vegetable crops, and in an Opuntia cactus crop (grown for its edible fruits). Armillaria also attacks various cultivated crops in Lithuania and the Ukraine (Dist. of Plant Dis. 1980), and in Azerbaijan, Armenia, Georgia, Belorussia, Tatar, and eastern Kazakhstan (Sokolov 1964). Details of Armillaria in European plantations have been reviewed by Peace (1962), Pawsey (1973), Schbnhar (1977), Rykowski (1980), Guillaumin (1982,1988), Guil¬ laumin and others (1982), Roll-Hansen (1985), and Phillips (1988). Greig and Strouts (1983) give a popular account of the disease in Britain. North America The behavior of Armillaria in both managed and un¬ managed forests has been extensively researched in North America since early this century (Byler 1984, see chapter 8). In many of these stands, Armillaria occurs as an important butt-rot agent and also influences forest successional development by killing seedlings, sap¬ lings, and more mature trees, particularly those already weakened by other causes (Byler 1984; Wargo 1980b, 1984b). Most reports of disease in forest plantings are from coniferous stands (table 9.1). Records on hard¬ woods have come mainly from ornamental or shade trees in the eastern half of the continent (table 9.2a). In conifer plantations in the west, Armillaria is one of sev¬ eral important root- and butt-rot disease fungi such as Phellinus iveirii (Murr.) Gilb. and H. annosum, which often occur in the same host or stand. Until recently, all North American records of Armillaria were attributed to A. mellea (or A. tabescens, see below). At least nine biological species of Armillaria are now known on the continent, some of which are related to European spe¬ cies (see chapters 1, 2). As in Europe, work is currently underway to define the ecological roles of these species and to identify those which cause disease in planta¬ tions and managed stands (Mallet and Hiratsuka 1988, McDonald and Martin 1988, Proffer and others 1987). Present forests in western North America originated largely by natural seeding following logging of the old growth forests that began during the 19th century. Planting is now carried out to improve stocking levels of desirable species. Early Armillaria research in plant¬ ed forests was done in coastal British Columbia, where trees in young plantations of Douglas-fir are killed by a species now identified as A. ostoyae (Bloomberg 1990; Buckland 1953; Hood and Morrison 1984; Johnson and others 1972; Morrison and others 1985a,b; Pielou and Foster 1962). Morrison (1981) reviewed the current understanding of the disease in this province. He con¬ sidered Armillaria to be comparatively unimportant in coastal conifer plantations because mortality is low (1-5%), and ceases after about age 25 years. By contrast, interior British Columbia experiences higher mortality in both plantations and natural, mixed conifer forests (Bloomberg and Morrison 1989; Morrison 1981; Morri- Planted Hosts 125 son and others 1988). In neighboring Alberta and other prairie provinces of Canada, disease impact has not been great, and mortality from Armillaria has occurred mainly in naturally regenerated conifers (Blenis and others 1987, Hiratsuka 1987, Mallet 1989). However, deaths also occur in plantations (Emond and Cerezke 1990, Hiratsuka 1987), and Armillaria root disease may become more common as management intensifies (Blenis and others 1987). Preliminary work suggests that the species pathogenic to conifers in Alberta are A. ostoyae, a form of A. cepistipes, and possibly A. mellea (Blenis and others 1987, Mallet 1989, Mallet and Hirat¬ suka 1988). In Europe, A. cepistipes is not considered to be a serious parasite (see chapter 6), and A. mellea is most important on hardwood hosts (Davidson and Rishbeth 1988, Guillaumin and others 1989a). Armillaria root disease (apparently A. ostoyae , Filip 1989a, Hadfield and others 1986, Morrison and others 1989) is widespread in western Washington and Ore¬ gon, but has the same low impact in conifer plantations as it has in adjacent coastal British Columbia (Filip 1979, Hadfield and others 1986, Johnson 1976). The disease also affects ornamental plants in urban areas in this region (Schmitz 1920). Armillaria kills planted pon- derosa pine in central Oregon (Adams 1974), but at present it has greater impact in managed natural stands of ponderosa pine and other species in central and eastern Oregon and Washington (Hadfield and others 1986, see chapter 8). Armillaria is of little consequence in planted and natu¬ ral coniferous forests in California (table 9.1), but it does attack many ornamental, orchard, and horticul¬ tural host species in this State (tables 9.2a,b; see below), suggesting that perhaps different species are involved (Adaskaveg and Ogawa 1990). Although Armillaria occurs in conifer plantations in Idaho and New Mexico (Weiss and Rifle 1971, table 9.1), the disease has only been reported from natural forests in Montana, Wyo¬ ming, Utah, and Colorado (Wargo and Shaw 1985). Armillaria is found across parts of the Great Plains, including North and South Dakota, Nebraska, Kansas, Oklahoma, and the eastern edges of Montana, Wyo¬ ming, and Colorado, where it occurs sporadically in over 25 tree species established as windbreaks, Christ¬ mas tree, recreational, roadside, and landscape plant¬ ings (Fuller and James 1986). The species identification is unknown. Armillaria is present in Canada's eastern maritime Provinces and has caused disease in planted conifers in Quebec (table 9.1, Magasi 1990). In Newfoundland, it causes serious disease in plantations of native and in- roduced species of fir, spruce, and pine (Hall and looley 1981; Hall and others 1971; Khalil 1977; Singh 7 8, l c) 80a,b, 1981b,c, 1983; Singh and Bhure 1974; Singh and Richardson 1973). Losses may exceed 30% but are usually lower. The disease also affects urban and shade trees in Newfoundland (Singh and Carew 1983). In Ontario, Armillaria frequently infects both natural forests (Basham 1958, Hord and Quirque 1956, Whitney and MacDonald 1985) and conifer plantations (Huntley and others 1961; Whitney 1983,1988b; Whit¬ ney and others 1978,1989a). Losses have not been seri¬ ous, in some cases because trees were established on abandoned farmland. The species pathogenic to coni¬ fers in this province is A. ostoyae (Anon. 1989, Whitney and others 1989a). Armillaria is also present in Manito¬ ba's forest plantations (C.G. Shaw III, pers. comm.). Across the southern border, the disease attacks conifer plantations in Minnesota (9% mortality; Livingston and others 1982), throughout Wisconsin (Patton and Riker 1959), and in Michigan (under 5%; Bruhn and others 1989). Several planted stands of red pine in Wisconsin experienced mortality ranging from 10% to 37% at age 10 years after the natural oak overstory was killed by aerial application of herbicide (Pronos 1977; Pronos and Patton 1977,1978). Although few reports docu¬ ment Armillaria in planted hosts in the northeastern USA, the disease is common there and causes losses in forests and Christmas tree plantations (table 9.1, Cook 1961, Longenecker and others 1975, Silverborg and Gilbertson 1962). Ornamental and urban shade trees are also attacked, and A. mellea and A. gallica both occur on hardwoods in this region (Dunbar and Stephens 1975, Motta and Korhonen 1986). Armillaria has little impact in eastern white pine plantations in North Caro¬ lina (Leininger and others 1970). In the southern and southeastern USA (Mississippi, Louisiana, Georgia, Florida) Armillaria attacks many different ornamental and shade trees (table 9.2a; Rhoads 1956, Sinclair and others 1987). It and other fungi have reportedly caused up to 25% mortality in plantations of sand pine (Barnard and others 1985, Ross 1970). The pathogen in this region was known as Clito- cybe tabescens as early as 1930 (table 9.2a), and the dis¬ ease is still attributed to A. tabescens in current reports (Sinclair and others 1987) although further examination is needed to establish its identity (Guillaumin and oth¬ ers 1989a). For reviews of Armillaria in North American forests, including plantations, see Boyce (1938), Singh (1980b), Wargo and Shaw (1985), and Sinclair and others (1987). Hepting (1971) supplied information for individual hosts, and a popular account of the disease was recent¬ ly prepared by Williams and others (1989). Armillaria severely affects orchard and horticultural crops in North America, particularly in California, Florida, and the Pacific Northwest (Washington, Ore- 126 Planted Hosts gon, and British Columbia). In California, it is consid¬ ered one of the most serious diseases of stone fruits (Wilson and Ogawa 1979), and it also infects citrus, walnuts, and grapes (table 9.2b). The disease has affect¬ ed these crops since the turn of the century on sites previously forested with native oaks, giving rise to the name "oak root fungus" (Gardner and Raabe 1963, Hewitt 1936). Armillaria infects California's pome fruits (table 9.2b), but it is apparently unimportant in apple and pear orchards (Wilson and Ogawa 1979). The dis¬ ease has also been recorded on California's avocado, blackberry, fig, kiwifruit, loquat, persimmon, and strawberry crops (table 9.2b). Wilson and Ogawa (1979) included olive, chestnut, hazelnut, commom pistachio, and common pomegranate as additional hosts in the State. In Washington and Oregon, hosts include currants, gooseberries, hazelnuts, hops, pome fruits, raspberries, stone fruits, strawberries, and walnuts (table 9.2b, Childs and Zeller 1929, Lawrence 1910, Piper and Fletcher 1903). Armillaria occurs in British Columbia's pome and stone fruits, and occasionally its raspberries and potatoes (table 9.2b). In Florida, Armillaria (cited as "A. tabescens ") has been reported on citrus (Rhoads 1948), and stone fruit trees have been attacked in sever¬ al southeastern States (table 9.2b; Weaver 1974). Other host crops in Florida have included banana, grape (also in Missouri), guava, litchi, pome fruits (also in Louisi¬ ana), and tung (also in Louisiana). Pecan trees have been attacked in Georgia, and stone fruit trees in Wis¬ consin, Illinois, Michigan, Ontario, and Quebec. The disease is recorded on fruit trees in the Eastern States of North Carolina and Maryland (Cooley 1943). Little work has been done to identify which species of Armillaria attack horticultural and fruit crops in North America. In Michigan, Proffer and others (1987) found A. ostoyae, A. mellea , and a species since described as A. calvescens (Berube and Dessureault 1989) attacking sour cherry trees planted on susceptible root stocks (see also Adaskaveg and Ogawa 1990). Central and South America Records of disease on planted hosts are much less fre¬ quent from Central and South America, but they sug¬ gest that Armillaria has a wide, if sporadic, distribution in both the tropic and temperate zones throughout the region (table 9.1, 9.2a,b). In warm-temperate, northern Mexico, A. ostoyae has caused root disease in radiata and Arizona pine planta¬ tions near Chihuahua (Hawksworth 1987, Shaw 1989a). Armillaria occurs in tropical America and has been recorded on conifers in Jamaica and Peru, mainly on pines such as slash pine (table 9.1). According to Ivory (1987), pines have been attacked in Cuba, Honduras, Surinam, and Ecuador. C.A. Garzon B. (pers. comm.) has observed mortality caused by Armillaria among planted eucalypts at high elevation near Popayan in western Colombia. The disease was reported in Colom¬ bia between 1975 and 1982 in slash pine, Mexican weeping pine, Mexican cypress, teak, and eucalypts (C. Alvarado, J.J. Castano, M. Gutierrez, E.R. Ordonez, M.; per C.A. Garzon B.), but it is not considered to be im¬ portant in that country. One account documents Armill¬ aria on hardwoods in Peru (table 9.2a), but it is not recorded in the extensive eucalypt, Gmelina, and pine plantations that have replaced natural rainforest in the Amazon Basin of Brazil, suggesting that the disease currently has little, if any, significance in the hot, low¬ land climate of this region (Brazilian Inst. Forestry Dev. 1976, Rankin 1985). Few records implicate the disease among cultivated crops in tropical America. Armillaria has been reported on cacao in Mexico, Colombia, and possibly Brazil; and it is listed on avocado in Ecuador (table 9.2b; Wood and Lass 1985). Locally, Armillaria has infected Cinchona plantations in Peru (table 9.2b) and economic food crops in the Dominican Republic, Guatemala, and Suri¬ nam (Dist. of Plant Dis. 1980). However, it is not nota¬ ble as a problem in cacao in Ecuador (J. Hedger, pers. comm.), and it was not detected in Trinidad and Vene¬ zuela (Dennis 1950, 1970). Nothing is known of the species responsible for disease in tropical America. In temperate South America, Armillaria has been re¬ ported from Chile and southern Brazil on introduced conifers, especially pines (table 9.1). The disease has killed groups of radiata pine less than 6 years old in Chile, but 1% or less of the forest is affected (H. Peredo L., pers. comm.). Reis (1974) noted that Armillaria had been recognized for a number of years in pine planta¬ tions in Chile (radiata pine) and Brazil (slash, loblolly, and Mexican weeping pine; May 1962a,b, 1964; Brazil¬ ian Inst. Forestry Dev. 1976). Gibson (1973) observed minor damage in slash pine plantations up to 12 years old near Sao Paulo. In its warmer, northerly range in southern Brazil, the disease is confined to higher eleva¬ tions (Brazilian Inst. Forestry Dev. 1976). Mortality varies between 1% and, rarely, 25-30%, and ceases after age 5 years (Brazilian Inst. Forestry Dev. 1976, Ferreira 1989, Hodges 1971). Among cultivated plants, Armillar¬ ia has been reported on cassava and pome fruits in Chile and grapes in southern Brazil (table 9.2b; May 1962a). No published records of the disease so far exist for plantations in Argentina (J.E. Wright, pers. comm.). Many Armillaria species have been described from tem¬ perate South America, including A. limonea and A. no- vae-zelandiae which are pathogenic in Australasia Planted Hosts 127 (Horak 1979; Kile and Watling 1983; Shaw and Cal¬ deron 1977; Singer 1953, 1969; Spegazzini 1922), but the species causing disease in pine plantations and horti¬ cultural crops have not been identified. However, the Northern Hemisphere species are very likely absent or of only minor importance in the south-temperate re¬ gions of the world. Africa Africa has provided many records of Armillaria in vari¬ ous planted hosts from numerous localities (tables 9.1, 9.2a,b; Ivory 1988). These are scattered from the north- temperate zone (Morocco, Algeria, Tunisia, Libya) across the tropics to temperate southern Africa (fig. 9.1). The impact and intensity of disease varies with location and host. A number of reviews, surveys, and literature collations have been published (Browne 1968; Fox 1970; Gibson 1967,1975,1979; Ivory 1987; Moham¬ med and others 1989). According to Mohammed and others (1989; cf Mwangi and others 1989), the two most common species on planted hosts are A. heimii, which was collected mainly above 2,000 m in eastern, central, and southern Africa, and a species culturally close to A. mellen which was found at lower altitudes on both sides of the continent (Pegler 1977). Both occurred on hard¬ wood and conifer plantation species and on horticul¬ tural crops. These distributions are interesting because Armillaria tends to infect plantations established above 1,000 m in eastern Africa where the climate is cooler and wetter. In central and west Africa (Ghana, Nigeria, Zaire, Uganda), the disease occurs in crops at both high and low altitudes (Blaha 1978, Fox 1970). Further sur¬ veys are needed, however, before the various altitudi¬ nal occurrences of disease on each side of the continent can be considered species-related. Plantations of fast-growing hardwood and softwood trees have been established in many African countries in order to replenish natural timber resources that are gradually becoming exhausted (Gibson 1967; Ofosu- Asiedu 1980,1988; Wingfield 1987). Losses from Armill¬ aria have been recorded in species of Araucaria, Finns, Widdringtonia, Acacia, Albizzia, Cassia, Cedrela, Cupres- s us, Eucalyptus, Gmelina, Grevillea, Khaya, Tectona, Termi- nalia, Toona, and Vitis (Gibson 1967, 1975; L.M. Mwangi, pers. comm.; fig. 9.4b). Other reports supple¬ menting those listed in table 9.1 and 9.2a are supplied by Gibson (1964, 1967,1975); Scharif (1964); Browne (1968); Saccas (1975); Bakshi (1976); Ofosu-Asiedu (1980); Nandris and others (1984); Piearce (1976,1984); Nicole and Mallet (1985); and Chipompha (1987). These document additional location records of the disease in planted forest species in Cameroon, Central African Republic, Gabon, Ivory Coast, Morocco, Nigeria, Su¬ dan, Swaziland, and Zambia (fig. 9.1). In West African s, incidence of disease is normally infrequent and occurrence is confined to localized infection cen¬ ters, but Gibson (1967) reported up to 60% loss of Albiz¬ zia falcata in Gabon. In North Africa the disease has been reported from Tunisia (eucalypts, occasional deaths, Gibson 1967) and Morocco (teak, up to 20% losses on some sites from Armillaria and Rigidoporus lignosus (Kl.) Imaz, Gibson 1967). Armillaria has been investigated intensively in forest plantations in Kenya, Malawi, and Zimbabwe (Chipompha 1987; Gibson 1957a,b, 1960, 1961; Gibson and Corbett 1964; Gibson and Goodchild 1960; Masuka 1989; Olembo 1972; Olembo and others 1971; Swift 1968,1970,1972). In these countries, attack occurs on cool, moist, higher sites formerly occupied by natural rain forest rich in hardwood species (Leach 1939) or on old hardwood plantation sites (Gibson 1979). Overall, the impact of the disease is minimal, but it can be local¬ ly severe (over 30% mortality) in younger stands of susceptible species such as teak, slash pine, and mlanji cedar. Normally of little or only local significance, Ar¬ millaria infects plantations of pine, teak, and occasional¬ ly other species in Uganda, Tanzania, Zambia, Swaziland, and South Africa (Gibson 1964,1967,1975; Kotze 1935; Liickhoff 1964; Lundquist 1986,1987; Lun- dquist and Baxter 1985; Piearce 1984; Wingfield 1987; Wingfield and Knox-Davies 1980; Wingfield and others 1989). The horticultural crops most commonly attacked in tropical Africa (table 9.2b) have been cacao (countries bordering the Gulf of Guinea, Central African Repub¬ lic, Zaire, Uganda and Madagascar; Saccas 1975), coffee (widespread; Guinea to Mauritius, Ethiopia to Zimba¬ bwe; Blaha 1978), tea (central and east Africa from Zaire to Mauritius, Kenya to Zimbabwe; Fassi 1959; Leach 1937,1939), and rubber (central Africa from Liberia to Uganda; Fox 1964,1970; Mallet and others 1985; Wastie 1986; D. Nandris, pers. comm.). Brief reviews of the disease have been published for cacao (Thorold 1975, Wood and Lass 1985), and for tea and coffee (Haarer 1963, Saccas 1975, Wallace 1935). In tea and coffee, the disease is known as "collar crack dis¬ ease." Armillaria has been reported less frequently on the fol¬ lowing cultivated hosts in Kenya, Uganda, Tanzania, Malawi, Zimbabwe (table 9.2b; Saccas 1975, Wiehe 1952): banana, cassava. Cinchona, citrus, fig, guava, geranium/pelargonium, granadilla, loquat, macadamia nut, mango, olive, papaya, pome fruit, stone fruit, sug¬ ar cane, and tung. Additional hosts in Zimbabwe are avocado, cotton, grapes, pecan, and strawberry (F.A. Chanakira-Nyahwa, pers. comm.). Records elsewhere in tropical Africa (table 9.2b; Saccas 1975, Turner 19, 0, Wardlaw 1965) include avocado (Ghana), banana (Gha¬ na), cassava (Ivory Coast, Central African Republic, 128 Planted Hosts Zaire), Cinchona (Guinea, Zaire), coconut (Ghana), Cola acuminata (Ghana), cotton (Zaire), Hydnocarpus anthel- mintica (Zaire), lime (Ghana), mango (Ghana), mulber¬ ry (Central African Republic), oil palm (Ghana, Zaire), and pome fruit (Zaire). Wardlaw (1972) reviewed the disease in banana. Further records of Armillaria in culti¬ vated plants are listed from Angola, and possibly Egypt and Sierra Leone (Dist. of Plant Dis. 1980). In temperate Africa, Armillaria has been recorded on fig and citrus in the north (Algeria, Libya, Morocco, Tuni¬ sia), and from banana, citrus, stone, and pome fruit trees in South Africa (Wingfield 1987). A variety of tropical ornamental trees and shrubs are subject to attack from Armillaria in Zimbabwe (F.A. Chanakira-Nyahwa, pers. comm.), and the same is no doubt true of other countries in Africa. Asia and the Pacific Records of Armillaria in planted hosts are widespread, but sporadic, from the Middle East to the Pacific Ocean, even though parts of this region (China, Japan) have a centuries-old reforestation tradition (Izumi 1988, Winters 1974). In north temperate Asia, the dis¬ ease has been reported in economic crops from Iran and apparently Iraq (Dist. of Plant Dis. 1980, Bakshi 1967) and in conifer plantations or hardwood trees from Pakistan, India, China, Korea, and Japan (table 9.1; Bakshi 1967). A record from cypress in Lebanon may refer to a plantation (Scharif 1964), and in north¬ ern Pakistan, infection has been observed on persim¬ mon (Zakaullah and others 1987). Armillaria occurs in the western Himalaya Ranges in northern India (Bakshi 1976,1977) but caused less than 3% mortality in young plantations of deodar cedar and pindrow fir (Singh and Khan 1979). Further east, Armillaria has been found in exotic pine and Japanese redcedar plantations in north¬ ern West Bengal (Bakshi 1976, Singh and Khan 1982). Armillaria is widespread in China (Zhang and Huang 1990). Jie (1982) described extensive attack in Heilong¬ jiang province in the northeast affecting plantations of Korean pine and larch. She reported Armillaria in the interior provinces of Gansu, Sichuan, and Yunnan, on both broadleaf and conifer hosts. Armillaria infects planted Korean pine trees but is not considered serious in Inner Mongolia province (Yang Li, pers. comm.). In Hebei province the disease is attributed to A. tabescens and is recorded in fruit, ornamental, and woodlot trees (apple, pear, peach, almond, white mulberry, locust, poplar, willow, elm, ailanthus, and jujube; Chang and others 1982). Armillaria is a serious problem on citrus in Sichuan province, and on tea and cocoa in Yunnan province in the south (Beijing Forestry University 1983). Armillaria occurs throughout Japan and kills trees in young plantations of Japanese larch (Bakshi 1967; Imazeki 1964; Kawada and others 1962; Ono 1965, 1970) and hinoki (Muramoto 1987,1988; Terashita and others 1983). Mortality rates for hinoki in Kagoshima Prefecture (Kyushu) are mostly under 10% (M. Mu¬ ramoto, pers. comm.). Armillaria has been reported on pine in Japan (Bakshi 1976; Kitijima 1934). It also affect¬ ed a cherry orchard near Osaka (Aoshima and Hayashi 1981). According to Guillaumin and others (1989a), A. mellea is one of the species commonly present in southern Japan where it occurs mainly on non-conifers although three isolates were obtained from hinoki. A Sakhalin spruce collection made from northern Japan was identified as A. ostoyae. This and other species have been reported from northern India (Chandra and Wa- tling 1981, Watling and Gregory 1980), but generally more work is needed to clarify our knowledge of the pathogenic species inhabiting plantations in temperate Asia (see chapters 1 and 2). Armillaria also infects coniferous and hardwood hosts in Korea (Bakshi 1967, Imazeki 1964, Lee and others 1987). The disease occurs in plantations of Korean pine (Sung and others 1989) and also mulberry plantations and orchards (Office of Forestry 1969). In tropical Asia, reports of disease are few, and while this may redect less disease research in some countries, Armillaria causes little or no impact in lowland areas over much of the region. It is rare in tropical India (Ivo¬ ry 1988), but Armillaria has affected green wattle in the south (Nilgiris hills) and Acacia and Albizzia in Sri Lanka (table 9.2a; Gibson 1975). Petch (1910,1928) studied the disease in Sri Lankan tea plantations and found that stumps of interplanted Acacia species fos¬ tered disease spread. Tropical crops such as cacao, coffee, and tea are commonly cultivated beneath a can¬ opy of quick-growing hardwood shade trees such as Acacia , Casuarina, Gliricidia, and Leucaena (Wood and Lass 1985). Generally, however, Armillaria has little impact in Sri Lanka and is not normally seen in horti¬ cultural crops there (A. de S. Liyanage, pers. comm.). Armillaria was reported on tea in southern India several times since 1960 (table 9.2b), but a record on tung has been disputed (table 9.2b). The pathogenic species in tropical India and Sri Lanka has been described as A. fuscipes which may be identical to or closely related to the African A. heimii (Chandra and Watling 1981, Kile and Watling 1988, Pegler 1986). Armillaria has been recorded in Vietnamese (Dist. of Plant Dis. 1980) and Philippine crops (Saccas 1975, Mallet and others 1985, Dist. of Plant Dis. 1980). Re¬ ports concerning conifer hosts in the region are few, but Armillaria has caused low levels of disease in plant- Planted Hosts 129 ings of Khasi pine and Bahaman pine in the Cameron Highlands of Peninsular Malaysia (Barnard and Bever¬ idge 1957; Ivory 1972,1975; M.H. Ivory, pers. comm.). In Indonesia, Armillaria (sometimes as A. fuscipes) was reported several times in the 1920's and 1930's in Cin¬ chona , tea, coffee, and citrus in Sumatra and java. These records were frequently at higher elevations. Armillaria has been found in planted hardwood tree species such as Albizzia, Leucaena, green wattle, and teak in Java and Sulawesi (table 9.2a,b; Gibson 1975, Hadi 1977, Imazeki 1964), but the disease is not common in Indonesia to¬ day (S. Hadi, pers. comm.). Records from Malaysia and Indonesia appear restricted to higher elevations, suggesting that Armillaria natural¬ ly inhabits the cooler, more temperate, montane forest types rather than lowland, tropical rainforests in this region (Fox 1970). In Peninsular Malaysia, Armillaria is absent from lowland plantations of rubber, oil palm, Caribbean pine. Acacia mangium, Gmelina arborea, and Paraserianthes falcataria (R.A. Fox; K.H. Chee; A.M. Tan; Lee S.S.; Norani Ahmad; Maziah Zakaria, pers. comrns.). In Papua New Guinea, too, Armillaria occurs only at mid or higher elevations where it causes root rot and mortality in planted pines and eucalypts (F. Arentz, pers. comm., Arentz and Simpson 1989, Shaw 1984). J.A. Simpson (pers. comm.) observed the disease in eucalypt plantations (swamp mahogany and southern blue gum) established mainly on certain sites cleared of natural southern-beech forest. Mexican weeping pine was also killed by Armillaria at Marafunga. Other euca¬ lypt species (e.g., flooded and New Guinea gums) showed no evidence of disease on most sites, and infec¬ tions in pines (Khasi, Mexican weeping, and Honduran at Lapegu and Kainantu) caused no economic loss. Armillaria has been recorded in Papua New Guinea on several cultivated crops such as coffee (Shaw 1984; table 9.2b), but is of little economic significance on these hosts. Armillaria collections made in Papua New Guinea have been identified as A. mellea, A. fellea, and A. heimii (Shaw 1984), and according to J.A. Simpson (pers. comm.), southern blue gum was attacked by A. novae-zelandiae , and swamp mahogany by A. fellea. Few records document the disease in the tropical Pacif¬ ic (Dingley and others 1981). Armillaria attacks Fiji's high- and low-elevation plantings of mahogany and slash pine on former rain forest sites (Singh 1978, Singh and Bola 1981; pers. obs.). It also occurs in the Solomon Islands (Corner, in McKenzie and Jackson 1986) but has not been reported in plantations. In Hawaii, Armillaria causes disease in young pine plantations over 1000 m l levation, and it also occurs in planted hardwood hosts Laemmlen and Bega 1974, Raabe and Trujillo 1963). The species responsible for the disease in these islands is unknown. Australasia Armillaria is of little consequence on planted hosts in tropical Australia but is widespread in the less arid parts of the temperate and subtropical regions. Even so, reports of disease have been infrequent in planted forests (table 9.1, 9.2a; Kile 1980a). Locally severe out¬ breaks have occurred in radiata pine in South Australia (table 9.1) and Tasmania (Kile 1980a), mountain ash in Victoria (Podger and others 1978), and slash, Hondu¬ ran, and radiata pines in southern Queensland (Bolland and Brown 1981). Mortality declines in pines after about 5 years, and infection centers are small. All plant¬ ed conifers (pines, Douglas-fir, and Queensland kauri) and hardwoods (eucalypts) appear susceptible to some degree (fig. 9.5), but the overall impact is minor. Kile (1980a) suggested that this may be because much of the newly planted land was converted from farming to forestry or from a natural cover of drier eucalypt for¬ ests in which Armillaria has a limited distribution. It is FIGURE 9.5 — Mortality gaps in plantation of alpine ash, caused by A. luteobubalina. Mt. Disappointment, Victoria, Australia. (G.A. Kile) Planted Hosts also possible that Australian Armillaria species have a low pathogenicity toward many introduced tree species. Armillaria has been more frequently reported on orna¬ mentals and garden plants, mainly in New South Wales (table 9.2a), Victoria (Smith and Kile 1981), and Western Australia (Kile and others 1983). The disease is considered serious in Melbourne and Sydney, and losses have been recorded in the botanical gardens of Perth, Adelaide, Canberra, and Sydney (Kile unpubl., Kile and Watling 1988, Smith and Kile 1981). Attacks have also occurred in orchards and horticultural crops (table 9.2b), particularly in citrus, pome, and stone fruit orchards in most states (Doepel 1962, Kable 1974, Ni- cholls 1915). Other listings (table 9.2b) include attacks on bananas (New South Wales), raspberries and logan¬ berries (Tasmania), grapes (Western Australia), hops (Tasmania), mulberry (Western Australia), passionfruit (Western Australia), and vegetables in most states (Do¬ epel 1962, Lea 1909). The species responsible for the disease in urban gardens, fruit orchards, vineyards, horticultural crops, and ornamental trees in Australia is A. luteobubalina (Kile and Watling 1988, Kile and others 1983, Smith and Kile 1981). Armillaria luteobubalina also caused mortality in planted mountain ash in Victoria (Podger and others 1978). In southern Queensland s forest plantations, A. novae-zelandiae and A. fumosa have caused minor disease in radiata pine, and A. pallidula has been associated with slash and Caribbean pines (Kile and Watling 1988). Earlier records ascribed to A. elegans in Australia refer to A. luteobubalina (Kile and Watling 1988). Armillaria occurs throughout New Zealand, and fre¬ quently kills woody hosts in parks and gardens. Re¬ ports on horticultural crops are less numerous (table 9.2b; Dingley 1969, Pennycook 1989), and according to Atkinson (1971), Armillaria is not important as a cause of disease in fruit orchards. Armillaria has infected stone and pome fruit trees in the Auckland district and on the South Island, but the impact has been compara¬ tively minor even though individual growers have occasionally sustained heavy losses. Armillaria is rare in citrus fruit (Atkinson 1971), but recently, the disease has become serious in many orchards of kiwifruit on the North Island (Horner 1985,1987, 1988,1990a,b). Attacks originate from the stumps of felled shelterbelt trees which act as inoculum sources (fig. 9.6). The spe¬ cies causing disease in kiwifruit orchards is A. novae- zelandiae (I.J. Horner, pers. comm.). Both A. novae-zelandiae and A. limonea are responsible for root disease in radiata pine planted throughout New Zealand where indigenous podocarp-hardwood or southern-beech forests have been cleared (fig. 9.7). Losses are spectacular in the first 5 years with up to FIGURE 9.6 — Mortality gaps in kiwifruit orchard caused by A. novae-zelandiae. Te Puke, New Zealand. Gaps follow lines of stumps of willow shelterbelt trees felled 5 years earlier. (I.J. Horner) FIGURE 9.7 — Mortality gaps where young trees have been killed by A. novae-zelandiae and A. limonea in a plantation of radiata pine on a site cleared of indigenous podocarp-hard¬ wood forest, but without stump removal (see fig. 11.1). Tuararangaia Forest, Raungaehe Range, Bay of Plenty, New Zealand. (J. Barran) 30% mortality, but may be more severe later in the rotation in the form of growth reduction and uprooting of final crop trees (MacKenzie 1987). Armillaria also occurs in second-rotation forests; however, its signifi¬ cance in these stands is unknown, but perhaps is great¬ er than previously thought (MacKenzie and Self 1988, van der Pas 1981a). Since the early studies of Birch (1937) and Gilmour (1954,1966b), much has been learned about disease development in forest plantations under New Zealand conditions (Benjamin and Newhook 1984a,b; Hood and Planted Hosts 131 Sandberg 1987,1989; MacKenzie and Shaw 1977; Roth and others 1979; Shaw and Calderon 1977; Shaw and Toes 1977; Shaw and others 1976b, 1980,1981; van der Pas 1981 a,b; van der Pas and Hood 1984). This informa¬ tion was recently reviewed (Hood 1989), and popular accounts of the disease are available (Shaw 1976, van der Pas and others 1983). Disease Development and Impact Although Armillaria occurs in many hosts and places, the same principles govern the behavior of the disease in most plantations throughout the world. This section examines the distinctive features of plantations that influence disease development and describes the effect of infection on crop production. For this discussion, a plantation is defined as a stand or crop created by sow¬ ing seed or by planting. Coppice stands derived from adventitious shoots or suckers and forests regenerated beneath seed trees after logging are excluded. Some factors governing disease development in amenity plantings have been considered by Miller (1940), Rhoads (1956), and Rishbeth (1983). The Significance of Plantations Plantations differ to a greater or lesser extent from natural forest in several respects. They are often even- aged monocultures in which plants are regularly spaced at an appropriate stocking density. Various forms of selection, including clonal propagation, may give rise to planted stock with a reduced genetic base. These features are intended to facilitate crop manage¬ ment and ensure high product yield. However, some aspects of plantations may encourage disease. Inoculum Potential Attack by Armillaria invariably involves inoculum in the soil consisting of woody material colonized by the fungus (see chapter 4). In natural, unmanaged forests, such a food base normally becomes available sporadi¬ cally as trees uproot or are killed by Armillaria or other agents. In plantations, by contrast, colonized stumps or debris left after harvesting a previous stand, generally by clearfelling, are particularly abundant when the crop is established and the young plants are most vul¬ nerable. Induced Host Stress Root systems of naturally established seedlings grow¬ ing under favorable soil conditions are normally well formed whereas those of planted seedlings are often formed or injured. Seedlings weakened in this way i re likely to die from Armillaria infection than are unstressed seedlings (see chapter 7). Singh and Richardson (1973) observed a higher incidence of mortality among bare-rooted stock than among con¬ tainer-grown seedlings after planting. Kessler and Moser (1974) found that plants established from seed survived Armillaria attack during drought stress better than planted trees (see also Buckland 1953, Thies and Russell 1984, Weissen 1981, Whitney and Timmer 1983). Choice of Species Planted hosts are often established outside their natural range, and may therefore be exposed to species and strains of Armillaria which they would not naturally encounter. Under these circumstances, introduced plants in plantations and gardens may conceivably be more prone to attack than hosts indigenous to the re¬ gion although evidence to support this hypothesis is meager. Exotic spruce and firs in a Newfoundland plantation proved more susceptible to Armillaria than indigenous species of the same genera (Singh and Rich¬ ardson 1973). In Californian walnut stands, the intro¬ duced Persian walnut is susceptible to the local species of Armillaria, and is therefore grafted onto rootstocks of the resistant, indigenous northern Californian walnut (Wilson and Ogawa 1979). Either exotic hosts may be inherently susceptible, or suceptibility may be induced by environmental features to which they are not adapted. Monocultures Deaths from Armillaria will be more numerous where greater numbers of susceptible plants occupy an infect¬ ed site. Establishing even-aged, uniformly stocked plantations of susceptible species creates an extreme situation conducive to disease expression that may not arise in floristically and structurally diverse natural forests. Moreover, the uniform, close spacing of many monocultures may facilitate disease spread between susceptible plants. Fedorov and Poleschuk (1981) attributed greater disease impact from Armillaria and H. annosum in the Soviet Union to large-scale planting of single-species forests (c/ Garrett 1956a). The general principles of disease risk in monoculture plantations have been discussed elsewhere (e.g., Gibson and Jones 1977, Peace 1957). Disease Dynamics Disease Establishment Outbreaks of disease typically occur in crops or planta¬ tions that replace natural forests or earlier plantings (fig. 9.7). Inoculum consists of residual infection de¬ rived from the original forest or previous crop. This builds up on stumps and root debris (fig. 9.8; see chap- Planted Hosts FIGURE 9.8 — Stump of recently felled tawa tree with root system colonized by A. limoned on site cleared of indigenous podocarp-hardwood forest prior to burning and planting in radiata pine (see fig. 9.7). Near Rotorua, New Zealand. (I.A. Hood) ter 4), from which it spreads to the new plants. Armill¬ aria is rare in plantations established on non-forested areas such as grasslands, or arable land that has been cultivated for many decades (Gibson 1957a,b; Huntly and others 1961; Kile 1980a; Liese 1939; Rhoads 1925; Singh 1981c), although disease occasionally occurs on these sites (Fedorov and Poleschuk 1981; Gilmour 1954; Rishbeth 1978b, 1988; van der Pas 1981a). Even under these circumstances, some form of woody material such as a thinning stump is needed to establish the primary inoculum (Swift 1972). Less commonly, prima¬ ry inoculum may consist of colonized wood material transported in flood waters (Dadant 1963b, Hewitt 1936, Magnani 1978) or during land contouring prior to planting (Horner 1987). Inoculum may be introduced on wooden stakes, posts, or infected nursery stock (Kable 1974). The fungus can also invade plantations from infected trees or shrubs established for shelter or shade, or as a source of green mulching material, in both tropical (Colonial Research Pesticides Unit 1959; Dadant 1960,1963b; Fassi 1959; Gadd 1940; Gibson and Goodchild 1961; Leach 1936; Milimo 1989; Petch 1922, 1928; Rishbeth 1980) and temperate crops (Beaumont 1954; Chapot 1964; Horner 1987, 1988; Smith 1971). Although the role of basidiospores has been disputed (Fox 1970, Kable 1974, Shaw 1981a, Swift 1972), current evidence indicates that the fungus may enter planta¬ tions in this form (Rishbeth 1964, 1970,1987). Airborne basidiospores appear unable to infect living trees di¬ rectly, with or without wounds (Roll-Hansen and Roll- Hansen 1981, Rykowski 1980), but they can colonize freshly cut wood during the fruiting season (Hood and Sandberg 1987, Molin and Rennerfelt 1959). Basidios¬ pores, which may be less ephemeral than previously assumed (Shaw 1981a), may invade stumps or other debris from which infection then spreads to adjacent, living trees (Fedorov and others 1985, Garrett 1956a, Horner 1988, Petch, in Rishbeth 1955, Rishbeth 1985b). In theory, new disease centers could be created when¬ ever suitable woody substrates become available dur¬ ing a rotation. When a plantation is established, incoming spores may supplement existing inoculum derived from the previous clearfelled forest, especially if this is substantially reduced during burning of the logging debris and slash prior to planting (Hood and Sandberg 1989, Sokolov 1964). Hot fires may kill still¬ living stumps, rendering them more susceptible to subsequent colonization. Thinning stumps (Fedorov and others 1985; Peace 1951,1962; Schonhar 1973) or stumps of shelter trees (Horner 1987) readily harbor the fungus and may act as sources of basidiospore-derived primary inoculum later in the rotation. In Britain, indi¬ rect but convincing evidence for basidiospore infection has been demonstrated by the occurrence of small, single-genotype clusters of Armillaria infection centered on thinning stumps in first rotation stands planted on former arable or heathland (Rishbeth 1978b, 1985b, 1988). In New Zealand, Horner (1988, pers comm.) has shown that infection centers in kiwifruit orchards are initiated by spores that colonize chemically killed stumps in felled willow shelterbelts (fig. 9.6). Armillaria infects the new crop when roots of estab¬ lished plants encounter the primary inoculum, either through direct contact or by rhizomorphs. Infection is governed by host susceptibility, pathogenicity of the species or strain of Armillaria, and the frequency of root or rhizomorph contacts (see chapters 4 and 6). Rhizo¬ morphs grow out from the inoculum source and are found mainly in the top 20 cm or so of soil (Redfern 1973; Rykowski 1981c; Singh 1978,1981b), although in some light soils they may live in colonized stump roots more than 2 m beneath the surface (Horner 1987). They can extend laterally up to 5 m from the inoculum source (Peace 1962), but the distance over which they are able to infect host plants is probably much less (see Planted Hosts 133 chapters 4 and 6). In some situations, they may serve as a bridge between roots nearly, or actually, in contact (Kable 1974). The extent of rhizomorph development depends pri¬ marily on the species of Artnillaria (Guillaumin and others 1984,1989a; Rishbeth 1985b); but the soil also has an effect (see chapters 4 and 6). Rhizomorphs are rare or infrequent in certain plantations in Southeastern United States (Rishbeth 1978a, Sinclair and others 1987), low and mid-elevation parts of Africa (Anon. 1953, Bottomley 1937, Boughey and others 1964, Fassi 1959, Fox 1970, Kotze 1935, Olembo 1972, Swift 1968, Wiehe 1952), northern India (Singh and Khan 1979), eastern China (Chang and others 1982), Papua New Guinea (J.A. Simpson, pers. comm.), Fiji (S. Singh 1978), and Australia (Pearce and Malajczuk 1990a, Podger and others 1978). Disease Distribution Pattern In young plantations, infected plants typically occur in groups centered on the primary inoculum (James and others 1982, Jie 1982, Peace 1962, Podger and others 1978, Swift 1972, van der Pas 1981b, Zondag and Gil- mour 1963). These groups are referred to as infection centers or disease foci. The number of dead trees in a focus is often small (Bolland and Brown 1981, Gibson 1973, Greig and Strouts 1983, Podger and others 1978, Whitney 1983). In these circumstances, the impact of mortality is probably comparatively minor since the limited land area temporarily lost to production is soon reclaimed as surviving tree root systems grow and reoccupy the site (Johnson and others 1972). However, disease centers may be larger and more significant. The shape, size, and distribution of disease foci are governed both by the spatial occurrence of Armillaria in the previous stand or forest and by the distribution pattern of the residual stumps. In western North Amer¬ ica, pathogenic Armillaria exists in large (often over 400 m across), centuries-old colonies (fig 8.6) in natural ponderosa pine forests (Anderson and others 1979, Shaw and Roth 1976). The same may be true in old- growth Douglas-fir stands nearer the coast (Hood and Morrison 1984). In plantations that replace these for¬ ests, many small disease centers arise, each consisting of only a few trees. All trees in every cluster over a wide area (more than 100 m across) are infected by a single Armillaria genotype, apparently derived from the colony of A. ostoyae that originally occupied the site (Adams 1974, Hood and Morrison 1984). By contrast, in selectively logged, old-growth, podocarp-hardwood rainforests in New Zealand, where Armillaria appears \ essentially non-parasitic, single-genotype colonies 11 and densely distributed (Hood and Sandberg 1987). Disease centers in radiata pine plantations subse¬ quently established on these sites are composed of different genotypes, some possibly originating from new introductions of basidiospore-derived material after clearfelling the natural forest (Benjamin and Newhook 1984a). In European forests, colonies of Ar¬ millaria species of one genotype seem to vary between about 10 m across up to 60 m (A. ostoyae), or to around 200 m or more for A. gallica and A. borealis (Durrieu and Chaumeton 1988; cf Rishbeth 1972a, 1982,1985b; Siep- mann 1985; Siepmann and Leibiger 1989; Thompson and Boddy 1983). Disease distribution is also affected by the distribution of stumps in the previous crop (Roth and others 1979, van der Pas 1981b). Many small, dead trees or stumps are more likely to ensure a widespread distribution of primary inoculum than are a few large ones (Pronos and Patton 1977), except when stumps are too small to act as effective inoculum. Secondary Disease Spread The primary inoculum eventually declines as a source of infection although the time required varies with stump size and host species. On some sites, hardwood stumps may act as inoculum for up to 30 years whereas conifers decompose more rapidly (Ivory 1987, Wing¬ field 1987). Whether the disease continues to spread through the plantation beyond the original infection center depends on whether or not infection is transmit¬ ted between healthy plants and adjacent infected plants of the same crop (the secondary inoculum). Secondary disease spread occurs in the same manner as primary spread (by root or rhizomorph contacts) and also by root grafting (Buckland 1953, Greig and Strouts 1983, Hintikka 1974, Peace 1962). It is limited by the distance between roots of neighboring trees and probably by the inoculum potential attained on infected hosts. Infected herbaceous plants, such as those found in vegetable or flower crops, are themselves unlikely to achieve suffi¬ cient inoculum potential for secondary disease spread; it is probable that only the initial, primary inoculum is functional in such plantings (Peace 1962, cf Wilson 1921, 1932). Rishbeth (1972b) suggested that very young, infected pine seedlings may also be too small to act as effective secondary inoculum, although apparent secondary spread has been observed among densely packed seedlings of radiata pine sown in nursery beds (pers. obs). Van der Pas (1981b), working with radiata pine up to 5 years old in New Zealand, monitored mortality rates that followed van der Plank's (1963) model for disease increase without multiplication (slope of log e [1/1-x] linear with time; x = proportion of dead trees) and 134 Planted Hosts concluded no secondary spread happens in very young plantations (cf Whitney 1988b). On the other hand. Swift (1972) fitted mortality rates in up to 8-year-old slash pine in Zimbabwe to the model for disease increase by multiplication (slope of log e [x/l-x] linear with time), implying that tree-to-tree spread of infection had oc¬ curred. Examining these results suggests that data from both authors may be used in either model with an ac¬ ceptable degree of probability; conclusions based on this statistical method should be treated cautiously. Other factors that govern the rate of spread of both secondary and primary disease are discussed in chap¬ ters 5, 8,10 and 11. In addition, spread of disease may depend on the presence of susceptible carrier weed species (S. Singh 1978). Horner (1987) found that infec¬ tion moved along kiwifruit roots faster than rhizo- morphs grew through the soil. At times, these factors effect different rates of spread in different directions and thus distort shapes of disease foci. For example, infection frequently spreads along planting rows (fig. 9.6) giving rise to elongated disease centers (Horner 1985, Marsh 1952, Rishbeth 1978b, S. Singh 1978). Kable (1974) observed a directional trend toward irrigation channels in a peach orchard. Average rates of extension of disease centers are about 1 to 2 m per year (Chipom- pha 1987, Ivory 1987, Kable 1974, Piearce 1984, Podger and others 1978, Rishbeth 1980, Shaw and Roth 1976). Subsequent Disease Development Little is known about how infection centers behave in older forest plantations, but their boundaries might be expected to become more diffuse and irregular, and centers may merge and coalesce (McNamee and others 1989, P. Singh 1981c, Stage and others 1990). If Armillar¬ ia is widely distributed, the disease may affect random¬ ly scattered trees rather than form discrete centers. MacKenzie (1987, cf Bloomberg and Morrison 1989) hypothesized a state of disease equilibrium in an older radiata pine plantation. He observed that although the percentage of basally infected trees remained fairly constant at 50-60% between ages 10 and 19 years, the root collars on 31 % of the trees recovered from infection over this period while those on a slightly higher per¬ centage of trees became newly infected. Chronically infected trees, often without crown symptoms (see chapter 5), have been reported in several forest planta¬ tions (Plavsic 1979, Rykowski 1980, Singh 1981c, Whit¬ ney and others 1989a); planted trees can resist and often recover from infection (Boullard and Gaudray 1975, Courtois 1979, Johnson and others 1972, Kawada and others 1962, Plavsic 1979). Observations in certain natu¬ ral stands suggest that faster-growing trees may be more prone to chronic infection because their larger root systems are more likely to encounter the inoculum (Bloomberg and Morrison 1989, Hrib and others 1983). Very little information describes how disease develops in successive crops planted on the same site except that Armillaria persists in subsequent rotations in both for¬ est plantations and orchards (New Zealand For. Res. Inst. 1954; Delevoy 1946; Gibson 1957a,b, 1960,1967; Holmsgaard and others 1961; Huntly and others 1961; Lundquist and Baxter 1985; Lysaght 1944; Millard 1949; Salmon and Ware 1937; Sisson and others 1978; Swift 1970; van der Pas 1981a). Knowledge is sparse partly because forest plantations which represent second or subsequent rotations are still uncommon and because in non-forest plantations the rotation status of the crop or stand is often unknown. Some authors considered that inoculum may dwindle and disappear after sever¬ al rotations of conifer species (Gibson 1975,1979; Peace 1962; S. Singh 1978; Wingfield 1987), but Redfern (1975) reported examples of disease in second- and third- rotation conifer crops following indigenous hard¬ woods. Inoculum may possibly increase in successive forest plantations, and Garrett (1956a) warned of a potentially greater need for eradication measures in planted stands than in natural forests. Multiple crop¬ ping may even introduce inoculum where it did not formerly exist (Delevoy 1946, Rishbeth 1978b). Armill¬ aria currently inhabits certain second-rotation radiata pine stands in Kaingaroa Forest, New Zealand, on sites not formerly covered in indigenous forest (Gilmour 1954, MacKenzie and Self 1988, van der Pas 1981a). Planting young stock among established trees is anoth¬ er practice likely to enhance inoculum in diseased or¬ chards (Kable 1974). This procedure, like multiple cropping, may also select for particular Armillaria species. Stress and Predisposition Disease development in plantations is influenced by two seemingly contradictory hypotheses of pathogen- host interaction often encountered in the literature. Some workers believe that Armillaria attacks secondari¬ ly or opportunistically (see chapter 7), being serious only on trees predisposed by various physical or biotic agents (Buckland 1953, Gremmen 1976, Huntly and others 1961, Johnson 1976, Sinclair and others 1987). Alternatively, attack may be primary; numerous exam¬ ples in the literature confirm that vigorous plants may be directly infected (see chapter 6). Whichever situation applies in a plantation probably depends on circumstances. Armillaria may be directly pathogenic on susceptible plant species but an opportun¬ ist on weakened, normally resistant hosts. Even so, it is not an easy matter to determine what is really occurring, due to the difficulty of deciding whether or not a host plant is actually under stress (Hiratsuka 1987, P. Singh 1980b). A major consideration is the pathogenicity of the Armillaria species concerned (see chapter 6; Guillaumin Planted Hosts 135 and others 1984,1989a; Rishbeth 1982,1985b) and its inoculum potential (see chapter 4). Peace (1962) suggest¬ ed that a three-way balance exists between the pathogen, the infected host, and the environment ( cf Davidson and Rishbeth 1988, Sinclair and others 1987). Physiological host stress disturbs this balance in favor of the pathogen (Gibson and Goodchild 1961). The effect of stress on the host-pathogen interaction in both natural stands and plantations is discussed in chapter 7. Disease Loss Loss and Crop Age In perennial plantations, the type and extent of disease loss is often closely related to the age of the crop (Rish¬ beth 1972a). In forest plantations, particularly those of conifers, mortality is the most common expression of disease early in the rotation since younger trees tend to be more susceptible and less tolerant of infection (Gibson 1975, Ivory 1987, Peace 1962, Sinclair and others 1987). In some stands, most mortality occurs during the first 8 years or so after planting (Bolland and Brown 1981, Fe¬ dorov and Poleschuk 1981, Fuller and James 1986, Longe- necker and others 1975, Pronos and Patton 1977, Redfern 1978, Shaw and Calderon 1977, Uscuplic 1980, van der Pas 1981a), while in others mortality may continue for at least 25 years (Johnson and others 1972, Morrison 1981, Piearce 1984, Singh and Khan 1979, P. Singh 1981c). Com¬ paratively early peak attack is also recorded in crop plants such as cinchona (Chevaugeon and Merny 1956), rubber (Anon. 1950, Pichel 1956), mulberry (G.-C. 1927), oil palm (Anon. 1948-1950,1958), olive (Leach 1931), and tea (Gadd 1928-1930). However, in some plantation spe¬ cies, such as fruit trees (Hendrickson 1925, Kable 1974) and chestnuts (Bazzigher 1956), killing is less closely related to age. Marsh (1952) found older apple trees to be more susceptible in Great Britain. Armillaria can kill even large specimens of some species; and significant losses may occur in older forest planta¬ tions or in urban plantings, particularly if trees are stres¬ sed (Greig and Strouts 1983; Kawada and others 1962; Manka 1953,1980,1981; Moriondo 1981; Podger and others 1978). However, production losses in older stands are more often caused by growth reduction, butt rot, breakage, and windthrow, as a result of chronic infection (Dariichuk 1986 a,b). Growth reduction due to Armillaria infection is rarely reported in annual crops (Conners 1936) and only occa¬ sionally in perennial horticultural plantations, possibly because reduced fruit yield is a more meaningful pa¬ rameter of production loss (e.g., grape vines: Nieder a), Sisson and others 1978; kiwifruit: Horner 1985). ’ rts of growth reduction are more frequent from Mentations (Peace 1962, Sinclair and others 1987, Williams and others 1989, cf Hrib and others 1983), but even in these crops, values are often presented only for tree height or stem diameter. Wood volume loss is rarely quantified (Morrison and others 1988, Shaw and Toes 1977, P. Singh 1980b, Terashita and others 1983). In older forest plantations, the increment of chronically infected trees can be depressed for extended periods although fluctuations may occur if circumstances change (Shaw and Toes 1977). By contrast, in acutely infected trees, which are usually relatively young, growth may drop sharply for 1-2 years prior to death (Lundquist 1988, Morrison 1981, Podger and others 1978, Szukiel 1980). As trees of certain species become older, infection progresses from the roots to the lower stem heartwood. Butt rot caused by Armillaria is frequently present in older forest plantations, often associated with other decay fungi (Kato 1967b, Schonhar 1969, Storozhenko 1974, Yde-Andersen 1958, Zhukov 1968). Decay is also occasionally reported in other perennial crops such as fruit trees (Adaskaveg and Ogawa 1990, Guillaumin and others 1989b, Petersen 1960). The extent of rot de¬ pends on the host species (Greig and Strouts 1983, Peace 1962) and also on the species of Armillaria. In Britain, conifers are decayed mainly by A. ostoyae, A. borealis, and A. cepistipes whereas hardwoods are de¬ cayed by A. gallica (Gregory 1989, Gregory and Watling 1985, Rishbeth 1982). Decay seldom extends more than a meter or so above ground level, depending on tree size, but it may be slightly more extensive in hardwoods than conifers (Greig and Strouts 1983). Even so, wood destruction represents volume loss from the more valuable butt log section, and the impact of this damage may therefore still be significant (Pegler and Gibson 1972). Losses also occur in butt rotted trees through stem breakage and windthrow (Greig and Strouts 1983, Ivory 1987, Mori¬ ondo 1981, Murray 1959, Sinclair and others 1987, Singh 1981c). The impact of windthrow and growth loss later in the rotation is probably more serious than that of early mortality since residual trees no longer balance the loss by compensatory growth during the remainder of the rotation (MacKenzie 1987). Butt rot also occurs in natural forests, and is discussed further in chapter 8 (which also includes examples from plan¬ tations, reference table 8.2). Evaluation of Disease Impact The overall economic loss caused by Armillaria root disease in plantations is rarely quantified effectively, probably due to the difficulties involved and the effort required. If attack is secondary, it is almost impossible to distinguish loss due to the predisposition stress from that caused by subsequent Armillaria infection (Chabro- Planted Hosts lin 1924, Hiratsuka 1987, Singh 1980b). For a complete and comprehensive economic evaluation, all aspects of disease loss, including uprooting, breakage, and less obvious effects of chronic infection such as growth loss and butt rot, must be considered. Most disease impact assessments in forest plantations contain either qualitative comments such as "of no importance," "severe attack," "scattered mortality," or numerical estimates of the proportion of trees killed. Mortality is the most dramatic expression of the dis¬ ease, and estimates have ranged from less than 3% (Morrison 1981, Singh and Khan 1979) to more than 50% (Ivory 1987, van der Pas 1981a). Mortality loss may be underestimated if counts are not made at regu¬ lar intervals since small dead trees soon become lost among surrounding weed growth. On the other hand, Gibson (1979) suggested that the impact of mortality may be over-emphasized, at the expense of that due to chronic infection, because of its often spectacular ap¬ pearance. Lower levels of mortality can represent a form of natural thinning, and are significantly compen¬ sated by increased growth of remaining trees (Courtois 1979, MacKenzie 1987). Other methods of impact as¬ sessment (see chapter 5) have been used, all of which to some extent underestimate the total economic loss. These methods include percentage of plantations or stands diseased in a forest, area or proportion of forest area out of production as a result of the formation of mortality gaps (Filip 1979, Jie 1982, Podger and others 1978, Redfern 1978, Shaw and Calderon 1977), and height (Singh 1981c) or diameter (Shaw and Toes 1977) increment reduction (see Lundquist 1988). Occasional attempts have been made to assess the total loss throughout a rotation. Gibson (in Ofosu-Asiedu 1980) presented estimates of annual wood volume loss¬ es from conifer plantations in Malawi. Johnson and others (1972) judged that mortality gaps in plantations on Vancouver Island in British Columbia, Canada, were not of sufficient area to support a 48-year-old tree. They concluded the disease would have little impact by age 40-50 years as long as gaps did not expand and assuming that infected trees with healing stem lesions would recover. Shaw and Calderon (1977), and later MacKenzie (1987), evaluated the losses in a radiata pine stand in New Zealand. It was estimated that Armillaria reduced volume production by 6-13% in stands with a projected rotation length of 28 years (MacKenzie 1987). Mortality more appropriately measures loss in or¬ chards and horticultural crops than in forest planta¬ tions. Some reports containing estimates of disease impact in orchards or other non-forest crops are given in Division of Botany, Department of Agriculture (1923) and by Leefmans (1927), Zeller (1932), Pastore (1955), and Horner (1987). In contrast to forest planta¬ tions, it is normally economically feasible to replace diseased trees in orchards. The impact of Armillaria in ornamental hosts varies greatly (Rhoads 1956). Losses are often high but diffi¬ cult to express in economic terms because of the prob¬ lem of assigning monetary values to plants grown for their aesthetic appeal. Costs can be quantified, howev¬ er. These include control measures, removal of dead plants, stumps, and roots, site preparation or resto¬ ration, and the purchase of replacement plants. Although the expenses incurred by individual land- owners are usually relatively low, the aggregate costs of Armillaria root disease in amenity plantings may be substantial. Plantation Management and Disease Control of Armillaria by reducing inoculum and other means is discussed in chapter 11. This section considers how routine tending procedures carried out in forest and horticultural plantations may indirectly influence disease severity, often without reducing the amount of inoculum. In practice, cultural management is rarely conducted specifically for disease control because such operations are costly and because reliable information on the expected economic gains is lacking (cf Pawsey and Rahman 1976a). Planting Young plants must always be considered vulnerable when exposed to a high inoculum potential. Using healthy, vigorous stock (Magnani 1978) and observing good planting practice (Birch 1937, Buckland 1953, Hadfield and others 1986, Johnson 1976, Ono 1970, Thies and Russell 1984) can minimize stress for trees planted on infected sites. The need for care at time of planting is supported by field observations (see also examples under "induced host stress"). Klomp and Hong (1985) found significantly higher Armillaria mor¬ tality among rooted radiata pine cuttings than among planted seedlings. They attributed this to better devel¬ oped root systems on the latter. Hall and others (1971) observed the disease in planted seedlings whereas natural regeneration was unaffected. A number of au¬ thors have recommended using seed or container stock rather than bare-rooted plants in order to reduce plant stress (Hiratsuka 1987, Kessler and Moser 1974, Singh and Richardson 1973, Weissen 1981), but Shaw and Roth (1978) noted that other management consider¬ ations do not always permit this. In theory, dense planting might be expected to favor disease spread due to competition stress and earlier root contact with adjacent plants. However, very little Planted Hosts 137 field or experimental evidence documents the influence of planting density on disease (Hiley 1923). Pielou and Foster (1962) did not find a relationship between densi¬ ty and disease severity in Douglas-fir plantations, and attributed this to the fact that all the stands they exam¬ ined were already old enough for root contact to have occurred. Cultivation and Weed Control Cultivation between rows of plants is a routine proce¬ dure in many orchards or planted crops. This practice controls weeds and improves soil texture, but it may also influence development of Armillaria root disease (Cutuli and Privitera 1986). Injuries sustained by crop plants during cultivation can cause stress and reduce disease resistance (Rosnev and Tsanova 1976). On the other hand, movement of infection across cultivated ground may be interrupted by the severing of roots. In a black currant plantation. Marsh (1952) observed a greater spread of disease along rows separated by par¬ allel strips of cultivated ground than between rows. However, cultivation may also stimulate fresh growth from the cut ends of damaged rhizomorphs (Redfern 1973, Rykowski 1981c, Sewell 1965). The amount of weed growth in a plantation is another factor that may influence disease development. Weeds may stress crop plants in young plantations through competition, especially in areas subject to droughts, rendering them more susceptible to infection, or dam¬ age if already infected. In addition, weeds may serve as bridges to promote disease spread between plants (Shaw and others 1976b, Singh and Bola 1981). How¬ ever, using herbicides to kill weed growth or unwanted shade or shelterbelt trees may also increase disease severity by providing additional inoculum substrate (Andruszewska 1973, Boyd 1986, Pronos and Patton 1977, Schutt and others 1978). Cutting woody weed species may similarly enhance inoculum if root systems die and become colonized by the fungus. Application of certain herbicides may promote or in¬ hibit the growth of Armillaria itself (Andruszewska 1973). Thinning and Pruning Little information describes how thinning impacts disease in forest plantations. Filip (1989a) found that thinning a number of conifer plantations had no signif¬ icant effect on Armillaria mortality 5 years later. Two factors in particular influence disease develop¬ ment when stands are thinned. Thinning may promote resistance to disease by reducing competition among residual trees (Johnson 1976, Singh 1981c, Williams and thers 1989). Davidson and Rishbeth (1988) found that 138 A. mellea, A. ostoyae, and A. gallica all caused extensive infections in oaks and pines weakened by crown sup¬ pression whereas only limited infections by A. mellea and A. ostoyae occurred in unsuppressed, subdominant oak and pine trees, respectively. On the other hand, thinning increases the amount of inoculum in a stand by providing fresh substrates for colonization, as noted earlier (refer under "disease establishment"). Thinning may affect diseased stands in other ways. It may lead indirectly to an unacceptably low stocking density if infected trees continue to die after the final thinning (Morrison 1981). Disease may be encouraged in final crop trees through stress from logging damage during commercial thinning (Johnson 1976). The timing of thinning operations might be expected to influence the level of disease in plantations, but no information is available about the precise effect. Rishbeth (1978b) suggested that thinning late in the rotation may result in fewer, smaller, new disease cen¬ ters, and so result in carryover of less inoculum into the subsequent rotation. Alternately, the bigger stumps created by late thinning may enable the fungus to de¬ velop a greater inoculum potential than on the smaller stumps from earlier thinnings. This could result in larger foci. The stress of pruning live branches is likely to be harm¬ ful to plants already infected by Armillaria. However, the effects of pruning on the disease are even less stud¬ ied than thinning. Chronically infected radiata pine trees were observed to die shortly after being pruned in a New Zealand stand (C.W. Barr, A. Zandvoort, pers. comm.). Excessive pruning of infected trees has also had serious effects in fruit orchards (Stahel 1950). Fertilization In many plantations, especially orchards and horticul¬ tural cultivations, application of fertilizers is an often routine part of management. Such treatment may espe¬ cially benefit chronically infected plants stressed by nutrient deficiencies. However, other effects may also occur. Greater root growth may increase the chance of encounter with inoculum. Development of the inocu¬ lum itself may be promoted or discouraged by particu¬ lar soil amendments (see chapter 4). Fertilizer treatments have generally benefited diseased plants in several field trials, but definite results are not always observed, suggesting that complex interactions are involved. Singh (1983) showed that trees potted in a nutrient-rich soil (pH 4.8) were larger, became infected later, and had a smaller proportion of roots infected than plants in a soil deficient in certain nutrients (pH 3.8). Infection and mortality were lower in the fertile Planted Hosts soil and plants demonstrated active resistance by resin bleeding and callus formation. In a series of field trials in young pine plantations, Rykowski (1976b, 1980, 1981a, 1983) in many cases also demonstrated im¬ proved health to chronically infected trees after fertiliz¬ ers were applied, although mortality rates were largely unaffected. Fertilizing appears to correct partially the tendency for root collar infection to hinder uptake of nitrogen and magnesium (Rykowski 1981b). Spurling and Spurling (1975) found that the damage caused by Armillaria in cultivated banana plants was reduced by applying potassium fertilizers. Clearly, additional trials are required before the effects of fertilization can be exploited in specific cases. Further work is also needed to clarify the effect of lime application on disease devel¬ opment (Anon. 1950, Pawsey and Rahman 1976a, Shields and Hobbs 1979, Sokolov 1971, van der Pas and Hood 1984). Fertilizing with organic material is not always benefi¬ cial. Soil applications of processed urban refuse in¬ creased the incidence of disease in plantations established on infected sites, due apparently to host stress caused by toxic matter in the waste materials (Courtois 1973, Schwarz and Zundel 1975). Even so, Courtois (1979) found that trees which survived on sites treated in this way were larger than untreated plants. This was attributed either to the direct effect of the organic additive or to the "thinning" response among residual, surviving trees. Control of Other Pests or Diseases Trees with chronic Armillaria infection may succumb to other debilitating pest or disease agents present in plantations (see chapter 7). Relieving stress by routine¬ ly controlling these disease organisms or agents may promote resistance to Armillaria. Copper-based fungi¬ cide sprayed to control Dothistroma needle blight [Do- thistroma septospora (Dorong.) Morelet (D. pini Hulbary)] in radiata pine stands in New Zealand re¬ duced the impact of Armillaria in a chronically infected stand (Etheridge 1968, Shaw and Toes 1977). Conclusions Reports in the literature during the past 60 years indi¬ cate that species of Armillaria cause root disease in many planted hosts throughout the world. Attacks occur in softwood and hardwood forest plantations, woodlots, hedgerows, shelterbelts, orchards, and horti¬ cultural crops. The disease is also widespread in shade and amenity trees, ornamental shrubs, and herbaceous plants established in gardens, parks, and on roadsides. Records are particularly numerous from Europe, North America, Africa, and Australasia, but the disease is also present in the Soviet Union, Asia, and South America. In tropical parts of East Africa, Southeast Asia, and South America, it occurs in plantations at higher eleva¬ tions where the climate is comparatively cool and moist. Attack typically occurs in plants established on sites formerly occupied by forests or orchards, and in hosts interplanted among infected trees in existing stands. Infection spreads to new plants when roots encounter rhizomorphs growing from stump roots or when crop roots directly contact roots of colonized stumps. Plants tend to become infected in groups centered on stumps or root inoculum present in the soil, and deaths give rise to unstocked gaps. As the primary inoculum de¬ cays and becomes ineffective, disease centers may ex¬ pand in perennial plantations through the creation of secondary inoculum in the crop itself. The rate of sec¬ ondary spread between adjacent plants is governed mainly by host susceptibility, the degree of interaction between neighboring root systems, and the effective¬ ness of root-to-root transmission of infection. The sub¬ sequent development of disease centers in older plantations is not well understood. Despite the merits of planting, and the obvious necessi¬ ty of growing food crops and forest trees in plantations, several features of these production systems tend to encourage disease development when Armillaria is present. On previously wooded sites, inoculum in¬ creases to a high level early in the rotation when plants are young and especially vulnerable; new plants are predisposed to disease through transplant stress and the malformation of root systems; and spread of dis¬ ease is favored by close spacing of even-aged stock. In monocultures the whole crop may be composed of a species susceptible to infection. Mortality is the most obvious form of disease loss in young perennial plantations, while growth reduction, butt rot, lower stem breakage, and uprooting character¬ ize chronic infection in older planted stands. Although often less spectacular than early mortality, chronic disease may have a more significant economic impact in forest plantations, but production loss has rarely been reliably quantified. The incidence of mortality is commonly quoted but has limited value unless disease centers are large, because increased growth of residual trees tends to compensate for earlier losses in tree num¬ bers. The financial losses in ornamentals are difficult to quantify, and the cost of remedial measures may be underestimated in such plantings. Disease development in infected plantations can be influenced by various management practices, such as the application of fertilizers, but choice of species, planting density, and the timing and intensity of thin¬ ning probably have the greatest effect in forest crops Planted Hosts 139 (see chapter 11). Apart from using less susceptible spe¬ cies on infected sites, operations are rarely conducted specifically to ameliorate the impact of disease due to uncertainty about the effectiveness of such procedures and about their economic benefit. Further research is required to identify management regimes which will maximize returns from infected plantations. TABLE 9.1 —Armillaria in planted conifer hosts, by country 12 World zone Continent Country (and region) References 2 North Temperate America (North) Canada (General) Canada(BC) Canada (Ontario) Canada (Queb. & Maritime Provs.) Canada (Newf.) Asia USA (General) USA (Calif.) USA (Wash., Ore., Idaho) USA (New Mexico) USA (Minn., Wis.Jnd.) USA (Penn., NY, Conn.) USA (Georg., Fla.) India (North) Atlantic Japan Portugal (Azores) Europe (Western) Belgium Denmark Eire Finland France German Fed. Rep. Italy Netherlands Norway Sweden Switzerland United Kingdom 65.2260 41.98; 42. 544; 57. 70; 63 160; 73. 4051 (FA); 82. 7166 (FA); 85. 787; 86. 1121 56. 799; 62. 418; 72. 1946, 73. 2753; 80. 941; 84. 374 (FA) 58 380 (?), 684 (?); 62. 418; 63 223; 67 158b, 68. 1315; 71. 3387o; 86.312 (FA) 63. 223; 70. 257a, 3032; 71 3213; 74. 1670 (FA); 75. 958 (FA), 980 (AE); 79. 3084 (FA); 80. 5607 (FA); 82. 1632 (FA), 1969, 3079, 4411 (FA); 83. 10258 (SF); 84. 914 86. 4102 48, 545 (ornamental?); 54. 79 (ornamental) 74. 6995 (FA), 7692 (FA); 76.420; 77. 4229; 80. 2375; 84. 2840 (FA) 72. 1995 28. 685; 60. 355; 77. 3733 (WA); 79. 3474; 83. 4468 (FA); 84. 5574 (FA) 26. 394 (ornamental); 62 259; 76.3873 (FA) 44. 504 (ornamental); 45. 435 (ornamental); 71. 1461; 73. 2051 56. 405; 80. 3431; 83. 4248 (FA) 62. 551; 63. 499; 66. 635; 71.2001 62. 487; 72. 4380; 77. 5835 29. 147; 36. 473; 46. 482; 49. 51; 50 187; 82. 1971 (FA) 27. 447; 28. 351; 59. 633; 61.497, 635; 62 69 40. 177; 45. 38 40. 54; 47. 222 (ornamental?); 73 1706; 75. 2738 (FA) 23. 431; 27. 586; 33. 798; 67. 2120; 68. 90; 73. 880, 1290; 74. 6724 (FA); 77. 292 (FA); 82. 364 (FA), 1915 (FA), 4359; 86. 2211, 2438, 2441 28. 349, 351; 31. 354, 698; 33. 739 (?); 38. 85; 40. 177; 52. 534; 55. 6, 760; 56. 800; 66. 2266; 68. 2882b; 70. 585, 2681; 73. 6898 (FA); 74. 220 (FA), 3031 (FA), 4396 (FA); 78. 3494 (FA); 82. 1368 (FA); 84. 875; 85. 3996 50. 185, 188; 53. 460 40. 195; 58. 745 51. 202; 66. 2978; 80. 4799 38. 752 (ornamental) 28. 351; 33. 195; 54. 570; 55. 328; 57. 69; 58 560 27. 197, 447; 29. 78; 33. 130; 35. 803; 38. 715; 40. 405, 506; 46. 428; 51. 591; 57 601 (orna mental); 59. 714; 62. 262; 63. 349; 64. 1164, 3023; 66. 888; 67. 3213; 68. 2564; 71. 2000; 72. 4375d; 73. 3431; 75. 2125 (FA); 76. 6820 (FA); 78. 4869 (FA); 79. 3491; 80. 5923; 82. 5974; 85. 2171; 86. 381, 2440; 88. 3140 140 Planted Hosts TABLE 9.1 — (Continued) World zone Continent Country (and region) References 2 South Temperate Tropics Europe (Eastern) Czechoslovakia 27. 213; 28. 351; 45. 257; 76. 2984 German Dem. Rep. (FA); 79. 856 (FA); 83. 1921 (FA); 84. 935; 87. 5207 (FA) 28. 351; 31.699; 32. 141; 33. 480, 739 (?); 37. Poland 647; 75. 4853 (FA) 26. 714; 45. 257; 53. 44, 54. 391; 55. 499; 62. Rumania 552; 64*. 2416, 2433; 69. 936; 73. 3875 (FA); 74. 3001 (FA), 5258 (FA), 7504 (FA); 75. 1 556 (FA), 7809 (FA); 78. 2234 (AE); 81. 730 (FA), 3032 (FA), 4126 (FA), 82. 381 (FA), 419, 2273 (FA), 3631 (FA); 84. 919; 85. 3770 (FA), 4827 (FA), 6070 (FA), 86. 5200; 87. 1573 (FA), 1893 (FA) 72.3608 Yugoslavia 76. 4309 (FA), 4312; 81. 6678; 82. 1632 (FA) USSR Soviet Union 28. 415; 65. 1283; 66. 1528; 75. 6374 (western, incl.Belorussia, Ukraine, (FA); 81.4516 (FA), 6092; 86.3004; 87. 2082, Caucasus) 2411 (FA), 4716 (FA) Soviet Union (Urals, Krasnoyarsk) 62. 417; 65. 1281 (?), 1978 (?) Africa South Africa 33. 142; 34. 425; 37. 355, 784, 54 America (South) Brazil (southern) 657; 56. 405; 81.2785; 83. 2174; 88. 1500 64. 2095; 67. 2848 Chile 63. 350; 65. 552; 67. 3227 Australasia Australia 23. 298; 71. 3384f; 82. 1632 (FA) New Zealand 34. 533; 38. 714; 45. 297; 53. 52; 54. 328, 329; Africa General 55. 267; 56. 565; 63. 156, 635; 69. 618c; 74. 2453 (FA); 82. 2277 (FA); 85. 2889 (FA) 82. 1634 (FA) Kenya 51. 141, 309; 54. 16; 58. 116, 190; 60. 509=61 Malawi 569; 61.436; 65. 261 5a 49. 200; 53. 669 Mauritius 46. 52 Tanzania 52. 225, 55. 350; 65. 1924, 2615a Zimbabwe 62. 762; 63. 727; 68. 3020; 72. 4431 America (Central) Jamaica 68.2958a /Caribbean America (South) Peru 78. 3635 Oceania Fiji 65. 1351b USA (Hawaii) 65. 849 (FA); 74. 6994 (FA) 'Published reports of Armillaria attack in coniferous plantation forests (with occasional records for ornamental plantings) All listings (except two) refer to Northern Hemisphere host genera, including tropical pines: Abies, Cedrus, Chamaecyparis, Cryptomeria, Cupressus, luniperus, Larix, Metasequoia, Picea, Pinus, Pseudotsuga, Thuja, Tsuga (exceptions:51.141 for Kenya and 53. 669 for Malawi, concerning Araucaria, Callitris, Widdringtonia). References to trees attacked within their natural distribution ranges (mainly North Temperate) are some¬ times doubtfully included; it is not always clear whether these are planted or naturally seeded. Sources: Review of Plant Pathology (Review of Applied Mycology), unless otherwise stated Code: year (not volume No.). No of abstract or [prior to 1964] page (abstract journal title abbreviation, as applicable). AE, Review of Applied Entomology, Ser. A; FA, Forestry Abstracts, hA, Helminthological Abstracts, Ser. B, HA, Horticultural Abstracts; PB, Plant Breeding Abstracts; SF, Soils and Fertilizers; WA, Weed Abstracts. Compilation: 1922-1972, manual search (keywords: ARMILLARIA, CLITOCYBE TABESCENS) 1972-June 1988, computer search (descriptor ARMILLARIA ( ) MELLEA; Commonwealth Agricultural Bureaux Dialog Information Retrieval Service, duplicate reports not listed) Planted Hosts 141 Table 9.2a —Armillaria in planted, non-conifer (angiosperm) hosts, by country.(a) Species used in commercial forestry, for shelter, or as ornamentals 12 Host group World zone Continent Country (and region) References 2 Northern deciduous North America Canada (Ont.) 64. 1755 (?) broadleaf trees Temperate (North) Canada (Queb., Newf.) 62. 418(?); 74 1669 (e g., beech, Fagus (?;FA); 83. 4775 birch, Betula; USA (Calif.) 59. 178; 71. 3187 chestnut, Castanea; USA (Central & East) 32. 411; 41. 183; 42. elm, Ulmus; oak, 272; 76. 7439 (AE); Quercus; poplar, 87. 78 Populus; willow. USA (Mississ., Fla.) 44. 417; 51. 1332 Salix) Asia Chinese Peoples' Rep. (?;FA);70. 864 47.421 Japan 60. 354 Pakistan 88. 5414 (FA) Europe (Western) Belgium 73. 6416 (FA) France 23. 431; 24. 5, 8; 27. 586; 28 290; 45 436; 47. 320; 66. 626; 82. 364 (FA); 84 3385 (FA), 6619 (FA); 85. 5785 (FA); 86. 2211, 2438,2441 German Fed Rep. 70. 1156; 72. 3694k Italy 54. 568; 61.493; 64. 3033; 79. 1785 (FA); 81.4684 Netherlands 31. 696; 40. 195 Spain 64.846 Switzerland 57. 69 (?), 673; 61.492 United Kingdom 24. 244; 27. 198; 28 126; 46 428; 48 399, 70. 1157; 72. 2871b; 74. 7708 (FA); 79. 899; 82. 1632 (FA), 5974; 83. 4450; 85. 2171; 86. 381, 2440 Europe Bulgaria 81.3154 (FA) (Eastern) Hungary 86. 781 Poland 74. 1768 (FA); 81.3032 (FA); 87. 2326 (FA) Yugoslavia 26. 705 (?); 27. 5 (?); 30. 278 USSR Soviet Union 59. 426 (?), 66. 1528 (western, Novosibirsk) (?); 69. 2009 South Africa South Africa 27. 237 Temperate Australasia Australia 55. 285 Other hardwood North Africa Tunisia 71. 285, 1396 trees Temperate America USA (Calif.) 28. 494; 71.3187 (?); (e.g., Acacia; (North) 80. 500 (FA) Casuarina, USA (Fla.) 30. 159, 41. 563; 42. Eucalyptus; 486, 497; 44. 417, 504; Grevillea; 48. 279; 51. 1332 (FA); Leucaena; 53. 406; 57. 560 teak, Tectona; Europe German Fed. Rep. 70. 1156 (?) Terminalia) (Western) Europe (Eastern) Hungary 86.781 South Africa South Africa 33. 142; 37. 784; 88. Temperate Australasia Australia 1500 59. 564 (?);77. 3 (hA; ?);82. 1632 (FA) Tropics Africa Ghana 27. 19 Kenya 51. 141; 60. 214, 509; 61. 569; 82. 7157 (FA) Madagascar 55. 107, 451 Malawi 29. 202, 40. 311; 53. 669 Mauritius 30. 507; 52. 537; 60. 633 142 Planted Hosts Table 9.2a — (Continued) Host group World zone Continent Country (and region) References 2 Shrubs and ornamental herbs Tanzania Uganda Zaire Zimbabwe America Peru (South) Asia India (south) Indonesia (Sumatra, Java, Sulawesi) Sri Lanka North America USA (General) Temperate (North) USA (Wash., Ore.) USA (Calif.) USA (Fla.) Europe (Western) France German Fed. Rep. Italy Netherlands Switzerland United Kingdom (incl. Jersey) Europe (Eastern) Czechoslovakia USSR Soviet Union (Georgia) South Africa South Africa Temperate Australasia Australia Tropics Africa Malawi Tanzania Zaire Zimbabwe Asia India (south) 33. 201; 36. 746 (?); 52. 225; 67. 1348 24. 509; 27. 15 51. 512 62. 126, 761; 63. 727 78. 3635 60. 123; 64. 558 25. 79; 31. 525 28. 745; 29. 470; 31. 275 33.696 32. 786; 48. 134; 70.1677 54. 79; 59. 147; 68. 2734; 69. 874; 72. 1551; 80. 1616 42. 497; 44. 504; 48. 23; 51. 1332 (FA); 57. 560 72. 4086; 82 4011; 86. 221 1, 2438 70. 1156 86.4983 40. 195 57. 69 29 628; 32. 376; 35 366; 36. 478; 38. 823; 48. 462, 53. 332; 54. 484, 605; 61. 267; 64. 2792; 85. 221 1; 86. 381 70. 2560 67.643 27.237 48. 275; 59. 564; 67. 660 34. 216; 35. 834; 36. 780 48. 158 51. 196 62. 126; 65. 598 64. 558 ’Published reports of Armillaria attack in forest plantations, woodlots, gardens, parks, roadsides, hedgerows, and farm shelterbelts. Includes ornamental nursery plants, trees established for shade or edible fruit supply (home or local, non-commercial use), and plants used to shelter production crops or provide green manure. References to trees attacked within their natural distribution ranges (mainly North Temperate) are sometimes doubtfully included, it is not always clear whether these are planted or naturally seeded. 2 As for Table 9.1. Planted Hosts 143 TABLE 9.2b — Armillaria in planted non-coniferous (angiosperm) hosts, by country, (b) Species used in eco¬ nomic production (except forestry) 12 Crop World zone Continent Country (and region) References 2 Avocado North America USA (Calif.) 35. 707; 49. 630 (?); 56. (Persea) Temperate (North) 907; 66. 1868 Tropics America Ecuador 60. 435 (South) Banana North America USA (Fla.) 32. 382; 42. 497 Temperate (North) South Africa South Africa 72. 1693 Temperate Australasia Australia 34. 356; 67. 1674(7) Tropics Africa Kenya 54. 141; 65. 2697a Malawi 76. 5091 (HA) Tanzania 33. 552; 53.324 Zimbabwe 62. 126 Berryfruit North America Canada(BC) 38. 49 - cane ( Ribes ; Temperate (North) USA (Wash., Ore.) 23. 278; 44. 112; 45. currant, gooseberry) 423 - bramble ( Rubus ; USA (Calif.) 52. 24 blackberry, logan- Europe United Kingdom 24. 525; 53. 320 berry, raspberry) (Western) USSR Soviet Union (Krasnodar) 72.4191 South Australasia Australia 49. 528; 60. 327 Temperate New Zealand 42. 29 Cacao (cocoa) Tropics Africa Cameroon 57. 383 Ghana & Togo 24. 325; 25. 463; 27 19, 659, 704; 28. 93, 565 Ivory Coast 36. 16 Madagascar 55. 107 Nigeria 69. 755c Sao Tome & Principe 26. 149; 80. 5680 Uganda 24. 509; 26. 17; 74. 1931 (HA) Zaire 47. 235; 49. 31,272 America Mexico 49. 328 (Central) America Brazil 39. 93 (?) (South) Colombia 60. 15 Australasia Papua New Guinea 52. 9 Cactus North Europe Italy (Sicily) 83.3151 (Opuntia ficus- Temperate (Western) indica / edible fruit) Cassava (Manihot) Tropics Africa Tanzania 33. 552 Zaire 57. 308 America Brazil 36. 278 (?) (South) Chesnut Refer Table 9.2 (a) (Castanea) Cinchona Tropics Africa Guinea 59. 94 (quinine) Zaire 44. 431; 46. 45, 154; 51. 196 America Peru 55. 675 (South) Asia Indonesia (Java, 23. 9; 24. 189; 25. 79; Sumatra) 28. 308; 30. 161; 31. 298; 37. 160; 39. 578 Citrus fruit North Africa Libya 60. 658 (grapefruit, lemon, Temperate Morocco 64. 3222 lime, orange, Tunisia 33.302 tangerine etc.) America USA (Calif.) 26. 358; 30. 766; 32. (North) 40; 41. 160, 360; 45. 225= 366; 49. 630; 51. 144 Planted Hosts fABLE 9.2b —{Continued) Europe (Western) Europe (Eastern) South Australasia Temperate USA (Fla.) Cyprus France (inch Corsica) Greece (Crete) Italy (incl. Sicily) Malta United Kingdom Yugoslavia Australia Tropics Africa Kenya Asia Malawi Indonesia (Java) Coffee Tropics Africa General Cameroon Central African Rep. Ethiopia Guinea Ivory Coast Kenya Madagascar Malawi Mauritius Sao Tome & Principe Tanzania Uganda Zimbabwe Asia Indonesia (Java) Australasia Papua New Guinea Co/a acuminata Tropics Africa Ghana (edible nut) Cork (Quercus suber) North Europe France Temperate (Western) Italy (Sardinia) Portugal Cotton Tropics Africa Zaire Fig (Ficus carica) North Africa Algeria, Morocco, Temperate Tunisia America USA (Calif.) (North) Europe France (Western) Flower production North Europe France Temperate (Western) Geranium/Pelar¬ Tropics Africa Kenya gonium (oil source; Tanzania see also Table 9.2a, Zaire) General Tropics Africa Tanzania Grapevine (Vitis) North America USA (Fla., Missouri) Temperate (North) USA (Calif.) 608, 54. 79; 55. 366, 641; 64. 520; 68. 1886; 69. 469; 70. 3661; 71. 1224d, 74. 2805 (HA); 77. 1943; 80. 5143, 5144 31.99; 32. 365; 42. 486, 497; 48 279; 72. 1495 32.696 56. 447; 86 719 39. 672 36. 213; 85. 4323, 88. 196 34. 81; 35. 618; 36.780 81.6692 (HA) 55. 297 23. 354; 27. 101; 33. 142; 36. 280, 37. 451; 42. 440; 44. 296; 46. 558; 47. 74, 337; 49. 453; 53. 175, 62. 151 85. 2544 33. 10 38. 162; 39. 794; 40. 143 37.454 55. 86 54. 537 67. 329; 69. 5; 70. 918 59. 94; 63. 612 50. 99 26. 299 (?); 30. 31 (?); 60. 228; 73. 2278 32. 699 (?); 34. 506 (?); 55. 107, 451; 58 408; 63. 683; 64. 2914 28 239; 62. 7 72. 8a 59. 745 (?); 80. 4618 33. 201; 34 114; 36. 261; 51. 509 23. 409; 24. 509; 28. 701; 33. 422 62.761 39.452 56. 423 27. 19 23.431 64.845 72. 4380 (?) 49. 31 24. 88 26. 37 (?); 48. 372 47. 328; 82. 4011; 86. 2438, 5626 Refer Table 9.2 (a) 56. 358 51.79 36. 746 25. 585; 27. 460 51.402; 64. 520; 73. 7573 (HA); 80. 1097 (SF), 9620 (PB) Planted Hosts 145 TABLE 9.2b — ( Continued) Crop World zone Continent Country (and region) References 2 Europe Austria 81. 1114 (HA) (Western) Belgium 57. 86; 64. 1 512v France 25. 525 (?); 27 563, 586; 30. 360 (?); 38. 653; 59. 118 (?); 60. 767; 61 507; 82. 4011, 4359, 86 2211,2438, 2441 Greece 55. 706 Italy 36. 774; 46. 436; 56. 658 Spain 34. 745; 36. 75, 541; 53. 419 Switzerland 56 415, 76. 3131 (HA) Europe (Eastern) Bulgaria 25. 460 USSR Soviet Union (western, Georgia) 64. 1535m; 77. 3152 South America Brazil (southern) 83. 3159 Temperate (South) Australasia Australia 27. 101 Guava (Psidium) North America USA (Fla.) 30. 159; 42. 497 Temperate (North) Tropics Africa Malawi 66. 700d Hazelnut (Corylus) North America USA (Oreg.) 48. 165 Temperate (North) Europe Italy 85. 4520 (Western) United Kingdom 26. 278 Hops (Humulus) North America USA (Oreg.) 49. 593 Temperate (North) Europe Greece 60. 210 (Western) United Kingdom 36. 478, 605; 37 822; 40. 364; 43. 39; 44 2 South Temperate Australasia Australia 60. 327; 63. 137 Hydnocarpus anthelmintica Tropics Africa Zaire 49.271 (medicinal oil) Kiwifruit North America USA (Calif.) 72.2721 (Actinidia) Temperate (North) Europe (Western) France 86. 2438 (?) South Temperate Australasia New Zealand 71.3066 Lavender North Europe France 34. 99 (Lavendula: Temperate (Western) United Kingdom 39. 724 perfume) Litchi (Litchi; North America USA (Fla.) 42. 497 (?); 56. 781; edible fruit) Temperate (North) 62. 618 (?) Loquat North America USA (Calif.) 27. 175; 49. 630 (?) (Eriobotrya) Temperate (North) Tropics Africa Tanzania 38. 15 Macadamia Nut Tropics Africa Zimbabwe 68.2588 Mangel ( Beta ; North America Canada (New Bruns.) 36. 632 cattle food) Temperate (North) Mango (Mangifera) Tropics Africa Ghana, Uganda, Zimbabwe Refer Table 9.2 (a) Mulberry North Europe France 25. 201; 27. 567 (Morns) Temperate (Western) Italy 29. 207; 30. 213 Europe (Eastern) Hungary 86.781 USSR Soviet Union (Uzbek., 46. 17 146 Planted Hosts TABLE 9.2b — ( Continued) Crop World zone Continent Country (and region) References 2 Kirghiz., Tadzhik. ?) South Australasia Temperate Australia Oil palm (Elaeis guineensis) Tropics Africa Zaire Olive North Temperate Europe (Western) France Italy Spain Tropics Africa Malawi Papaya (papaw) Tropics Africa Kenya Tanzania Passiflora spp. (passionfruit, South Temperate Australasia Australia granadilla) Tropics Africa Zimbabwe Pecan (Carya North America USA (Georgia) illinoensis) Temperate (North) USA (Calif.) Persimmon North America (Diospyros) Temperate (North) Canada(BC) Pome fruit (Malus, North America Pyrus; apples, pears) Temperate (North) Europe (Western) USA (Wash., Ore.) USA (Calif.) USA (Louis., Fla.) France German Fed. Rep. Italy Malta Netherlands Spain Switzerland United Kingdom South Africa South Africa Temperate America (South) Australasia Chile Australia Tropics Africa Kenya Tanzania Zaire Zimbabwe Rubber (Hevea) Tropics Africa Cameroon Central African R< Chad, Congo 8J or Gabon Congo Nigeria Uganda Zaire 27. 101 47. 235; 49. 31,272; 50. 236, 292; 51 155, 512; 57. 513 24. 348; 82. 4011,86. 1420, 2438 46. 492, 51 235; 56. 658; 62. 358; 76 1641 (HA) 46. 508 31.707 48. 483; 71. 5b 48. 483; 50. 145 27. 101 65. 598 71. 294 48. 372 (?); 49. 630 (?) 23. 304; 26. 146 24. 89; 26. 746; 30. 212 23. 394; 32. 40; 25. 21; 26. 37; 34. 552; 40. 24; 48. 372; 64. 520 (?), 2855; 72. 1631 41. 169; 42 497 23. 431 (?); 59. 125; 86. 2438 35. 677 (?); 71. 1867 (?) 74. 2033 (HA) 36. 780; 38. 589 51.420 34. 745 (?); 36. 75 (?), 541 (?); 42. 338 50. 443 24. 259; 26. 278; 37. 822, 53. 320; 54. 91; 60. 178 (?); 70. 1426 37. 784 (?) 69. 2282 23. 353; 25. 399 (?); 26. 105; 27. 101; 28. 304 (?); 35. 451 (?); 45. 318 (?); 56. 613 (?); 60. 327; 62. 158; 67. 2775 51. 309 48.224 51.196 36. 705; 38. 46; 66. 2a 66. 1166 53. 507 64.3003 64. 3003, 3172; 66. 1166 24. 509 47. 466; 48 123, 382; 49. 271; 50. 236, 292; 51. 512; 54. 413; 55. 583; 56. 925; 57 308; 61. 486 Planted Hosts 147 TABLE 9.2b — ( Continued) Crop World zone Continent Country (and region) References 2 Stone fruit North America Canada (BC, Ont., 26 146; 47 429 (?); 75. (Prunus ; almond, Temperate (North) Queb.) 2151 (HA) apricot, cherry, USA (Wash., Ore.) 26. 746 peach, plum, etc.) USA (Calif.) 25. 681; 26. 37; 31. 323; 34 552; 36. 518, 40. 24; 41. 161; 45. 453; 46. 446; 48 372; 49 630, 52. 68; 53. 24; 54. 36; 64. 520 (?), 2855; 72. 4174; 77. 1943 USA (Wise., III., Mich.) 23. 481; 54. 733; 88. 4191 (FA) USA (Mary., Nth 30. 159; 41.68, 562; Carol., Sth Carol., 42. 497; 44. 25; 53. Georg., Fla.) 682; 54. 434; 55. 603; 61 370; 62. 398; 63. 618; 69. 1842 Europe France 23. 431 (?); 26. 304; 27. (Western) German Fed. Rep. 563 (?); 30. 116, 465; 32 791; 49. 527; 50. 265; 54. 161; 56. 109, 305, 82. 4011, 4359; 86 2211, 2438, 2441; 87. 9220 (HA) 35. 677 (?); 71. 1867 (?); 73. 63 (HA) Italy 34. 706; 56. 658; 63. 562; 68. 2215 Netherlands 48. 244 Spain 34. 745 (?); 36. 75 (?), 541 (?) United Kingdom 28. 647; 37. 822; 60. 178 (?) Europe (Eastern) Hungary 47. 344; 48. 140; 86. 781 Yugoslavia 86. 808 South Africa South Africa 27. 237; 37. 784 (?) Temperate Australasia Australia 25. 399 (?); 28. 304 (?); 35. 451 (?); 45 318 (?); 47. 285; 48. 27; 56. 613 (?); 65. 1354g; 67. 2775; 73. 2099f Tropics Africa Kenya 37. 796 (?); 51. 309; 58. 136 Zimbabwe 36. 705; 38. 160 Strawberry North America USA (Wash., Ore.) 29. 727; 31. 163; 32. (Fragaria) Temperate (North) USA (Calif.) 727; 39. 402; 45. 423 61. 617 Europe (Western) United Kingdom 27. 650 Sugar cane (Saccharum) Tropics Africa Tanzania 33. 552 Tea North Asia India (northeast) 40. 369; 83. 3975; 85. Temperate 2155 Tropics Africa Kenya 53. 513 (?); 58. 512; 60. 214; 61.73, 723; 76. 10851 (HA) Malawi 28. 275; 29. 202; 33. 10; 34. 216; 35. 14, 36 780; 37. 209, 564; 40. 311; 49. 200; 74. 8119 (HA); 81.3935 Mauritius 52. 538 Mozambique 50. 89 Tanzania 36. 261; 53. 513 (?); 55. 350 Uganda 24. 509; 29. 756; 37. 838 Zaire 49. 272; 57. 309 (?); 60. 440 Zimbabwe 59. 295; 65. 598 Asia India (south) 55. 489; 60. 123; 64. 558, 66. 595b; 76 8835 (HA) Indonesia (Java, 23. 9; 24. 5, 64, 611; Sumatra) 26. 585; 31.409; 38 162, 202; 39. 579 Malaysia 37. 657 (?) Sri Lanka 23. 295; 28. 745; 29. 469; 31. 275; 40. 678 (?) US PI mi ted Hosts TABLE 9.2b — (Continued) Q 0 p World zone Continent Country (and region) References 2 Tung (Aleurites; oil) North Australasia America Papua New Guinea USA (Louis., Fla.) Temperate Tropics (North) Africa Malawi Vegetables North Asia America India Canada(BC) (carrot, parsnip, Temperate (North) Belgium potato; also Europe tomato; see (Western) United Kingdom elsewhere for USSR Soviet Union cassava, mangel) South Australasia (Leningrad) Australia Walnut (Jug la ns) Temperate North America USA (Ore.) Temperate (North) Europe (Western) USA (Calif.) France South Temperate Italy Europe Bulgaria (Eastern) Czechoslovakia Hungary Australasia Australia 56. 423 37. 426 (?); 41. 169, 44. 504; 49. 365 40. 626; 49. 200; 51. 295; 53. 703; 62. 734 50. 588 (?; see 55. 267) 37.832 39. 724; 71.460b 22. 357; 48. 462 46. 316 27. 101; 33. 142; 34. 257; 37. 118 24. 89; 42. 310; 49. 94; 51. 594; 52. 463; 59. 39 26. 37; 34. 552; 45. 453; 48. 103,372 22. 35, 77; 24. 179; 25. 201, 577; 26. 526; 27. 200, 426, 563, 586; 28 686; 32. 95; 34. 366 (?); 36. 763; 72. 4403 (?); 84.4723 (FA) 46. 427 76. 6817 (FA); 77. 871 (FA), 4716 27. 6 48. 140; 86. 781 54. 610 ’Published reports of Armillaria attack to planted trees, shrubs, or herbaceous species used as commercial food crops or for processed products (except timber or pulpwood). 2 As for Table 9.1 Planted Hosts 149 CHAPTER 10 Modeling the Dynamics, Behavior, and Impact of Airnrillaria Root Disease Charles G. Shaw III, Albert R. Stage, and Peter McNamee nformation on the ecological, biological, and pathological attributes of Armillaria spp. and the root disease they cause comprises the major por¬ tion of this book. Integration of this material, particularly as it relates to the portrayal of disease dy¬ namics and the quantification of disease impacts, would markedly enhance its utility for foresters, or¬ chard managers, and scientists. Models can accomplish this objective, and some have been developed for root disease caused by Heterobasidion annosum (Fr.) Bref. (Alexander and others 1985, Pratt and others 1989) and Phellinus weird (Murr.) Gilbn. (Bloomberg 1988). This chapter describes how information on Armillaria root disease has been used to develop a predictive model of disease dynamics, behavior, and impact. For¬ esters currently make decisions about root disease management using their mental model of the disease process in the affected area as a guide to select treat¬ ment alternatives for the land. The process of building a predictive model combines existing data and the key features of the mental models of several knowledge¬ able forest managers and scientists in a set of mathe¬ matical equations. By pooling and structuring the knowledge of many, we should have a better model for individual stand management as well as for overall forest planning than would be assembled by any single manager or scientist. An important benefit of the mod¬ el-building process is a highlighting of our still inade¬ quate understanding of many biological aspects of Armillaria root disease, increased knowledge of which is necessary to improve management. Pathologists have a wealth of information about Armill- aria spp. and the root disease they cause. However, even when these data are published they frequently are not available to managers in a form that directly assists decision making. Because pathologists have the best biological understanding of root disease dynamics as well as the limitations of available data, it is imperative that they define the biological assumptions necessary to develop a predictive model. If research pathologists diligently perform this role, then the resulting model not only becomes a tool of immediate use to managers, but also becomes a quantitative description of a series of hypotheses about root disease dynamics, behavior, and impact. As such, it can aid scientists in identifying serious data gaps and thus help to define and prioritize research needs. Scientists trained in the research process may find devel¬ opment of a management-oriented model troublesome because professional judgement, rather than statistically analyzed data, often becomes the only, or at least prima¬ ry, basis for a generalized assumption that can markedly influence the outcome of a model prediction. The tradi¬ tional researcher is far more comfortable with the model of a scientific paradigm where behavior is judged at lev¬ els that are often far removed from the decision criteria that are required of a predictive model for management. Consequently, knowledge gaps can be left to future re¬ search as no immediate opportunity exists for application of the model. Forest managers who routinely encounter stands severely impacted by root diseases are desperate for tools to deal with these complex and damaging prob¬ lems. Thus scientists, who best understand these prob¬ lems, even if that understanding comes primarily from their professional judgements and experiences, can no longer take a laissez faire, hands-off approach to manage¬ ment-oriented modeling. This chapter describes the integration of our current understanding of Armillaria root disease dynamics and the damage the disease causes in various conifer eco¬ systems in western North America (see chapter 8) into a predictive model for management use in silviculture and forest planning (Stage and others 1990). The hy¬ potheses or assumptions that underlie the quantitative relationships contained in the model are discussed and referenced to information presented elsewhere in this book. In addition, direction is provided to indicate how model users (both managers and scientists) can exam¬ ine alternative hypotheses about the dynamics and behavior of Armillaria root disease. Modeling Root Disease The process used to build the Western Root Disease Model (Brookes 1985, Eav and Shaw 1987, Shaw and others 1985) can serve as a prototype for modeling the dynamics and behavior of Armillaria root disease in other forest ecosystems or orchards. History and Structure of the Western Root Disease Model Recognizing the serious economic impact of annually losing 6.8 million cubic meters of timber in the Western United States to root diseases (Smith 1984), the USDA Forest Service initiated a project to develop a root dis¬ ease model (Brookes 1985). The protocols of Adaptive Environmental Assessment, as outlined by Holling (1978), were used to develop the Western Root Disease Model. In this procedure, serial workshops allow vari¬ ous experts in disease recognition, biology, and man¬ agement to meet with potential model users for short periods of intense interaction. Through the direction and assistance of model coordinators, they develop a conceptual model of the problem and possible manage¬ ment actions to mitigate damaging effects (Brookes 1985). The coordinator is then responsible for convert¬ ing this information into a working, predictive model that is further refined at subsequent workshops through additional input from specialists and potential users. The process itself is not new, but it creatively extends the scientific method from the individual in¬ vestigator to a corporate surrogate (Walters 1986). A recognized strength of the procedure is that it gives ownership of the final product, and thus a desire to have a quality item produced in a timely manner, to all who were involved with its development. Also, be¬ cause the model building is cooperative, scientists can appreciate the need to provide managers with the best current understanding of root disease spread and im¬ pact, and managers can recognize the critical uncertain¬ ties in our knowledge of root disease biology and the need for further research. The model was developed as a tool to aid foresters with overall, forest-level planning and with the design of silvicultural treatments in individual stands affected by root disease. The model can project the effects of various levels of Armillaria root disease on future stand composition and structure which, for timber purposes, can be converted into volume losses. The latter is particularly significant since current expecta¬ tions of timber yields over the next decade from certain forest areas in western North America may be over¬ estimated by 50% because effects of Armillaria root disease have not been considered. Even with the wealth of available empirical informa¬ tion, relationships that are appropriate for modeling disease behavior at the stand level need to be postulat¬ ed. In modeling the dynamics and behavior of Armill¬ aria root disease in forests of western North America, we found that the available information for specific components ranged from virtually no hard data to two or more conflicting data sets or opinions. Therefore, it became critical to document assumptions made during the modeling process because: (1) if model perfor¬ mance is questionable in certain areas, then the as¬ sumptions can be checked to see if they help to explain the concern; (2) if new information becomes available, then the current assumptions can be appropriately modified; and (3) if theory or concepts change in areas where little empirical information exists, then docu¬ mentation of initial assumptions is necessary to consid¬ er any possible changes. The model dynamically represents the spatial and tem¬ poral epidemiology of pathogenic Armillaria species or P. weirii (McNamee and others 1989, Stage and others 1990). It can project up to 40 growth cycles of stand development, normally of 10 years each, and operate in stands up to 100 ha. The three main components or submodels are root disease per se, "other agents," and an interface to a vegetation model (fig. 10.1). The root disease submodel provides the status and spread of forest management FIGURE 10.1. — Relationship among the three models of the Western Root Disease Model. Modeling Root Disease 151 root disease and contains a Keyword mechanism to modify relationships to meet particular conditions (Stage and others 1990). This feature allows the user to explore alternative hypotheses concerning root disease dynamics. The "other agents" submodel simulates the effects of wind-throw and three types of bark beetle behavior. This submodel is important because it struc¬ tures the interactions between root diseases and other mortality agents that can be important and damaging factors in forests of western North America (Shaw and Eav 1991). The stand-interface submodel links the stand-development model, to which the Western Root Disease Model must be attached, currently Prognosis (Stage 1973, Wykoff and others 1982), and the root disease and "other agents" submodels. Critical Model Relationships and Associated Assumptions and Hypotheses Spatial Resolution The Western Root Disease Model spans two levels of organization: individual trees and the aggregation of these individuals into stands. Within a stand, two stra¬ ta are defined with respect to root disease. The first consists of areas that are clearly beyond the influence of currently diseased trees. The second stratum consists of a number of root disease centers, each of which con¬ tains infected trees, uninfected trees, and other inocu¬ lum sources such as infected stumps. The size and separation of areas in these two strata define the spatial resolution of the model. Within each stratum, the actual spatial proximity of individual trees is not maintained. When rates of pathogen spread to uninfected trees are calculated, however, the individual trees in a sub-sample of the first stratum are assigned x-y coordinates according to whether the stand is of natural origin (a random distribution is assumed) or is evenly spaced as in a plantation. Center Dynamics The model addresses three important characteristics of root disease centers: the dynamics of infection and inoculum within root disease centers; the expansion of root disease centers; and the carry-over of root disease to a new stand following stand entry. Inside Established Centers Progression Within Single Trees The relationship that describes how live root systems become infected, trees are killed, and infection spreads in dead, infected roots (fig. 10.2) is a fundamental func¬ tion of the model. This relationship was developed from the experiences and judgements of those who participated in model development. Chapters 4 and 5 provide background information relevant to these as¬ sumptions. The relationship that describes the time between initial infection and death of a Douglas-fir tree on Douglas-fir habitat in the interior region of the Western United States is shown in fig. 10.3. This relationship is modi¬ fied for other species and habitat types, but the hypoth¬ esis is that all trees react similarly to infection. The relationship represents, with some reference to pub¬ lished information (Hadfield and others 1986), the best professional judgement of the pathologists who partici¬ pated in model development. They realized that, as modeled, the relationship may not be appropriate under situations of scattered mortality, a situation needing further research. For example, how is the per¬ centage of a root system that is infected when a tree dies affected by Armillaria species, tree species, stress, etc? In recognition of these uncertainties, critical points of the relationship can be modified using the Keyword system (Stage and others 1990). For example, one Keyword specifies the level of root infection at which trees die and allows users to vary the level for different tree species and sites. Another Key¬ word can be used to change the time-to-death for in¬ fected trees. A third Keyword allows users to modify infection and mortality dynamics by tree size. A consensus of pathologists in western North America suggested the values in table 10.1 for the average por¬ tion of a root system that is colonized by Armillaria when a tree dies. Following tree death, the model Infection in a tree total root death colonization decay FIGURE 10.2 — Pattern of root pathogen spread and inoculum buildup and decline in a single tree root system. 152 Modeling Root Disease infection levels causing death habitat species pathogen <^=o FIGURE 10.3 — Time required from infection by Armillaria to tree death for Douglas-fir on a Douglas-fir habitat type. assumes that Armillaria colonizes all remaining por¬ tions of infected root systems within 5 years. Support¬ ing evidence for this comes from Morrison (1981) and Shaw (1980). The model assumes that dead trees or stumps can only become inoculum if, prior to their death or cutting, their root systems were already colonized to some de¬ gree with a pathogenic species of Ay miliaria. Even if a tree only has a small lateral lesion somewhere on its roots (Shaw 1980), its entire root system will, under this assumption, become inoculum within 5 years. Contrar- ily, trees not already infected at cutting, regardless of location, will not become inoculum. Even though this modeling assumption contrasts with certain hypothe¬ ses about the competitive saprophytic ability of Armill¬ aria (see chapter 4, Garrett 1970), the model does provide sufficient inoculum for disease to progress in a manner judged to be reasonable by knowledgeable forest pathologists in western North America. This assumption may be logical for modeling in conifer¬ ous forests of western North America. These forests show limited rhizomorph development by pathogenic species TABLE 10.1 — Average percentage of root systems assumed to be infected at the time a tree is killed by root disease. Fungal Species Tree species Armillaria P. weirii Douglas-fir (% root system infected) 80 60 Pines 30 85 True fir 80 60 Hemlock 80 80 Spruce 75 65 Larch 00 75 W. redcedar 75 85 of Armillaria (Shaw 1980), and pathogenic lesions fre¬ quently occur on trees with little above-ground evidence of infection other than proximity to trees with obvious symptoms or signs of infection (see chapter 5). Perhaps the earlier assumptions on competitive saprophytic abili¬ ty, developed primarily in the United Kingdom, need to be re-examined regarding current information on the pathogenicity of various Armillaria species and their rela¬ tive in vivo abilities to produce rhizomorphs (see chapters 4 and 6). In the model, how disease spreads through root systems of dead trees is independent of how the trees died, even though in reality the speed and mechanism of death may affect either the proportion of the root system actually colonized or the viability of resulting inoculum. This assumption relates to a fundamental research need re¬ garding Armillaria root disease: the importance of, and mechanisms for, interaction of root diseases with other agents (both biotic and abiotic) of stress (see chapters 7 and 8; Shaw and Eav 1991). The maximum lifespan of effective inoculum also may be affected by habitat type or other environmental parame¬ ters; however, users can modify these parameters. As modeled, the lifespan of effective inoculum is assumed to be a function of stump size and species, with rather rapid deterioration after maximum build-up (fig. 10.4). Species are grouped into heartwood (Douglas-fir, pines, and larch) and nonheartwood types (true firs, hemlocks, arid spruce), with the latter decaying more rapidly. Inoculum is assumed to decay at a rate that reduces the radial extent of infected root systems by 75% during the first one-third of their lifespans. The remaining infected roots are as¬ sumed to decay at a steady rate over the remaining two- thirds of their lifespan. The pattern of inoculum decay is undoubtedly influenced by habitat type, tree rooting habit, temperature, moisture, and other abiotic factors not captured in the model but discussed in chapters 4 and 7. Modeling Root Disease 153 Spread from Tree to Tree Pathogen transmission to adjacent, living trees is mod¬ eled as a probabilistic process of two parts. First, an uninfested root system overlaps an infested system; and second, the pathogen will be transmitted given that the root systems overlap. The latter probability is controlled by a species-dependent Keyword. The first probability is calculated by simulation on a map which plots individual trees. Their spatial distribution is mod¬ eled as random (Poisson) for natural regeneration or a lattice for plantations, and can be changed by the user during the simulation. Kellas and others (1987) also suggest modeling tree distribution in Australian mixed species eucalypt stands with a Poisson distribution modified for stumps colonized by Armillaria. Pathogen transmission via rhizomorphs is not explicit¬ ly modeled though one can change infection probabili¬ ties. Thus, increasing the probabilities of root system overlap, or pathogen transmission given root overlap, could be used to accommodate the activity of rhizo¬ morphs where they are considered important agents of infection (see chapter 4). area is determined by the relation of root extent to tree diameter and species, with rooting patterns assumed to be circular. If rhizomorph networks extend the influ¬ ence of inoculum beyond the actual root systems boundaries (see chapter 4), then the areas could be increased. This action was considered to be unneces¬ sary in forests of western North America because of limited in vivo rhizomorph production by the primary pathogenic species (see chapters 4 and 6). As described above, the size of these areas declines with time to rep¬ resent the decay of root systems after tree death. We know these assumptions have inherent inaccura¬ cies and simplifications. For example, the density of roots is not uniform across a radius drawn outward from a stump, but decreases with distance from the stump. And, of course, trees, particularly when grow¬ ing on slopes, do not have roots with a uniform, circu¬ lar distribution. However, the important attribute is area, which is only slightly different for an ellipse than for a circle. A more detailed representation of actual root system geometry would have considerably com¬ plicated the model, with a limited likelihood of im¬ proving predictions. Quantity of Inoculum Expansion of Centers The quantity of inoculum available in a stand is esti¬ mated from the area occupied by infected roots. This Armillaria FIGURE 10.4 — Lifespan of Armillaria inoculum for trees with and without heartwood, see text for detailed description. The simulation for enlarging infection centers has two main components: estimating the average rate of en¬ largement and translating that rate into a new stand area encompassed by root disease. Rate of disease spread into uninfested area is simulated by a subroutine with enhanced spatial resolution. It maps a sample of the trees still outside the infection centers onto a smaller square area within which trees are assigned x-y coordinates. Then, the same relations that describe increases in area occupied by infected roots (fig. 10.2) and the probability of transmission, given that the root area of an uninfected tree overlaps that of a diseased tree, are used to estimate the time required for the disease to propagate across the width of the map. Width of map divided by the estimated time defines the radial spread rate of disease centers. The spread rate is thus not an input parameter but is calculated by the model. As such, it provides a means to gauge model performance as data on rates of infec¬ tion center enlargement are available (see chapter 8). As an option, one can override this model function and input a static spread rate. When this radial increment is added to the radius of each existing infection center, some centers may over¬ lap. However, the new area of infection is calculated after adjusting for overlaps. The increase of infected area divided by the previously uninfected area pro- 154 Modeling Root Disease vides a proportion for moving trees from the uninfect¬ ed tree inventory to the infected tree inventory. Carryover to Regeneration after Harvest How root disease centers are affected by clearcutting and regeneration of a new stand is poorly understood. The modelers considered three different "carryover" scenarios: (1) root disease centers from the former stand cease to exist after clearcutting, and root disease in the new stand arises in a small number of new cen¬ ters located within previously infected areas; (2) root disease centers from the former stand retain their integ¬ rity, and, as the new stand matures, these centers en¬ large, starting at their old boundaries; and (3) after a clearcut and regeneration, root disease centers form around certain individual pieces of inoculum through¬ out the area affected in the former stand and these centers gradually expand and coalesce. These three scenarios actually form a continuum that depends on inoculum density and the probability of a piece of inoculum initiating a new center that is capable of expanding. New disease centers have equal probability of occur¬ ring anywhere root disease occurred in the previous stand and no probability of occurring elsewhere. This assumption implies that all disease in the previous stand was noticeable and detected in the stand exami¬ nation (see chapter 5) and that spores do not initiate new centers. We realize that the latter event must occur at some time (see chapter 9). However, for Armillaria root disease in western coniferous forests, in contrast to root disease caused by H. annosum in these same forests (Shaw and others 1989b), its occurrence seems to be infrequent enough that it can be ignored for stand-level modeling purposes—particularly when modeling stands that are already infected. Information presented in chapters 7, 8, and 9 supports this view. A ring of root systems around the outside of each dis¬ ease center represents trees that have just been infected. When these trees are cut, the prompt colonization of their entire root systems by the fungus causes disease centers to expand rapidly. In the model, the mean di¬ ameter of all root systems in the stand at the time of the cut is the distance by which radii of disease centers increase. Evidence for such action is found in Morrison (1981) and Shaw (1980). A major unknown is how far root disease actually does "jump out" from the recog¬ nized, above-ground edge of a center after clearcutting. Cursory examination of model behavior suggests that this is a sensitive parameter and thus it can be con¬ trolled by use of a Keyword. This feature of the model was most useful in preliminary work on adapting the Western Root Disease Model to represent root disease caused by H. annosum (Shaw and others 1989b). Representation of Management Actions Inoculum Removal The Western Root Disease Model can simulate inocu¬ lum removal through "pushing" or removing infected stumps and their root systems (see chapter 11). This option can be requested in a specific year, with a factor specifying the efficacy with which roots are removed and the minimum diameter of dead trees and stumps to be removed. Even though this practice is an accepted management alternative in certain stands (Roth and others 1977), it is not universally applicable (Wargo and Shaw 1985). Silvicultural Treatments Regeneration systems ranging from single-tree selec¬ tions to clearcutting can be simulated. Besides harvest¬ ing existing trees, new stands can be introduced following site preparation either by natural regenera¬ tion or by planting with species selected for disease resistance (see chapter 11). Likewise, particular species can be favored during thinning or during other partial stand harvests. A full range of treatment alternatives may be consid¬ ered when regenerating diseased stands, depending on economic constraints and stand management objec¬ tives. The most frequent approach to managing root disease problems in timber stands throughout western North America is regeneration to site-suited tree spe¬ cies that are disease tolerant (Hadfield and others 1986, Morrison 1981). The model can be used to compare the effects of various approaches. For example, the follow¬ ing options are among the many that may be compared and considered: — No action—leave the stand "as is," but recognize presence of root disease. — A clearcut, seed-tree cut, or shelterwood cut fol¬ lowed by natural regeneration. The mixture of species in the resulting regeneration will depend on the habitat type. — Overstory removal to leave an understory of tree species that might be disease susceptible, disease tolerant, or a mixture of the two. — A clearcut with stump removal followed by plant¬ ing of a disease-susceptible but otherwise preferred species. — A clearcut without stump removal followed by planting of a disease-susceptible but otherwise preferred species. Comparison to the preceding option provides an estimate of the control value of stump removal (see chapter 11). — A base simulation, without invoking the Western Root Disease Model, of the Prognosis model for Modeling Root Disease 155 stand development, perhaps followed by planting of a preferred but disease-susceptible tree species. These scenarios represent "control" simulations for the Western Root Disease Model. Besides aiding management decisions, gaming with the model by preparing such scenarios can help scientists identify research needs relating to treatment alterna¬ tives. For example, the efficiency of stump removal (i.e., the proportion of inoculum removed) is a sensitive parameter in the model which suggests that carryover of root disease as a function of stump-removal efficien¬ cy is an important research area, particularly since little information exists on the topic. What Data are Required? The model is designed to start with sample inventories of actual stands. For example, the compartment exami¬ nation procedure described by Stage and Alley (1972) and in the Forest Service Handbook for Region 1 (USDA Forest Service 1986) can supply the necessary stand data if it is augmented to include stumps infected with root disease (see chapter 5). Besides the customary tree-size attributes, the model uses information on the frequency of tree infection by root pathogens. This value can be compiled by the model from disease status codes of the individual sam¬ ple trees, or supplied by the user from an overall esti¬ mate based on an independent sample of the stand. The Western Root Disease Model also uses data on the area of the stand and the sizes and distribution of dis¬ ease centers to initiate the simulation. The user may specify a total area in root disease and the number of centers. In this case, the model randomly locates root disease centers throughout the stand. Initially, each center will be of equal size, calculated as the total area in root disease divided by the number of centers. The alternative is to provide a list of root disease centers with X and Y coordinates and a radius for each center. The model can start from bare ground by planting, by invoking the Regeneration Establishment component of the Prognosis model for stand development (Fergu¬ son and others 1986), or from the stand description contained in the list of trees sampled in the inventory. Conclusions We believe that the Western Root Disease Model pro¬ vides a workable framework for others to consider when modeling the behavior of Armillaria root disease in orchards or other forest situations. The current mod¬ el should continue to improve as new information be¬ comes available. The model is currently undergoing an analysis of its sensitivity to changes in the various pa¬ rameters that control it and thus the assumptions and hypotheses under which it was developed (Marsden, unpubl.). We believe that the items to which the model is the most sensitive (longevity of inoculum and the quality, quantity, and type of input data) are the ones where additional resources could best be put to im¬ prove model performance. Thus, a list of research needs relevant to improving model reliability can be generated through a structured sensitivity analysis. If the procedure we have outlined is used in model development, then it is critical that participants in the process represent a cross section of interested and knowledgeable scientists, managers, and administra¬ tors. Furthermore, it is paramount that scientists be willing to extrapolate beyond the limits of available data to help meet existing management needs. In so doing, however, they must insist that all extrapolations and assumptions are thoroughly documented. Also, such disease models need to be developed so that they can function in concert with existing models that may predict other stand or orchard attributes such as yield or watering regimes. Based on how well users have accepted the Western Root Disease Model for both short-term, site-specific management decisions and long-range applications in planning, we strongly encourage others to pursue this avenue for transferring technology on Armillaria root disease dynamics into a useable tool for managers. We also contend that the process of doing so will help sci¬ entists clarify the current state of knowledge and help to focus management-oriented research needs. 156 Model in g Root Disease CHAPTER 11 Avoiding and Reducing Losses from Armillaria Root Disease Susan K. Hagle and Charles G. Shaw III s the forest and agricultural land base is in¬ creasingly utilized, careful stewardship of remaining productivity becomes increas¬ ingly important. Armillaria epiphytotics can not only cause marked reductions in fruit and fiber production (see chapters 8 and 9), but they may also carry high economic, social, and ecological costs for con¬ trol. In some forest settings, properly applied cultural control methods are efficient and effective. But for many crops, we lack convenient, cost-effective methods for control. In fact, it has been said (Schiitt 1985) that while our biological knowledge about Armillaria has increased markedly since the time of Hartig, the efficiency of con¬ trol measures, with some exceptions, has not improved very much. However, advances in our ability to identify species accurately (see chapters 1 and 2), determine their relative pathogenicities (see chapter 6), and model the disease process (see chapter 10) provide us a sharper image of disease problems, and should allow a more systematic evaluation of control options. In this chapter, we examine various approaches and techniques for control and avoidance of Armillaria root disease in forests, orchards, and amenity plantings. These include use of resistant species, avoidance of haz¬ ardous sites, cultural manipulation, chemical applica¬ tion, biological methods, and integrated biological methods. Chapter 9 contains related material on man¬ agement practices in plantations that can reduce losses from Armillaria root disease. Armillaria species cause three types of disease in indig¬ enous forests (see chapter 8). In one type, tree and shrub species are attacked and killed by an aggressive, pri¬ mary pathogen (Filip 1977, Gibson 1960, Kile 1981, MacKenzie and Shaw 1977). In another type, the fungus lives primarily in chronic infections it causes on roots that may but seldom become aggressive. The third type causes butt rot that may or may not be related to other disease types (see chapters 5 and 6). The first disease type often requires radical measures to effect control. The other two types may cause little dam¬ age if stand management maintains the resistance or tolerance of infected trees. In these latter two, a shift in the balance between the host and the pathogen induced by stresses such as drought, insect attack, other dis¬ eases, or anthropogenic activities can allow the fungus to expand and kill the host (see chapter 7). The pathogenic behavior of Armillaria species in planta¬ tions, orchards, and amenity plantings also ranges from aggressive to benign. Control options may, how¬ ever, differ from those that are feasible in indigenous forests, and they also may be more costly; however, the higher commodity values may offset higher costs (see chapter 9). The type of root disease expression (see chapter 5) and the extent of damage are related to species and geno¬ types of Armillaria (Shaw and others 1981, Guillamin and Lung 1985, Rishbeth 1982, Kile and Watling 1988, Roll-Hansen 1985, Intini 1989a), inoculum characteris¬ tics (see chapter 4), inherent host resistance or tolerance (Thomas and Raphael 1935), host adaptation to site (Intini 1989a, Singh and Richardson 1973), stand struc¬ ture and species composition, management history, and site factors which directly affect the pathogen (Redfern 1978, Blenis and others 1989; see chapter 6). Where Armillaria acts as a secondary pathogen on plants that are predisposed in some way, control efforts should focus on the predisposing condition (see chap¬ ter 7). This problem becomes especially acute in situa¬ tions such as those created by atmospheric deposition or photochemical oxidant injury, which are not only difficult to document but also difficult to control for so¬ cietal reasons. In such cases, we could find ourselves treating symptoms at great expense with little benefit. Where Armillaria is a primary pathogen, infection often leads to rapid death even if the plants were vigorous prior to attack. This distinction in pathogen behavior generally determines the type of control measure to use. Cultural controls often are used to reduce damage to natural stands or plantations of indigenous species. Reducing Losses 157 while direct methods of inoculum removal, either alone or in combination with cultural control methods, may be required to reduce damage in plantations, arboreta, seed orchards, or amenity plantings. Diseases can be avoided by using the natural balance and diversity of indigenous forests to prevent Artnil- laria epidemics, even though the fungus is present as a minor pest and natural thinning agent. Examples occur worldwide where Armillaria causes insignificant dam¬ age in indigenous forests but inflicts significant losses when these forests are cleared to establish exotic plan¬ tations (see chapters 8 and 9). The cost, both economic and ecological, of converting indigenous forests to ex¬ otic plantations must be weighed against any increased commodity value derived from the exotic species. Needs Assessment In many situations, Armillaria may be present in a for¬ est or orchard and cause little damage. Thus, mere fun¬ gal presence is not sufficient cause to treat. Plants that are resistant to Armillaria throughout their lives or, as is the case with many conifers, through most of their lives, are capable of maintaining stand or orchard pro¬ ductivity. In such cases, Armillaria may act as a thin¬ ning agent in young stands (Filip and others 1989, Morrison 1981, Rishbeth 1972a) and as a nutrient recy¬ cler in old stands (Durrieu and others 1985, Mason and others 1989, see chapter 8). Disease often is severe in the first few years after plantation establishment, but subsides thereafter. Where this happens, primary in¬ oculum from stumps or other buried woody material is the likely source of disease; secondary inoculum is not effective. Disease in New Zealand's radiata pine plan¬ tations (Roth and others 1979), in western North America's young ponderosa pine stands, whether planted or naturally regenerated (Hadfield and others 1986, Hagle and Goheen 1988, Morrison 1981), and in Europe's first-rotation conifer plantations on cleared hardwood sites (Hartig 1873b, Nechleba 1915, Pawsey 1973) follow this pattern. A high incidence of mortality following establishment may be alarming, but without secondary spread of dis¬ ease, the economic impact may be insufficient to justify control. Such is generally the case in indigenous pon¬ derosa pine and coastal Douglas-fir stands of western North America (Morrison 1981, Hadfield and others 1986) and in many first-rotation conifers on former hardwood sites in Europe (see chapter 9). In contrast, radiata pine plantations on high-risk sites in New Zealand may lose 50% of the crop within the first 5 > ears after planting (van der Pas 1981b) which consti- utes a severe impact. Direct reductions in primary in- i 'urn (fig. 11.1) may be economically feasible in such FIGURE 11.1 — Reduction of primary inoculum by removal of stumps and roots of indigenous forest cover in New Zealand prior to establishment of radiata pine plantations. Such actions can markedly reduce disease incidence and severity in first rotation crops (see fig. 9.7). (C. Shaw) cases; even so, other alternatives also should be consid¬ ered. For example, increased planting densities that al¬ low for full stocking after suffering losses due to primary inoculum may, if effective, prove more eco¬ nomical and environmentally acceptable than efforts to reduce inoculum levels through stump removal or chemical treatment at the beginning of the rotation. Patchy killing of trees in the plantation may make thin¬ ning the remaining stand necessary after mortality has subsided. Contrarily, in orchards and amenity plantings, the economic importance of losing a few or perhaps even a single tree may be sufficient to justify inoculum removal or other costly control procedures. The lack of assessment data in these situations (see chapter 9) complicates decisions to implement control. As indicated by Rishbeth's survey (1983) of gardens and forests in southern England, the species of Armillaria found on a stump, tree, or shrub can affect the decision to initiate control. For example, A. gallica had spread widely from an ash stump in a garden with no signs of attacking other trees or shrubs that it had encountered. However, A. mellea had spread from a Primus stump and killed species of apple, stonefruit, birch, and sequoia. As identification of Armillaria spe¬ cies becomes more routine (see chapters 1 and 2), its use is likely to become standard before control is recommended. Control Options Silvicultural Considerations for Natural Forests In natural forests, silvicultural control of Armillaria root disease is frequently an option. Local tree species grown in natural mixtures and densities may resist 158 Reducing Losses Armillaria root disease even though they are known hosts for the local species of Armillaria. Where Armillaria root disease is a major concern in coniferous forests in western North America (see chapter 8), only indigenous tree species are grown in production for¬ ests. Even so, careful selection among species, seed sources, and cultural methods to match site conditions, particularly those related to habitat type (Daubenmire 1952) or site type (Corns and Annas 1986), is necessary to avoid economic losses. Using locally adapted seed sources for indigenous species that tolerate the disease is recommended for control in the Northwestern United States and Western Canada (Hadfield and oth¬ ers 1986, Morrison 1981, Williams and others 1989). Substantial losses occurred in mixed coniferous forests of southern Oregon after selective harvest of ponderosa pine overstories (Filip 1977) caused a species composi¬ tion shift to highly susceptible true firs. Severe root dis¬ ease problems have been attributed to similar changes in species composition over much of the Western United States due both to selective logging of pines and larch and to fire suppression which favored shade tol¬ erant true firs and Douglas-fir (fig. 11.2, see chapter 8). Dense Douglas-fir and true fir forests are unnatural on these sites and their development often results in dis¬ ease conditions much like those found in exotic planta¬ tions. Current silvicultural practices in such areas, developed in part to reduce root disease losses, aim to re-establish pine, larch, and pine/larch mixtures with Douglas-fir and true firs composing less than 40% of the regeneration (Hagle and Goheen 1988). Even when planted within their natural range, some species adapt poorly to certain sites. Although Dou¬ glas-fir is well distributed over diverse montane envi¬ ronments, the species has differentiated populations that are closely tied to elevation, latitude, and longi¬ tude (Monserud and Rehfeldt 1990, Rehfeldt 1982). Each population has adapted to local environmental conditions and fails to thrive when planted elsewhere. Other conifer species appear to behave similarly (Balmer and Williston 1983, Lotan and Perry 1983, Rehfeldt and others 1984). Thus, attention to seed sources is important for culturing these species. For ex¬ ample, substantial increases in Armillaria damage to Scots pine plantations in the German Democratic Re¬ public followed a drought in 1969. Even after the drought, however, wildling pines were seldom affected by the pathogen, leading Kessler and Moser (1974) to recommend development of seed-saving methods to take advantage of natural resistance by regenerating stands through seeding with these sources. Where use of locally adapted seed sources is not an op¬ tion because the natural forests were removed, genetic differentiation within artificial populations can be used. Lung-Escarmant and Taris (1989) reported a method to test the Armillaria resistance of various pine species in natural stands. They suggested using the FIGURE 11.2 — A natural forest in western Montana where ponderosa pine is more resistant to Armillaria root disease than most associated species. Al¬ though few large pon¬ derosa pines remain in the overstory, past manage¬ ment practices that favored removal of ponderosa pine and excluded natural fire have allowed a Douglas-fir and true fir understory to develop that is more susceptible to Armillaria root disease. (S. Hagle) Reducing Losses 159 method to test Maritime pine for population, family, and clonal differentiation in resisting A. ostoyae within the pine's natural range in southwest France. Whether considering indigenous or exotic trees, genetic differen¬ tiation should be matched to the natural site conditions where the trees are growing. Intraspecific variation in adaptation to sites may be great, but the extension of a species' range may still be limited. For example, die- back and declines of silver fir plantations in central and northern Italy are frequently associated with infection by A. ostoyae. The diseases appear to be drought-trig¬ gered and to be concentrated in fir plantations estab¬ lished in an area "phytoclimatically inferior and warmer" than natural sites for the species (Intini 1989a). Excessive moisture may have been responsible for the demise of several ponderosa pine plantations which were established during the 1940's in Idaho using non¬ local seed sources (Hagle unpubl.). The parent trees, growing more than 500 km away, were on very differ¬ ent sites than those where the plantations were estab¬ lished. The plantations were installed to determine if genotypes adapted to dry pine sites would produce su¬ perior growth when planted on more mesic, grand fir climax sites. The trees grew exceptionally well for about 40 years but died rapidly thereafter from a com¬ bination of pests, among which Armillaria root disease was most prominent (Hagle unpubl.). Ponderosa pine has since been found to have "seed zones" of limited range (Squillace and Silen 1962) and planting outside these zones is not recommended. McDonald (1990) dis¬ cusses how the potential for ecophysiological maladap- tation of species to specific sites may influence their susceptibility to Armillaria root disease. Avoiding Hazardous Sites Matching indigenous species with suitable sites is one way to minimize disease hazard. Sites can be hazard¬ ous because they predispose the host in some way, as with off-site plantings. Sites with heavy inoculum loads of pathogenic Armillaria species may also be haz¬ ardous. Whether a naturally high frequency of Armillaria infections occurred in the previous stand or human or other activity increased the level of infection, the influence of inoculum loading is little disputed (see chapter 4). Hazardous sites also may result from site conditions that are unusually favorable to disease de¬ velopment; however, these conditions are difficult to discern because of our limited knowledge of Armillaria ecology. Site hazard varies within indigenous forests. For example, soil-related differences in disease severity were reported in Norway spruce stands in central Eu- r< ; e (Gramss 1983). Enhanced survival of stands was Gv attributed to the "poor podzolic highland soil types" to which the spruce appeared to be better suited. The highland soils distinguish low-hazard sites for growing Norway spruce. Ono (1970) found edaphic, topographic, and vegetational relationships with the level of Armillaria damage to Japanese larch plantations in Hokkaido, Japan. Williams and Marsden (1982) related root disease patch occurrence in conifer¬ ous forests in Montana and Idaho to certain productive soil and habitat types. Byler and others (1990) found that the more productive habitat types in these areas had greater root disease severity than the less produc¬ tive types, a result that was partially supported in a preliminary report by McDonald and others (1987a). Damage in exotic plantations or orchards can be mini¬ mized by establishing them on sites with a low disease hazard. Sokolov (1964) reported soil types in the Soviet Union influenced the severity of Armillaria root dis¬ ease in mulberry plantations. Severe Armillaria root disease in Norway spruce stands in Poland was related to a combination of soil type and elevation (Manka 1980). The spruce plantations were established on sites previously supporting indigenous stands of silver fir and common beech. Whitney (1984) found Armillaria root disease to be more severe on conifers in Ontario, Canada, where soils were coarse-textured and sandy rather than finer-textured and silty. Vegetation on a site prior to clearing for establishing an orchard or plantation may indicate differences in root disease hazard. For example, Leach (1939) reported site hazard differences for tea plantations associated with indigenous stands of Muula trees in eastern Africa. Muula roots remain alive but moribund for years after cutting, and these were most often associated with root disease in tea plantations. In Kenya and New Zealand, differences in root disease severity also were noted on lands converted to radiata pine from different indig¬ enous forest types (Gibson 1960, Shaw and Calderon 1977). Hendrickson (1925) reported Armillaria root disease in fruit orchards to be of "widespread economic impor¬ tance in California." He considered oak roots from the indigenous forest to be the most important source of inoculum, but secondary spread among orchard trees maintained the disease long after the land was cleared of oaks. Cooley (1943) surveyed Eastern U. S. fruit or¬ chards and found little disease except in the sandhill section of North Carolina. Most orchards in this area had been established on land cleared of hardwood for¬ ests, indicating that a heavy load of primary inoculum that harbored a pathogenic species of Armillaria may have been responsible for the frequency of disease. Or¬ chards themselves may maintain a high hazard from one rotation to the next. For example, Armillaria inocu- Reducing Losses lum from a highly susceptible cherry rootstock created hazardous site conditions in replanted orchards of sev¬ eral species in Michigan (Proffer and others 1987). Vegetation maintained on a site from one crop to the next may affect disease hazard. Although conifer plan¬ tations in Britain are damaged by Armillaria in the first 10-15 years after planting on sites freshly converted from hardwoods, succeeding rotations of conifers sus¬ tain much less damage (Pawsey 1973). Balsam fir Christmas tree plantations are damaged in the first few years after establishment on sites converted from indig¬ enous mixed hardwood and pine forests (Wargo and Houston 1987). Similarly, radiata pine plantations in New Zealand may be severely damaged within 5 years after conversion from indigenous forest but subsequent rotations of pine on the sites may be little damaged (Shaw and Calderon 1977). If this effect is found to be consistent, such areas would be high-hazard sites only for the first rotation after conversion. Recent work in New Zealand, however, suggests that the cursory ob¬ servations on limited disease incidence in second- or third-rotation pine crops by Shaw and Calderon (1977) may have been premature (MacKenzie and Self 1988). Resistance In situ host resistance to root disease is a complex topic as it involves the genetics of both host and pathogen as well as environmental influences. It also can involve managing mixtures of genotypes with varying levels of resistance. Some species with superior resistance or tolerance to infection in one location may be quite sus¬ ceptible in other locations. For example, Douglas-fir is among the species recommended for planting in Britain where Armillaria root disease is especially damaging to Scots pine plantations (Greig and Strouts 1983). It is also considered more resistant than Sitka spruce and grand fir in France (Delatour and Guillaumin 1985). Within its natural range in western North America, however, Douglas-fir is considered rather susceptible to Armillaria root disease (Hadfield and others 1986, Hagle and Goheen 1988). In all three locations, A. ostoyae is the most common parasite of conifers (Rishbeth 1982, Guillaumin and Lung 1985, Morrison and others 1985a). Even within limited geographic areas, such discrepan¬ cies in resistance can be seen. For example, ponderosa pine's resistence is superior to true firs and Douglas-fir over most of western North America (Morrison 1981, Hadfield and others 1986), but natural ponderosa pine stands in some south-central Washington, sites are se¬ verely damaged by Armillaria root disease (Shaw and others 1976a, Shaw and Roth 1976). Douglas-fir associ¬ ated with pine on these sites suffers less damage (Roth and Rolph 1978). A similar situation exists in the Jemez Mountains of northern New Mexico (Wood 1982, Shaw unpubl.). Thus, intraspecific variation in adaptation to sites may be as great as interspecific variation within the natural ranges of any two or more species. Radiata pine planted in New Zealand suffers consider¬ able damage from Armillaria root disease (Shaw and Calderon 1977), but the same species is not particularly damaged in its natural range in western North America (Raabe 1979a). This variation is probably related to dif¬ ferences in Armillaria species, inoculum loads, or edaphic, climatic, or physical site characteristics. Whether endemic or exotic, genetic differentiation in tree species should be compared to site conditions and pathogen species in natural settings in which plants are to be grown, and plantations should be monitored for suitability of the genotypes to the site. Diseases and in¬ sect attack are likely to be among the earliest indications of poor compatibility of trees with growing sites. Relative resistance of many species has been observed in forests, orchards, and parks as well as by controlled inoculation experiments (see chapter 6). Much of this work was completed before many of the pathogenic Armillaria species were recognized. As such, the infor¬ mation is only useful in a general way. Morquer and Touvet (1972b) noted considerable variation in resis¬ tance of several conifers, but found no species immune to infection by Armillaria. Pines were notably suscep¬ tible while certain ecotypes of Norway spruce were relatively resistant. Mugala and others (1989) found that white spruce succumbed more readily than lodgepole pine in potted-seedling studies, but these results were inconsistent with field observations where white spruce was damaged less than lodgepole pine (Blenis and oth¬ ers 1987,1989). In Kenya, Gibson (1960b) noted that slash pine was more severely affected than either ra¬ diata or Mexican weeping pine. Rishbeth (1972a) re¬ ported that, in a 17-year-old mixed stand of Scots pine and Norway spruce, large patches of pine died while the spruce was virtually unaffected. Day (1927b) ob¬ served that, in adjacent 3-year-old plantations of Scots pine and Japanese larch, the pines were more frequently infected than larch but relatively fewer pines were killed. Up to 10% of the larch were killed by Armillaria. Resistant rootstocks have been developed for both fruit and fiber species (Raabe 1966a). Rhoads (1948) reported considerable variation in susceptibility of citrus root¬ stocks to Armillaria and Thomas and others (1948) tested rootstocks of prune and apricot for resistance in Califor¬ nia. Armitage and Barnes (1968) reported that loblolly pine resisted Armillaria, and a heterospecific graft of slash pine onto loblolly rootstock was sufficiently resis¬ tant to replace slash pine killed by the fungus. Peach, al¬ mond, apricot, and cherry trees are severely damaged by A. mellea in France, while plum is generally resistant Reducing Losses 161 (Guillaumin and others 1989b). The use of resistant rootstocks appears to be the only practicable control op¬ tion in French stone fruit orchards (Guillaumin and oth¬ ers 1989b). Two rootstocks in particular, resulting from interspecific crosses between diploid plum and peach, appear to satisfy both Armillaria resistance and other cultural demands. Heaton and Dullahide (1989b) rec¬ ommended a number of Armillaria-resistant plum, pear, and grape rootstocks for Granitebelt orchards in south¬ ern Queensland, Australia. Perhaps gene manipulation techniques can improve a species' adaptability and physiological resistance to dis¬ ease, produce populations or clones immune to Armillaria, or improve economic qualities of endemic, resistant species (Hubbes 1987). Rootstocks for fruit or¬ chards are prime candidates for receiving resistance genes. Superior quality rootstocks which produce de¬ sired growth and compatibility characteristics could be made resistant to Armillaria if genes known to produce successful resistance reactions in other species or geno¬ types can be identified and transferred to the genome of the otherwise superior rootstock. Research to determine the relative resistance of different species and geno¬ types under a variety of conditions must continue. Ad¬ ditional information is also needed on the nature of resistance (i.e., physiological, genetic, or environmental) and its interaction with various Armillaria species and genotypes (see chapter 6). Using species resistant to Armillaria may be economi¬ cally practicable in some cases, and their suitability may be enhanced by combination with other control procedures. Planting mixtures of species with differing resistance to Armillaria may reduce secondary spread of disease in a plantation. For example, Morrison and oth¬ ers (1988) observed fewer and smaller disease patches in plots planted with highly susceptible lodgepole pine or Douglas-fir in alternating rows with resistant west¬ ern redcedar or paper birch, compared to plots planted only to lodgepole pine or Douglas-fir. Presumably the benefit was derived from limiting secondary disease spread, which limited the size of infection centers. Other Cultural Considerations Regeneration methods may influence tree condition and susceptibility to Armillaria root disease. For ex¬ ample, Newfoundland conifer plantations that origi¬ nated from bareroot stock were significantly more damaged by Armillaria than those that had been broad¬ cast seeded (Singh and Richardson 1973). Similar re- i ts were reported for Scots pine plantations in the ■ wm Democratic Republic (Kessler and Moser 1974). is not always acceptable, however, because of ivantages that accrue from planting (Page 1970, ■ hubert and others 1970). Manipulation of rotation length may minimize losses in some situations. As previously mentioned, Norway spruce may be only moderately affected by Armillaria root disease. Still, in Czechoslovakian plantations, diameter growth significantly decreased in 70- to 80-year-old spruces (Hrib and others 1983). The current recommendation is to harvest stands by this age, which represents the culmination of mean annual increment. Such pathological rotations are used primarily for eco¬ nomic reasons. They often are an option in hardwood forests in England (Greig and Strouts 1983) and the Eastern United States (Marquis and Johnson 1989), and conifer stands in Ontario, Canada (Whitney 1988b). Stands or individual trees are harvested prior to the age at which they are expected to succumb to Armillaria root disease (Marquis and Johnson 1989). A similar principle underlies a recommendation to use training and trellising techniques to promote early cropping in Australian fruit tree orchards where sec¬ ondary spread of Armillaria causes high losses (Heaton and Dullahide 1989b). This practice allows an early re¬ capture of investments and minimizes the economic impact caused by eventual loss of trees to Armillaria root disease. Both precommercial and commercial thinning to re¬ duce damage from Armillaria should be considered ac¬ cording to the type of disease caused. Where host condition has little influence over infection and killing by Armillaria, thinning is unlikely to reduce losses and may increase damage by providing additional food bases. Precommercial and commercial thinning to re¬ duce intertree competition may effectivelv reduce dis¬ ease losses where Armillaria is a secondarv pathogen, as it is in hardwood forests in southern England and the Eastern United States. Suppressed oaks and pines (completely shaded by overtopping trees) were exten¬ sively infected by A. mellea and A. ostoyae in southern England. Subdominant (codominant) trees in those stands were only slightly infected and apparently re¬ sisted extension of infection by these pathogens (Davidson and Rishbeth 1988). Red spruce stands in the Northeastern United States have been severely damaged by Armillaria root disease following stagna¬ tion from overstocking (Wargo and Shaw 1985). Early thinning has avoided such problems in most managed stands. Precommercial thinning of conifer stands in western North America has produced mixed results (Hagle and Goheen 1988). In one ponderosa pine plan¬ tation, mortality rates in thinned and unthinned plots were similar 20 years after precommercial thinning (Filip and others 1989); however, net productivity on the thinned plots increased because of superior volume growth. Singh (1981a) recommended precommercial thinning in pine and spruce stands in eastern Canada to reduce stress and improve resistance to Armillaria. Reducing Losses Morrison (1981) advised delaying precommercial thin¬ ning in western Canada to 30 years in infection centers of conifer stands where losing residual trees after early thinning could cause low stocking. Blenis and others (1987) observed infections in young lodgepole pine stands in Alberta, Canada, and noted that rhizomorphs spread from the previous stand's debris. They con¬ cluded that precommercial thinning could be done af¬ ter such debris was no longer effective as inoculum. Interpreting how Armillaria root disease responds to commercial thinning or other forms of partial harvest is difficult because stands are often affected by mixtures of two or more root pathogens capable of varying re¬ sponses. However, in ponderosa pine stands infected with Armillaria alone, a partial harvest increased mor¬ tality even though the total area affected had remained unchanged (Shaw and others 1976a). Rishbeth (1978b) reported indirect evidence of A. mellea spores coloniz¬ ing oak and ash thinning stumps and thereby causing a low incidence of new infection patches. On this basis, he recommended delayed thinning. Such a delay might, however, lead to even larger and more numer¬ ous new infection patches through spore infection of even larger stumps. Severe damage has occurred in New Zealand kiwifruit orchards following spore infec¬ tion of stumps created by removing windbreak trees (Horner 1988). Kellas and others (1987) surveyed for A. luteobubalina infection in mixed eucalypt stands in Victoria, Austra¬ lia, where shelterwood cutting has become common. Cutting intensity did not appear to influence the inci¬ dence of disease in residual trees and stumps, but fre¬ quent, low-intensity cuts may have increased infection. The high infection frequency on residuals in thinned stands also may have reduced growth and increased mortality sufficiently to negate any growth response in the thinned stands. Partial harvest for commercial thinning or other pur¬ poses has caused considerable damage to conifer stands in the Western United States. Byler and others (1986) reported that root disease frequency doubled in stands that had at least one harvest entry compared to those with no tree cutting. Losses to root diseases have been so great following partial harvests which leave stands composed primarily of Douglas-fir or true firs (fig. 11.2) that these silvicultural methods are not rec¬ ommended on sites prone to root disease in the West¬ ern United States (Hadfield and others 1986, Williams and others 1989). Such management activities have truly exacerbated the incidence and severity of Armillaria root disease. Soil pH, organic content, and nutrient status are par¬ tially alterable in forestry and orchard operations. Very little direct evidence, however, establishes the effects of fertilization or other soil amendments on Armillaria root disease in forest crops. This may be, in part, be¬ cause the nutritional requirements of forest species are not well understood. Rykowski (1981a) was not able to decrease mortality rates by applying fertilizers in young Scots pine plantations, but the appearance of chronically infected trees did improve. Fertilization increased the fungistatic effects of extracts from periderm and phloem tissues but also improved the nutritional quality of wood used during the saprophytic phase of Armillaria colonization (Rykowski 1983). Application of potassium did, however, significantly reduce damage from Armillaria in banana plantations in Malawi (Spurling and Spurling 1975). Use of fertilizers as a management tool in orchards and plantations is further discussed in chapter 9. Direct Reduction of Inoculum On the speculation that buried roots and stumps were a source of infection in young orchard trees, Barss (1913) suggested that stumps and roots be removed and that non-orchard species be cropped for several years before establishing an orchard. Since then, recommendations for physical removal of inoculum (Shaw and Roth 1978, 1980) have involved removing diseased trees, uprooting stumps (Roth and others 1980, Arnold 1981), destroying stumps and root remnants (Morrison and others 1988), and turning the earth over to a considerable depth (Heaton and Dullahide 1989b, Horne 1914, McGillivray 1946, Reitsma 1932, Sokolov 1964). The quantity and location of inoculum that must be re¬ moved to prevent disease buildup and spread and the cost of removal justified by the crop's future value are difficult to balance. Complete eradication of the fungus by mechanical, biological, or chemical means is improb¬ able (Williams and others 1989) and of doubtful value. Even sites that have been devoid of woody material for decades, such as land supporting an herbaceous crop, may be re-invaded, albeit slowly, through spore infec¬ tion when placed into woody plant production (Rishbeth 1978b). Soil disturbance may also stimulate fresh rhizomorph production, increasing the disease risk to newly planted trees (Morrison 1976, Redfern 1970). Excessive removal of woody debris from a site may be detrimental to mycorrhizae as well (Harvey and others 1981, Maser and others 1984). Such additional risk must, however, be evaluated in context with the large quantities of inoculum removed. Stump and root removal is commonly practiced in pre¬ paring sites for fruit orchards and other high-value crops, whether converting from indigenous forest or re¬ moving infected trees from the previous crop. This pro¬ cedure has long been standard where Armillaria is a Reducing Losses 163 recognized threat (Wallace 1935, Thomas and Raphael 1935, Hendrickson 1925). In forests, Sokolov (1964) rec¬ ommended inoculum removal in the USSR, and Morrison and others (1988) successfully reduced inocu¬ lum on an infested site in western Canada. The latter treatment involved pushing the trees, roots and all, from the ground followed by a raking which removed most roots over 1.5 cm in diameter. Douglas-fir, lodge- pole pine, western redcedar, and paper birch were planted on treated and untreated sites. After 20 years, mortality of Douglas-fir and lodgepole pine in untreated plots was 5-10 times greater than in treated plots. Areas within a managed, natural ponderosa pine forest in south-central Washington were so severely damaged by Armillaria that they had little likelihood of persisting as a commercial forest unless the disease was con¬ trolled (Shaw and others 1976a). Here, recently killed sapling and pole-sized trees, and those positioned where they were likely to be infected (Shaw 1980, Shaw and Roth 1974), were pushed over with a bulldozer (fig. 11.3). This method removed nearly intact root sys¬ tems. Removing infected pine trees and stumps during thinning in this forest has also effectively reduced dis¬ ease losses (Roth and others 1977, Roth and Rolph 1978). Special guidelines (fig. 11.4) are used to mark symptomatic trees before pushing them over (Roth and others 1977,1980). Extraction of stumps up to 50 cm in diameter with a vibratory stump puller has also been successful in this area (Arnold 1981). In New Zealand, Armillaria root disease increased the cost of growing first-rotation radiata pine on sites with a high disease hazard by approximately 40% (Shaw and Calderon 1977). This cost approximates the maxi- GURE 113 — Pushing trees over with a bulldozer, rather cutting them off with a saw, is an effective way to )e root systems and remove inoculum on certain soil i he technique may also be used to create a "root free may serve as a barrier to further disease spread (see he 11.4). (C. Shaw) mum amount available for disease control. Initial stump uprooting and removal trials (fig. 11.1) sug¬ gested that the high disease losses encountered on nontreated sites could be reduced at a cost that was within the calculated economic limits (Shaw and Calderon 1977, Shaw unpubl., van der Pas 1981b). Uti¬ lizing stumps and roots may partially offset removal costs (Arnold 1981, Hakkila 1974). In addition, stump removal and plowing can benefit general seedling per¬ formance (Department of Forestry Queensland 1972, Morrison and others 1988, Roth and others 1977, Sorochkin 1972). The value of plants in urban forests, gardens, and or¬ chards often justifies the much higher treatment costs that are associated with inoculum removal. Recom¬ mended site preparation for orchards in the Australian Granitebelt consists of ripping to 35 cm so large roots can be removed, then ripping again to 25 cm to remove roots as fine as 1 cm (0.4-inch) diameter, followed by hand removing remaining roots. Without such rigorous site preparation, A. luteobubalina kills up to 50% of the newly planted fruit trees (Heaton and Dullahide 1989b). In New Zealand, soil has been extensively sifted to remove even the smallest infected root pieces (fig. 11.5A) to rehabilitate highly susceptible kiwifruit orchards that have suffered severely from Armillaria (Horner 1988). In urban settings, Pawsey (1973) recom¬ mended removing stumps and roots to whatever de¬ gree is practicable. Stump and root removal is most effective in the follow¬ ing situations. First, removal works well where the pathogen causes a primary disease from existing inocu¬ lum but does not continue to spread from secondary inoculum. Second, if secondary inoculum is important in the disease process, then it occurs in distinct patches (not diffuse in stands) and thus makes careful removal over concentrated areas possible. Third, where second¬ ary inoculum is important but the crop can be man¬ aged on a short rotation, such as intensively managed fruit orchards, the pathogen has little time for second¬ ary spread from unremoved inoculum pieces. Stump removal followed by a brief fallow period may increase the effectiveness of inoculum reduction where secondary inoculum is of concern. The presumably small, residual roots would likely decay quickly—par¬ ticularly in tropical regions. Pits dug for removing in¬ fected coffee roots in Kenya either are left open for several months or are treated with a soil sterilant be¬ fore covering. Still, a short fallow of 3-4 months after treatment is recommended (Baker 1972). Extended fal¬ low periods without stump removal might also prove effective in temperate regions although costly in terms of lost production time (Roth and others 1977). Rishbeth (1972b) showed that hardwood stumps cut 40 Reducing Losses O oo o O ° o Uon Diameter ■ 9 O o oo infectorf ttgivtp 5 10 yeot. old □ O e o o o o O O oo O O o h#oItky □ o old ttgmp o o o m o or o o o„ ° o o o o O o O u; O o o O o O O O o o O o °o° o o o o □ o..o 14 POORLY DEFINED INFECTIONS Q Mark according »o fobutolad dutoncoi o* ° O 9 o o o o o ° I root in * ♦ O * i. • O • a O ° * ° • T* o o Moon d iomoto r • 5 CP o o o, II * O o o • f □ Moan Diameter ■ 11 o •o 0 o° o o o § ° oo DISCRETE ACTIVE POCKET □ Developed around largo infected »tump» Q moro thon 1 0 ^oa r» old 23 o o o o o recenl O _ k,ll • 0 • □ dead O IS f«p.. • O «D II IB O o o o o o v o o o o □ O o o o o - is • o 0 .. ° O o SCATTERED MORTAUTT No t roa Imont roqui rod 0 o oO O O SAEE DISTANCES Ave f ago Active Old IIS*) Intervening OIH killing mortality ItOOl removed (old mortality s' is 10 I II 73 I 3 I I .0 14 7S loot O Healthy treat □ Healthy tree removed b.cog.e ol vno.oidoble logging domog. □ Old-growth tlumpi o O Dead or recogniiobly dneoted treat • Treat removed b.coo.e the, p-obobly o,e ml..ted bo. .how 0 n q abovo ground tymptomt FIGURE 11.4 — Guidelines developed by Roth and others (1977) to treat Armillaria root disease in ponderosa pine in south-central Washington. Trees at the left of the Armillaria center are too small for commercial thinning and are not disturbed except for removals to establish a barrier at the limits years earlier produced rhizomorphs, but at much lower levels than from younger stumps. The disease is rarely troublesome in first-rotation forests established on natural grassland and former agricultural land. How¬ ever, the economics of extending the rotation by delay¬ ing establishment may prohibit leaving land unstocked for lengthy periods—unless the land can otherwise be profitably used (e.g., livestock grazing or herbaceous cropping) in the interval. Annual cropping with cereals or alfalfa 4-5 years following clearing of orchard sites may deplete most of the nutritional sources for Armillaria (Guillamin 1977, Mallet and others 1985) while providing income. Simply avoiding old planting sites when replanting sections of orchards killed by Armillaria is advised for many crops (Horner 1990, Baker 1972, Heaton and Dullahide 1989b). This, combined with stump and root removal, provides an effect similar to fallow before root closure occurs. Heaton and Dullahide (1989b) rec¬ of the disease pockets. This action is done in conjunction with treatment of the commercial stand at the right by pushing trees over to remove their root systems (see fig. 11.3). (L.F. Roth, Journal of Forestry) ommended avoiding specific locations from which in¬ fected trees have recently been removed in orchards by relocating new orchard rows midway between the former rows. Trenching at least 1.1 m deep to isolate infected plants from healthy parts of a vinyard or fruit orchard is used in France to control spread of Armillaria root disease (Guillamin 1977). Trenching 0.6 m (20 inches) in cocoa plantations (Rishbeth 1980) and 1.1 m (3.5 feet) deep in coffee plantations (Wallace 1935) has been used suc¬ cessfully in Africa. Laying a plastic barrier in a trench and then backfilling it with removed soil has also been used for disease control in kiwifruit orchards in New Zealand (fig. 11.5B). Pruning and girdling diseased roots (Kendall 1931), or drying and aerating the root collar (Munnecke and oth¬ ers 1976, Kendall 1931), may be useful on individual, high-value trees in orchards, parks, gardens, forest Reducing Losses 165 B FIGURE 11.5 — Control of Armillaria root disease in kiwlfruit orchards in New Zealand (see fig. 9.6). A: Sifting of soil to remove even small pieces of infected roots to rehabilitate a severely diseased orchard. This method is no longer cost effective due to a reduction in kiwifruit profitibility. (I.J. Horner) B: If properly placed, trenches lined with plastic and backfilled serve as a mechanical barrier to root and rhizomorph growth into yet-unaffected portions of the orchard. (R.A. Hill) campsites, and seed orchards (Shaw and Roth 1978, i 980). Levitt (1947) used a water jet to expose root col- s of infected citrus trees. Rackham and others (1966) d that exposing root collars effectively con- rmillaria in citrus orchards, but they also noted that the large craters formed by this control method over a number of years could pose a hazard to workers in the orchards. Sokolov (1964) reported control of Armillaria in young Siberian larch by exposing the root collar. Generally, these procedures would be inappro¬ priate in commercial forests because of prohibitive costs. Chemical Protectants, Eradicants, and Curatives Apart from soil fumigation with carbon disulphide, methyl bromide, or chloropicrin after removing woody debris, very little experimental evidence supports the effectiveness of the most commonly advocated chemi¬ cal treatments (Shaw and Roth 1978). Justification for using many of these treatments is based on superficial and subjective criteria (Pawsey and Rahman 1976a). Reviews of chemical control of Armillaria have been presented by Pawsey and Rahman (1976a), Shaw and Roth (1978, 1980), and Thies and Russell (1984). As em¬ phasized by Shaw and Roth (1978 1980), managers must understand whether chemical applications are in¬ tended to protect uninfected plants, eradicate the fun¬ gus in infected stumps and roots, or treat or cure infected, living plants. Chemical soil fumigants that destroy Armillaria in root fragments are especially useful in orchard, vineyard, and floriculture operations where agricultural methods are applicable (Kissler and others 1973). Methvl bro¬ mide, a chemical demonstrated useful for this purpose in 1935 (Richardson and Johnson 1935), is still the most extensively used fumigant because of its non-specific action and good penetrability in soil (Vanachter 1979). Activity of chloropicrin against Armillaria in prune root sections was demonstrated in 1936 (Godfrey 1936). It is still a much-used fumigant because it will destroy even the most resistant soil pathogens, although penetration in soil is difficult to achieve. Fruit crops in California have benefited from using carbon disulphide injected at regular intervals over an infected site after removing stumps (Bliss 1951). Heaton and Dullahide (1989b) also recommended using methyl bromide fumigation in or¬ chards by injection into root-free soil and sealing with plastic. Systemic fungicides which have effectively suppressed A. ostoyae, A. mellea, and A. gallica in vitro are hexaconazole, flutriafol, and fenpropidin (Turner and Fox 1988). Chemicals of the ergosterol biosynthesis in¬ hibitor type are promising candidates for protectants and curatives (Schwabe and others 1984). In fact, sys¬ temic fungicides, "which can act both directly on the fungus in the soil and within the plants at some dis¬ tance from the point of application, have raised great hopes for the control of soil-borne fungal diseases" Reducing Losses (Louvet 1979). However, as Louvet (1979) also points out, substantial problems need to be solved before ap¬ plication of systemic chemicals is successful. First, trans¬ locating systemics in plants is acropetal whereas basipetal translocation would be more useful in treating roots. Second, strains of pathogens resistant to their ac¬ tion may rapidly appear in crops although evidence for such action in Armillaria is currently lacking. Armillatox, a phenolic emulsion containing 48% active ingredients (unidentified), has been marketed for spe¬ cific use against Armillaria. Apparently, the compound was developed after successfully controlling Armillaria with creosote (Bray 1970). Pawsey (1973), however, con¬ sidered creosote to be phytotoxic, and of doubtful value. Penetration of the material into the wood is minimal. Armillatox did produce some phytotoxic effects at the recommended dilution, even though rhizomorph pro¬ duction was somewhat reduced (Redfern 1971, Pawsey and Rahman 1976b). There was no evidence of remedial effect of Armillatox on established root infections. Pawsey and Rahman (1974) suggest that repeated, regu¬ lar use of Armillatox might protect against rhizomorph- initiated infection. Redfern (1971) found no beneficial effect from the chemical. Maneb (Pawsey and Rahman 1976a) and boric acid (Hesko 1971, Pawsey and Rahman 1976a) applied to tree root collars and stumps have successfully reduced some rhizomorph production, although Shaw (unpubl.) aban¬ doned trials with boric acid because of severe phytotox¬ icity. Rykowski (1974b) suggested that field applications of sodium pentachlorophenate (NaPCP) protected young Scots pine from Armillaria infection, helped eradicate the fungus in infected stump roots, and did not injure the tree. However, Shaw and others (1980) found NaPCP did not reduce infection on radiata pine inoculated with Armillaria, but they did notice some de¬ creases in host vigor. The long-term benefit of such treatments has yet to be demonstrated, and considerable doubt remains about the phytotoxic effects of both boric acid and NaPCP (Shaw and Roth 1978), and about NaPCP's potential effects on human health (Shaw and others 1980). Filip and Roth (1987) applied chemical to the root col¬ lars of small-diameter ponderosa pines to prevent mor¬ tality caused by A. ostoyae in south-central Washington. After 10 years, none of the seven chemicals (benomyl, captan, copper sulfate, iron sulfate, copper wire, vorlex, or chloropicrin) appeared to reduce mortality. Although single applications of the chemicals to protect pines from lethal infections were not effective, some of the chemicals may protect pines in high-value areas, such as seed orchards, recreation sites, or ornamental plantings, where economics may justify repeated applications. Fedorov and Bobko (1989) tested several fungicides for controlling existing infections in live hosts by applying them to the rhizosphere. They reduced rhizomorph production using cuprozan, fundazol, derozal, topsin- M, and copper oxychloride, but doubted the overall benefit of the treatments because Armillaria remained alive in host tissues. Recently, treating stone fruit trees in Australian orchards with potassium phosphite (fig. 11.6) has shown promising results; 75% of the treated trees appear to be recovering from Armillaria infection (Heaton and Dullahide 1989a). In many chemical tests, effects on rhizomorph produc¬ tion have been the main criterion for effective treat¬ ment. However, Redfern (1975) reported a significant negative correlation between the percentage of trees killed by Armillaria isolates and dry weight of rhizomorphs produced by the isolate. Rishbeth (1985a) also reported greater rhizomorph production by weakly parasitic A. gallica compared to the more ag¬ gressive A. mellea and A. ostoyae. Conceivably, treat¬ ment could alter the stump or rhizosphere environment such that the resident Armillaria species change, which may result in a difference in rhizomorph abundance. Considering our current understanding of rhizomorph production among different species in situ, this criterion for evaluating treatment effectiveness should be reconsidered (see chapter 4). Filip and Roth (1977) successfully controlled Armillaria in ponderosa pine stumps using methyl bromide, Vorlex, chloropicrin, carbon disulphide, and Vapam (fig. 11.7). Chloropicrin, Vortex, and methyl bromide eliminated the fungus from the stumps. In high-value crops and ornamentals where stump removal may not FIGURE 11.6 — Injection of a peach tree with potassium phosphite as treatment for prior infection by Armillaria root disease. Development of epicormic branches along the stem indicates success. Infected trees can apparently recover following such treatments, but resumption of full production remains to be shown. (J.B. Heaton) Reducing Losses 167 FIGURE 11.7 — Chemical treatment of ponderosa pine stumps to eradicate Armillaria. Holes are drilled in stumps (A) as entry ports for liquid eradicant chemicals (B) or gaseous fumigants (C). Many such treatments successfully eliminated Armillaria, but costs were considered too high for general applications in forestry. (Filip 1976, Filip and Roth 1977). (G. Filip) be desirable or possible (such as where access with heavy machinery needed for removal is limited), fumi¬ gants may be a useful option. Fumigant injections to es¬ tablish barriers which prevent vegetative spread of Armillaria may also be valuable (Houston 1975, Filip and Roth 1977) in forestry applications. The significant economic losses caused by Armillaria justify further efforts in chemical control (Pawsey and Rahman 1976a). In certain situations, chemical treat¬ ments may alter disease development at the epiphy- totic level (Filip and Roth 1977). However, certain aspects of such work need to be stressed. For example, field studies must define treatments by specific objec¬ tives—i.e., protecting, eradicating, or curing. Disease condition prior to treatment (i.e., proportion of stump colonized) must be known. Techniques for assessing ef¬ fectiveness must be both valid and definitive. The cost/ benefit of treatment must be evaluated in context with alternative measures and crop value. Detrimental ef¬ fects of the treatment on the environment or society re¬ quire consideration. One potential advantage for chemical protectants is that the critical region for appli¬ cation is likely to be the root collar, thus limiting the requiring treatment. For seedlings, a protectant mical should be relatively inexpensive, safe, easy to k\ easy to apply at planting, nonphytotoxic, or fungistatic, and persistent in the region of application. As discussed below, the control achieved by some chemicals may be interrelated with their ef¬ fects on other microorganisms. Biological Control and its Integration with Other Methods Biological control of a plant pathogen has several inherent advantages (Hunt and others 1971). Among others, it is more likely to be accepted by the public than either chemical control or the expense and initial unsightliness of stump and root removal. To control Armillaria, a rhizosphere or wood-inhabiting organism might function by inhibiting or preventing rhizomorph and mycelial development, by limiting the pathogen to substrate already occupied, by actively preempting the substrate, or by eliminating Armillaria (perhaps through replacement) from substrate already occupied. Pursuing these potential benefits must, however, be tempered with the feasibility of the technique (Shaw and Roth 1978,1980). Rishbeth (1976) noted two important features that make control of Armillaria by introduced organisms difficult. First, Armillaria has a positional advantage over introduced fungi since it already may occupy a portion of the substrate. Second, although Armillaria does not colonize wood quickly, it spreads rapidly in Reducing Losses the cambial zone of freshly killed trees. He suggested that antagonistic organisms might not be able to pre¬ vent Armillaria from becoming established in stumps, but they may restrict further stump colonization and thus limit the available food base. The same logic has been used to suggest that Armillaria species of limited pathogenicity may serve as biological control agents for Heterobasidion annosum (Fr.) Bref. (Morrison and Johnson 1978; Shaw 1989b,c). Perhaps the most thoroughly studied antagonists of Armillaria are Trichoderma species (fig. 11.8) from which two fungitoxic substances, trichodermin and an uni¬ dentified compound, have been isolated (Ishikawa and others 1976). Aytoun (1953) studied in vitro interac¬ tions of Trichoderma and Armillaria and concluded that Trichoderma must be considered a possible controlling factor in the spread of pathogenic fungi. Sokolov (1964) found fungi in six genera, including Trichoderma, Peni- cillium, and Peniophora, antagonized Armillaria. He rec¬ ommended using T. viride Pers.:Fr. as a control for Armillaria root disease. Dubos and others (1978) found that the medium in which T. viride inoculum was grown altered the degree to which the antagonist in¬ hibited rhizomorph production by Armillaria. Morquer and Touvet (1972a) suggested growing T. viride for Armillaria control on a "lactoserum" medium. Trichoderma species are common and ubiquitous soil in¬ habitants (Aytoun 1953, Griffin 1972), which might suggest that applying Trichoderma inoculum is gener¬ ally unnecessary. Trichoderma has been implicated in Armillaria control using sublethal doses of fumigants (Bliss 1951, Ohr and others 1973, Filip and Roth 1977), FIGURE 11.8 — Test of antagonism between A. luteobubalina and Trichoderma sp. Karri wood blocks previously colonized by 4. luteobubalina (left) and Trichoderma sp. (right) were placed face-to-face in soil for 6 weeks, separated and split, and isolations made at 1-cm intervals back from the contacting faces of each block. Green pins indicate recovery of Trichoderma sp., red pins 4. luteobubalina. Penetration of Trichoderma sp. into the wood block previously colonized by 4, luteobubalina was apparently stalled by the zone line. (E. Nelson) sublethal heating or drying treatments (Rackham and others 1966, Munnecke and others 1976), and possibly fire (Reaves and others 1990). Scytalidium lignicola Pesante also produces a toxin with antifungal proper¬ ties toward Armillaria (Cusson and LaChance 1974). Armillaria growth in culture is halted by either Scytalidium or its toxin, scytalidin. Since both Trichoderma and Scytalidium are common in soil, the basis for improving their ability to control Armillaria lies in shifting the balance among the fungal popula¬ tions. An inability to maintain effective populations of organisms antagonistic to Armillaria under field condi¬ tions has been the main factor limiting successful bio¬ logical control (Shaw and Roth 1978,1980). Bliss (1951) demonstrated the ability of T. viride to re¬ place Armillaria in artificially infected root segments fu¬ migated with carbon disulphide (CS.,). Garrett (1957, 1958) showed that CS, can directly damage Armillaria mycelium; pure cultures of T. viride, in the absence of fumigation, also killed Armillaria. Apparently, both di¬ rect fumigant toxicity and subsequent action of T. viride were killing Armillaria in fumigated soils. After fumiga¬ tion (fig 11.70, Filip and Roth (1977) frequently iso¬ lated T. viride from pine stumps in which Armillaria was no longer viable. Munnecke and others (1973) sug¬ gested that after fumigation with CS 2 or methyl bro¬ mide, a lag period for Armillaria growth occurred, indicating a "weakening" of the Armillaria. Trichoderma viride, being more tolerant of the chemical (Ohr and others 1973), was able to exploit the lag period and ex¬ ert an antagonistic action on Armillaria. Riffle (1973) noted that two mycophagous nematodes greatly reduced mortality of ponderosa pine seedlings inoculated with Armillaria. The nematodes apparently reduced fungal vigor and growth. In vitro studies of how mycophagous nematodes affect mycelia of Armillaria and Trichoderma species indicated a possible role of Aphelenchus avenae in controlling Armillaria in a French vineyard. The nematode destroyed the hyphae of Armillaria but grew well on T. polysporum (Link ex Pers.) Rifai without reducing its growth (Cayrol and others 1978). Armillaria produces antibiotic compounds (see chapter 3). Oduro and others (1976) suggested that such activ¬ ity may be an important factor in surviving attack by antagonistic soil microorganisms. Significantly, Ohr and Munnecke (1974) showed that sublethal methyl bromide fumigation prevented the production of anti¬ biotics by Armillaria. Munnecke and others (1976) sug¬ gested that heating or drying may similarly affect Armillaria. The critical factor is that Armillaria is stressed. The factors causing the stress may concur¬ rently stimulate antagonistic organisms, resulting in further damage to the already weakened Armillaria. Reducing Losses 169 Direct competition for the woody substrate may be an important natural control of Armillaria. Garrett (1956b) hypothesized that root-inhabiting parasites would have a low competitive saprophytic ability (see chapter 4). Redfern (1968) suggested that Armillarin probably cannot survive indefinitely as a saprophyte and that control is perhaps most easily achieved in the sapro¬ phytic phase. Leach (1937) observed that Rhizoctonia lamellifera Small prevented Armillarin from colonizing tea roots. Sokolov (1964) also observed that spruce stumps colonized by Lenzites saepiaria Fr. and Peniophora gigantea (Fr.) Massee were not invaded by Armillarin. From laboratory tests, Orlos (1957) thought Fames pinicola (Swartz:Fr.) Cke. might be useful in con¬ trolling Armillarin because of its greater growth rate and ability to exclude Armillarin from occupied media. Fedorov and Bobko (1989) tested 10 basidiomycetes which were capable of excluding Armillarin from occu¬ pied substrates. Two of these, Peniophora gigantea (Fr.) Massee and Pleurotus ostreatus (Jacq.:Fr.) P. Kumm., also effectively prevented Armillaria growth in freshly cut stumps into which they had been inoculated. Coriolus versicolor (L.:Fr.) Quel, Stereum hirsutum (Willd.:Fr.) S.F. Gray, and Xylaria hypoxylon (L.:Fr.) Grev. inoculated into karri thinning stumps simulta¬ neously with A. luteobubalina (fig 11.9) each signifi¬ cantly reduced colonization by Armillaria (Pearce and Malajczuk 1990b). The eucalypt stumps were colonized both above and below ground by the competing fungi, but they were more effective antagonists above ground. A naturally occurring, cord-forming species of Hypholoma proved to be even more competitive with Armillaria, in some cases excluding it entirely. Such cord-forming, wood-decay fungi have a similar niche to Armillaria and are perhaps the most exciting recent discovery in relation to its possible biological control. They are capable of subcortical mycelial growth in stumps and occupy the same initial sites as RE 11.9 — Successful establishment of antagonistic fungi arri stumps inoculated with A. luteobubalina. Stump ' on with either Coriolus versicolor (A) or a Hypholoma '■uhcantly reduced colonization by A. luteobubalina. •■ -’ting fungi colonized the eucalypt stumps both above B and below ground, but were more effective competitors above ground. A naturally occurring, cord-forming species of Hypholoma proved to be even more competitive with Armillaria, in some cases excluding it entirely (Pearce and Malajczuk 1990b). (E. Nelson) Reducing Losses Armillaria (Redfern 1968). According to Rayner (1977), cord formers “closely paralleled A. mellea in their be¬ havior, except for their lack of pathogenicity." Further studies have indicated that several species of cord¬ forming basidiomycetes, in particular Phanerochaete velutina (DC per Pers.:Fr.) Karst., Hypholoma fasciculare (Huds.:Fr.) Kumm, and Steccherinum fimbriatum (Pers.:Fr.) J. Erikss., have considerable potential to spread and colonize woody debris in field sites. Several produce networks of mycelial cords in soil and litter (Dowson and others 1988a) which can infest additional woody substrates (Dowson and others 1988b). Popula¬ tions of some cord formers can be manipulated by chemically treating stumps (Rayner 1977), a finding that indicates potential to artificially induce biological control of Armillaria. Ammonium sulphamate appears particularly useful as it increased colonization and de¬ cay by cord-formers of below-ground portions of treated beech and birch stumps (Rayner 1977). Stump fumigation has excluded or eradicated Armillaria directly in lethal doses (Bliss 1951, Rackham and others 1966, Filip and Roth 1977). Sublethal doses, however, do not kill Armillaria directly, but allow com¬ peting fungi less affected by the chemical to replace it. (Munnecke and others 1973, Ohr and others 1973). Sub- lethal doses of methyl bromide injected into orchard soil in California successfully controlled Armillaria (Munnecke and others 1981). Trichoderma spp., which resisted the methyl bromide, may have been respon¬ sible for controlling Armillaria in the fumigated soil. Formaldehyde was used in sublethal doses to control Armillaria in apple and pear orchards in China (Chang and others 1983), where Trichoderma populations were stimulated by the treatment and were credited with the control. Silvicides which can rapidly kill host tissues are used to kill trees before cutting in tropical regions where the killed roots decay rapidly and pathogens are readily re¬ placed by saprophytes (Mallet and others 1985). How¬ ever, in temperate regions, the rapid killing by herbicides may benefit Armillaria. For example, Rishbeth (1976) reported that stump treatment with 2,4,5-T favored Armillaria colonization; and Pronos and Patton (1979) found more rhizomorph production 10 years after treatment in stumps of herbicide-killed oaks than in girdled oaks. Rapid death from the herbicide treatment was thought to have favored Armillaria over competing saprophytes. While stump colonization is an important factor in Armillaria root disease, the nutritional quality of stumps and roots will influence the longevity of the pathogen. Leach's report (1937) that ring-barking effec¬ tively controlled the spread of Armillaria in African tea plantations led to several investigations into the nutri¬ tional suitability of altered stumps for Armillaria. Leach (1939) suggested that decreased concentrations of stored carbohydrates within the roots of ring-barked trees rendered them unsuitable for Armillaria coloniza¬ tion. Ring-barking also could have rendered the roots more easily colonized by other, saprophytic fungi and thus quickly reduced the volume of material available for Armillaria. Redfern (1968) found that ring-barking or poisoning mature oaks 1 year prior to felling in Britain resulted in more rapid decay of the roots by Armillaria compared to those felled without prior treatment. Rhizomorph production may have been a major influence in Redfern's study. While rhizomorphs may be scarce in African soils (Wiehe 1952), they are abundant and pro¬ liferate epiphytically on live tree roots in Britain where localized lesions on live roots are also common. Ring¬ barking and silvicide treatment favored the invasion of the already present parasite. The roots of the treated oaks either initially were not a good substrate, or dete¬ riorated quickly because significantly fewer rhizomorphs were produced from sections of treated roots compared to roots of non-treated trees 5 years af¬ ter felling. Neither ring-barking nor silvicide treatment was effective in reducing mortality in subsequent plan¬ tations. Lanier (1971) indicated that girdling old Scots pine and common beech a year before felling reduced the num¬ ber of young pines attacked. Disease incidence was low, however, even in untreated parts of the forest. Punter (1963) indicated that girdling reduced neither the mortality of young trees nor the number of Armillaria basidiomes on stumps. Swift (1970) con¬ cluded that ring-barking effectively prevented invasion of stumps from external inoculum sources, but spread of the fungus from pre-existing lesions was not inhib¬ ited and probably was enhanced. Heating and drying methods such as those employed by Birmingham and Stokes (1921) and Rackham and others (1966) are costly, difficult to apply, and almost certainly limited in utility to orchard and ornamental situations. Broadcast burning after clearfelling of indig¬ enous forest in New Zealand significantly reduced the number of viable rhizomorphs compared to counts be¬ fore clearfelling (Hood and Sandberg 1989). However, all the pine plantations described in New Zealand as having suffered severe losses from Armillaria (Shaw and Calderon 1977) were on sites burned during prepa¬ ration for planting. Apparently, such reductions in rhizomorphs are not sufficient to control the disease. Focan and others (1950) noticed that root disease in¬ creased among perennial plants established after burn¬ ing, and Trichoderma spp. markedly declined. However, Reaves and others (1990) reported that recovery of Reducing Losses 171 Trichoderma isolates from soil was unaffected by fire. The species composition shifted such that, after burn¬ ing, the most frequently isolated species had a greater antagonism towards Armillaria. Mycorrhizae have been suggested as a protection against parasitic attack by Armillaria. Gaudray (1973) postulated that the formation of mycorrhizae on exotic Sitka spruce in France is incomplete and thus affords inadequate protection against Armillaria. Studies in vitro have shown that mycorrhizal fungi can inhibit Armillaria (Eghbaltalab and others 1975). Direct protec¬ tion by mycorrhizae seems unlikely, however, as the main infection sites for Armillaria are on coarse roots rather than the fine roots where mycorrhizae develop. Examining natural populations of organisms that are antagonistic to the identified, parasitic species of Armillaria present on sites that express different severi¬ ties of root disease may be useful. Such examinations may indicate characteristics that could be manipulated through management so as to favor antagonistic organisms. Conclusions Whether to invest in Armillaria control and selecting control methods are decisions that need to be based on the value of losses in the absence of control. Assessing impact to commodity production or other features such as amenity value may, in itself, be costly, but it is an important precursor to any control decision. Both mon¬ etary and environmental costs of various control alter¬ natives should be justified by commodity or other values derived. Effectiveness of control in a particular situation is another important consideration. For ex¬ ample, stump removal that reduces inoculum by 60% may greatly improve the productivity of a crop for which primary inoculum is the major concern, or of a crop that is produced in a short time. But stump re¬ moval with this same level of inoculum reduction may only slightly improve the productivity of a crop subject to secondary disease spread. Control projects must be monitored long-term in most crops to measure relative gains from treatment (Jancarik 1955). Fruit orchards that are subject to sec¬ ondary spread of Armillaria may require monitoring over decades to evaluate treatment fully. Forests may require a century or more to reach maturity. If disease spread from secondary inoculum is of concern, then onitoring for at least 20-30 years may be required to sess control effectiveness. How the effectiveness may o\ er into subsequent rotations also needs to be ered because little information is available on this subject (see chapter 10). In such cases, interim evaluation and some degree of faith in projections are necessary. In summary, the following checklist needs to be con¬ sidered before any attempts are made to control Armillaria root disease. (1) Critically evaluate disease impact to ensure that the level of loss justifies control. The use of disease models may aid this effort (see chapter 10). (2) Control through cultural modifications should be given first priority, particularly in forests. As our current forest management rarely emulates nature's processes, pathologists must work in direct coopera¬ tion with foresters to understand and modify dis¬ ease-stimulating practices. (3) Utilize resistant or tolerant species, genotypes, or rootstocks, if known, that are compatible with other necessary values. Ensure that the host genotype se¬ lected for resistance is suitable for planting on po¬ tential sites, and will provide for the planned end use of the fruit or fiber. Pursue opportunities to ge¬ netically engineer Armillaria- resistant or tolerant species. (4) When establishing new plantations or orchards, exercise care in site selection. Small-scale trials to evaluate disease potential should be established prior to large-scale land clearing and plantation or orchard development. If the site is found to have a high disease hazard, then one must be prepared for costly preestablishment actions such as inoculum re¬ movals by more thorough site preparation, post¬ ponement of plantation or orchard establishment for some unknown period, or elimination of the site from further consideration. (5) Maintain the general health of the forest, orchard, or amenity planting by preventing damage from other agents, avoiding adverse sites, and discourag¬ ing detrimental human activities. (6) Direct reductions of inoculum levels by physical removal of stumps and roots requires careful eco¬ nomic and ecological analysis. The effectiveness of such treatments in orchards, exotic plantations, and amenity plantings is generally appreciated; their ef¬ fectiveness in natural forests will be better under¬ stood within the next few years when results from existing, long-term trials become available. (7) When considering chemical treatments, clearly differentiate among protectants, eradicants, and 7 71 Reducing Losses curatives. Except for high-value fruit or amenity trees, curatives are likely to be uneconomical. Even in orchards and amenity plantings, chemical appli¬ cations need to be realistically evaluated for their relative cost/benefit. For chemical treatment of stumps, consider compounds that can be translo¬ cated, particularly basipetally. Protectants should be inexpensive, easy to handle and apply, nonphytotoxic, fungitoxic or fungistatic, and rela¬ tively persistent. Possible environmental and human health hazards require consideration. (8) Fumigation, girdling, and silvicide treatment be¬ fore felling may be useful methods to employ in pre¬ paring land for orchards, ornamentals, and some forestry applications such as seed orchards and test plantations. Fallowing after such treatment may im¬ prove effectiveness, especially where disease spread from secondary inoculum is anticipated. (9) Biological control is desirable but requires further development for practical application in most situations. Research on antagonists, particularly cord-formers, needs to continue as does work on the various actions (i.e., fire, chemicals) that might be used to alter conditions in a way that favors devel¬ oping and maintaining populations of desirable, an¬ tagonistic organisms. Armillaria control needs to be a thoughtful, reasoned process, not a random or haphazard one. Evaluating the necessity for control and, if found necessary, deter¬ mining the best option to implement are integral to prudent stewardship of forests, orchards, and amenity plantings. Our increased understanding of species identity, their pathogenic behaviors, and ecological re¬ lationships offers the opportunity for a systematic evaluation of approaches to controlling Armillaria root disease. 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Degradation of raspberry suberin by Fusarium solani f. sp. pisi and Armillaria mellea. Phytopathologische Zeitschrift. 110: 192-199. Zollfrank, U.; Hock; B. 1987. Infection of Norway spruce by Armillaria under controlled conditions. European Journal of Forest Pathologv. 17: 266-270. Zondag, R.; Gilmour, J.W. 1963. Forest pathology. In: Report of the New Zealand Forest Research Institute, 1962: 39-51. Zycha, H. 1970. Hallimasch [Armillaria mellea (Vahl ex Fr.) Kumm.] als Kernfaule- Erreger an Fichte ( Picea abies Karst.). Fortwissenschaftliches Centralblatt. 89: 129-135. Literature Cited appendix Scientific and Common Names of Plants Noted in This Book Common Names — Scientific Names COMMON NAMES SCIENTIFIC NAMES acacia Acacia African oilpalm Elaeis guineensis Agathis Agathis ailanthus Ailanthus albizia Albizzia falcata Merr. albizia Albizzia alder Alnus almond Primus amygdalus Batsch. alpine ash Eucalyptus delegatensis R.T. Bak. American chestnut Castanea dentata (Marsh.) Borkh. American beech Fagus grandifolia Ehrh. apple, pome fruit Malus apricot Primus armeniaca L. araucaria Araucaria Arizona pine Pinus arizonica Engelm. Arizona pine Finns ponderosa var. arizonica (Engelm.) Shaw ash Fraxinus avocado Persea Bahaman pine Pinus caribaea var. bahamensis Barr, ex Golf. balsam poplar Populus balsamifera L. balsam fir Abies balsamea (L.) Miller banana Musa banksia Banksia barkclothtree Brachystegia beech Fagus beefwood Casuarina beet Beta bigtooth aspen Populus grandidentata Michx. birch Betula black currant Ribes nigrum L. black oak Quercus velutina Lamarck black spruce Picea mariana (Mill.) B.S.P. blackberry, raspberry broad-leaved Rubus peppermint Eucalyptus dives Schau. broom Cytisus COMMON NAMES SCIENTIFIC NAMES brown barrel Eucalyptus fastigata Deane & Maid. brown salwood Acacia mangium Willd. cacao Theobroma cacao L. cactus Opuntia cane Arundinaria Caribbean pine Pinus caribaea Morelet carrot Daucus cassava Manihot ceanothus Ceanothus cedar Cedrus cedrela Cedrela chaulmoogratree Hydnocarpus cherry Primus chestnut Castanea chocolatetree Theobroma cinchona Cinchona citrus Citrus cocoa Theobroma coconut Cocos coffee Coffea colanut Cola common ash Fraxinus excelsior L. common fig Ficus carica L. common teak common Tectona grandis L. f. chaulmoogratree Hydnocarpus anthelminticus Pierre common beech Fagus sylvatica L. common pistachio Pistacia vers L. common pomegranite Punica granatum L. common tea Camellia sinensis (L.) Ktze. cork oak Quercus suber L. Corsican pine Pinus nigra var. maritima (Ait.) Melville cotton Gossypium cryptomeria Cryptomeria currant, gooseberry Ribes cypress Cupressus cypress pine Widdringtonia cypress pine Callitris dacrydium Dacrydium Armillaria Root Disease 221 COMMON NAMES SCIENTIFIC NAMES COMMON NAMES SCIENTIFIC NAMES dawn redwood Metasequoia Japanese larch Larix leptolepis (Sieb. & Zucc.) deodar Cedrus deodora G. Don ex Loud. Gord. Douglas-fir Pseudotsuga menziesii (Mirb.) Franco jarrah Eucalyptus marginata Donn ex Smith Douglas-fir Pseudotsuga jujube Zizyphus downy oak Quercus pubescens Willd. karri Eucalyptus diversicolor F. eastern white pine Pinus strobus L. Muell. eastern hemlock Tsuga canadensis (L.) Carr. kauri Agathis australis Salisb. elm Ulmus Khasi pine Pinus kesiva Boyle ex Gordon Engelmann spruce Picea engelmanni Parry ex (P. insularis Endl.) Engelm. khaya Khaya English oak Quercus robur L. (Q. kiwifruit Actinidia pendunculata Ehrh). Korean pine Pinus koraiensis Sieb. & Zucc. eucalypt-gum Eucalyptus larch Larix European larch Larix decidua Mill. lavender Lavandula falsecypress Chamaecyparis lead tree Leucaena fi g Ficus lime Citrus aurantifolia fir, true fir Abies (Christmann in L.) Swingle fish pelargonium Pelargonium hortorum Bailey litchi Litchi flooded gum Eucalyptus grandis Hill ex loblolly pine Pinus taeda L. Maid. locust Robinia geranium Pelargonium lodgepole pine Pinus contorta Dougl. ex Loud gliricidia Gliricidia loganberry Rubus loganobaccus L. Bailey gmelina Gmelina loquat Libotrya gmelina Gmelina arborea L. macadamia Macadamia gooseberry, current Ribes Mahaleb cherry Primus mahaleb L. granadilla Passiflora mahogany Sivietenia grand fir Abies grandis (Dougl. ex D. mango Man gif era Don) Lindl. maple Acer grape \/it is Maritime pine Pinus pinaster Ait. greatcone banksia Banksia grandis Willd. messmate stringybark Eucalyptus obliqua L'Herit. green wattle Acacia decurrens (Wendl.) Mexican weeping pine Pinus patula Schiede & Deppe Willd. Mexican cypress Cupressus lusitanica Mill. grevillea Grevillea mlanji cedar Widdringtonia whytei Rendle guava Psidium Morinda spruce Picea morinda Link Hankow willow Salix matsudana Koidz. mountain pine Pinus uncinata Mill, ex Mirb. hazelnut Corylus mountain ash Eucalyptus regnans F. Muell. hemlock Tsuga mountain gray gum Eucalyptus cypellocarpa L. hickory Cary a Johnson hinoki Chamaecyparis obtusa Endl. mountain hemlock Tsuga mertensiana (Bong.) Honduran pine Pinus caribaea var. Carr. hondurensis Barr. & Golf. mulberry Morns Honduras mahogany Sivietenia macrophylla King Muula Parinari mobola F. Muell. ex hops Humulus Benth. Hungarian oak Quercus frainetto Ten. myrtle-beech Nothofagus cunninghamii incense-cedar Calocedrus decurrens (Torr.) Florin (Libocedrus decurrens narrow-leaved (Hook, f.) Oerst. incense-cedar Torr.) peppermint Eucalyptus radiata Sieb. ex DC Calocedrus New Guinea gum Eucalyptus deglupta Blume Indian fig Opuntia ficus-indica Mill. nightshade Solanum Indian pipe Monotropa hypopitys L. northern California Indian pipe Monotropa walnut Juglans hindsii Jeps. ex Smith Indian pipe Monotropa uniflora L. Norway spruce Picea abies (L.) Karst. iron tree Metrosideros oak Quercus jack pine Pinus banksiana Lamb. ohia Metrosideros polymorpha Japanese redcedar Cryptomeria japonica (L.) D. Don (Gaug.) Rock Ill Armillaria Root Disease COMMON NAMES SCIENTIFIC NAMES oilpalm Elaeis olive Olea orchid Gastrodia cunninghamii Hook. f. orchid Gastrodia elata Bl. orchid Gastrodia orchid Galeola orchid Galeola septentrionalis Reichb. f. papaya Carica paper birch Betula papyrifera Marsh. paraserianthes Paraserianthes paraserianthes Paraserianthes falcataria (L.) I. Nielsen parsnip Pastinaca passion fruit Passiflora pawpaw Asimina peach Prunus persica Sieb. & Zucc. pear, pome fruit Pyrus pecan Garya illinoiensis (Wangenh.) K. Koch Persian walnut Juglans regia L. persimmon Diospyros pindrow fir Abies pindrow Royle pine Pinus pistachio Pistacia planetree Acer pseudoplatanus pomegranite Punica ponderosa pine Pinus ponderosa Dougl. ex Laws. poplar Populus potato Solanum tuberosum L. provence broom Cytisus purgans (L.) Boiss. quaking aspen Populus tremuloides Michx. Queensland kauri Agathis robusta F.M. Bailey radiata pine Pinus radiata D. Don red alder Alnus rubra Bong. red maple Acer rubrum L. red oak Quercus rubra L. red pine Pinus resinosa Ait. red spruce Picea rubens Sarg. rimu Dacrydium cupressinum Sol. ex Lambert rose Rosa rubber tree Hevea Sakhalin spruce Picea glehnii (Schmidt) Mast. sand pine Pinus clausa (Chapm.) Vasey scarlet oak Quercus coccinea Michx. Scots pine Pinus sylvestris L. senna Cassia sequoia Sequoiadendron Siberian larch Larix sibirica Ledeb. silver birch Betula verrucosa Ehrh. silver fir Abies alba Mill. silver-beech Nothofagus menziesii (Hook, f.) Oerst. COMMON NAMES SCIENTIFIC NAMES silver maple Acer saccharinum L. silver wattle Acacia dealbata Link Sitka spruce Picea sitchensis (Bong.) Carr. slash pine Pinus elliottii Engelm. snowbrush Ceanothus velutinus Dougl. sour cherry Prunus cerasus L. sour orange Citrus aurantium L. southern blue gum Eucalyptus globulus Labill. ssp. bicostata (Maid et al.) Kirkp. southern-beech Nothofagus spike barkclothtree Brachystegia spiciformis Benth. spruce stonefruits, apricot. Picea cherry, peach, plum Prunus strawberry Fragaria subalpine fir Abies lasiocarpa (Hook.) Nutt. Sudan colanut Cola acuminata (Pal.) Schott & Endl. sugar maple Acer saccharum Marsh. sugarcane Saccharum officinarum L. sunbush Bossiaea sunbush Bossiaea laidlawiana Tovey & Morris swamp mahogany Eucalyptus robusta Sm. sweet orange Citrus sinensis Osbeck sweetcane Saccharum sycamore Platanus occidentals L. sycamore Platanus tawa Beilschmiedia tawa (Cunn.) Kirk tawa Beilschmiedia tea Camellia teak Tectona terminalia Terminalia thuja Thuja tomato Lycopersicon toon Toona tung Aleurites tungoiltree Aleurites fordii Hemsley turkey oak Quercus cerris L. walnut Juglans wandoo Eucalyptus wandoo Blakely western hemlock Tsuga heterophylla (Rafn.) Sarg. western larch Larix occidentals Nutt. western redcedar Thuja plicata Donn ex D. Don western white pine Pinus monticola Dougl. ex D. Don white fir Abies concolor (Gord. & Glend.) Lindl. ex Hildebr. white mulberry Morns alba L. white oak Quercus alba L. white spruce Picea glauca (Moench) Voss willow Salix Armillaria Root Disease 223 Scientific Names — Common Names SCIENTIFIC NAMES COMMON NAMES Cary a hickory C. illinoiensis (Wangenh.) SCIENTIFIC NAMES COMMON NAMES K. Koch pecan Abies fir, true fir Cassia senna A. alba Mill. silver fir Castanea chestnut A. balsamea (L.) Miller balsam fir C. dentata (Marsh.) Borkh. American chestnut A. concolor Casuarina beefwood (Gord. & Glend.) Lindl. Ceanothus ceanothus ex Hildebr. white fir C. velutinus Dougl. snowbrush A. grandis (Dougl. ex D. Don) Cedrela cedrela Lindl. grand fir Cedrus cedar A. lasiocarpa (Hook.) Nutt. subalpine fir C. deodora G. Don ex Loud. deodar A. pindrow Royle pindrow fir Chamaecyparis fa lsecy press Acacia acacia C. obtusa Endl. hinoki A. dealbata Link silver wattle Cinchona cinchona A. decurrens (Wendl.) Willd. green wattle Citrus citrus A. mangium Willd. brown salwood C. aurantifolia (Christmann in Acer maple L.) Swingle lime A. pseudoplatanus planetree C. aurantium L. sour orange A. rubrum L. red maple C. sinensis Osbeck sweet orange A. saccharum Marsh. sugar maple Cocos coconut A. saccharinum L. silver maple Coffea coffee Ac tin id ia kiwifruit Cola colanut Agathis Agathis C. acuminata (Pal.) A. australis Salisb. Queensland kauri Schott & Endl. Sudan colanut A. robusta F. M. Bailey kauri Corylus hazelnut Ailanthus ailanthus Cryptomeria cryptomeria Albizzia albizia C. japonica (L.) D. Don Japanese redcedar A. falcata Merr. albizia Cupressus cypress Aleurites tung C. lusitanica Mill. Mexican cypress A. fordii Hemsley tungoiltree Cytisus broom Alnus alder C. purgans (L.) Boiss. provence broom A. rubra Bong. red alder Dacrydium dacrydium Araucaria araucaria D. cupressinum Sol. ex Arundinaria cane Lambert rimu Asimina pawpaw Daucus carota L. carrot Banksia banksia Diospyros persimmon B. grandis Willd. greatcone banksia Elaeis oilpalm Beilschmiedia tawa E. guineensis African oilpalm B. taum (Cunn.) Kirk tawa Eucalyptus eucalvpt-gum Beta beet E. cypellocarpa L. Johnson mountain grey gum Betula birch E. deglupta Blume New Guinea gum B. papyrifera Marsh. paper birch E. delegatensis R.T. Bak. alpine ash B. verrucosa Ehrh. silver birch E. diversicolor F. Muell. karri Bossiaea sunbush E. dives Schau. broad-leaved B. laidlawiana Tovey & Morris sunbush peppermint Brachystegia barkclothtree E. fastigata Deane & Maid. brown barrel B. spiciformis Benth. spike barkclothtree £. globulus Labill. ssp. bicostata CaUitris cypress pine (Maid et al.) Kirkp. southern blue gum Calocedrus incense-cedar E. grandis Hill ex Maid. flooded gum C. decurrens (Torr.) Florin E. marginata Donn ex Smith jarrah (Libocedrus decurrens Torr.) incense-cedar E. obliqua L'Herit. messmate Camellia tea stringvbark C. sinensis (L.) Ktze. common tea £. radiata Sieb. ex DC. narrow-leaved Carica papaya peppermint 224 Armillaria Root Disease SCIENTIFIC NAMES COMMON NAMES £. regnans F. Muell. mountain ash E. robusta Sm. swamp mahogany E. xvandoo Blakely wandoo Tagus F. grandifolia Ehrh. beech American beech F. sylvatica L. common beech Ficus fi g F. carica L. common fig Fragaria strawberry Fraxinus ash F. excelsior L. common ash Galeola orchid G. septentrionalis Reichb. f. orchid Gastrodia orchid G. elata Bl. orchid G. cunninghamii Hook. f. orchid Gliricidia gliricidia Gmelina gmelina G. arborea L. gmelina Gossypium cotton Grevillea grevillea Hevea rubber tree Humulus hops Hydnocarpus chaulmoogratree H. anthelminticus Pierre common Juglans /. regia L. J. hindsii Jeps. ex Smith chaulmoogratree walnut Persian walnut northern California walnut Khaya khaya Larix larch L. decidua Mill. E. leptolepis (Sieb. & Zucc.) European larch Japanese larch Gord. E. occidentalis Nutt. western larch E. sibirica Ledeb. Siberian larch Lavandula lavender Leucaena leadtree Libotrya loquat Lit chi litchi Lycopersicon tomato Macadamia macadamia Malus apple, pome fruit Mangifera mango Manihot cassava Metasequoia dawn redwood Metrosideras irontree M. polymorpha (Gaug.) Rock ohia Monotropa Indian pipe M. hypopitys L. Indian pipe M. uniflora L. Indian pipe Morns mulberry M. alba L. white mulberry Musa banana SCIENTIFIC NAMES COMMON NAMES Nothofagus southern-beech N. cunninghamii (Hook, f.) Oerst. myrtle-beech N. menziesii (Hook, f.) Oerst. silver-beech Olea olive Opuntia cactus O. ficus-indica Mill. Indian fig Paraserianthes paraserianthes P. falcataria (L.) I. Nielsen paraserianthes Parinarium Parinarium P. mobola F. Muell. ex Benth. Muula Passiflora passion fruit, granadilla Pastinaca parsnip Pelargonium geranium P. hortorum Bailey fish pelargonium Persea avocado Picea spruce P. abies (L.) Karst. Norway spruce P. engelmanni Parry ex Engelm. Engelmann spruce P. glauca (Moench) Voss white spruce P. glehnii (Schmidt) Mast. Sakhalin spruce P. mariana (Mill.) B.S.P. black spruce P. morinda Link Morinda spruce P. rubens Sarg. red spruce P. sitchensis (Bong.) Carr. Sitka spruce Pinus pine P. arizonica Engelm. Arizona pine P. banksiana Lamb. jack pine P. caribaea Morelet Caribbean pine P. caribaea var. bahamensis Barr, ex Golf. Bahaman pine P. caribaea var. hondurensis Barr. & Golf. Honduran pine P. clausa (Chapm.) Vasey sand pine P. contort a Dougl. ex Loud. lodgepole pine P. elliottii Engelm. slash pine P. kesiva Boyle ex Gordon (P. insularis Endl.) Khasi pine P. koraiensis Sieb. & Zucc. Korean pine P. monticola Dougl. ex D. Don western white pine P. nigra var. maritima (Ait.) Melville Corsican pine P. patula Schiede & Deppe Mexican weeping pine P. pinaster Ait. Maritime pine P. ponderosa Dougl. ex Laws. ponderosa pine P. ponderosa var. arizonica (Engelm.) Shaw Arizona pine P. radiata D. Don radiata pine P. resinosa Ait. red pine P. strobus L. eastern white pine P. sylvestris L. Scots pine P. taeda L. loblolly pine P. uncinata Mill, ex Mirb. mountain pine Armillaria Root Disease 225 SCIENTIFIC NAMES COMMON NAMES Pistacia pistachio P. vers L. common pistachio Platanus sycamore P. occidentalis L. sycamore Populus poplar P. balsamifera L. balsam poplar P. grandidentata Michx. bigtooth aspen P. tremuloides Michx. quaking aspen Primus stonefruits, apricot P. amygdalus Batsch. cherry, peach, plum almond P. armeniaca L. apricot P. cerasus L. sour cherry P. mahaleb L. Mahaleb cherry P. persica Sieb. & Zucc. peach Pseudotsuga Douglas-fir P. menziesii (Mirb.) Franco Douglas-fir Psidium guava Punica pomegranite P. granatum L. common Pyrus pomegranite pear, pome fruit Quercus oak Q. alba L. white oak Q. cerris L. turkey oak Q. coccinea Michx. scarlet oak Q. frainetto Ten. Hungarian oak Q. pubescens Willd. downy oak Q. robur L. (Q. pendunculata Ehrh). English oak Q. rubra L. red oak Q. suber L. cork oak Q. velutina Lamarck black oak Quinine cinchona Ribes currant, gooseberry R. nigrum L. black currant SCIENTIFIC NAMES COMMON NAMES Robinia locust Rosa rose Rubus blackberry, raspberry R. loganobaccus L. Bailey loganberry Saccharum sweetcane S. officinarum L. sugarcane Salix willow S. matsudana Koidz. Hankow willow Secjuoiadendron sequoia Solatium nightshade S. tuberosum L. potato Swietenia mahogany S. macrophylla King Honduras mahogany Tec torn teak T. grandis L. f. common teak Terminalia terminalia Theobroma chocolatetree T. cacao L. cacao Theobroma cocoa Thuja thuja T. plicata Donn ex D. Don western redcedar Toona toon Tsuga hemlock T. canadensis (L.) Carr. eastern hemlock T. heterophylla (Rafn.) Sarg. western hemlock T. mertensiana (Bong.) Carr. mountain hemlock Ulmus elm Vitis grape Widdringtonia cypress pine W. whytei Rendle mlanji cedar Zizyphus jujube 6 Armillaria Root Disease About the Authors Charles G. Shaw III, better known as Terry, is Princi¬ pal Research Plant Pathologist and Project Leader for the USDA Forest Service's research project on pest im¬ pact assessment technology. From 1977 to 1986, he was principal research plant pathologist at the Forest Service's Forestry Sciences Laboratory in Juneau, Alaska. He received a B.S. degree in forestry from Washingtoxr State University in 1970 and a Ph.D. in plant pathology from Oregon State University in 1974. His thesis dealt with Armillaria root disease in pon- derosa pine. He continued working on that disease from 1974 to 1977 while serving as a research scientist with the Forest Research Institute in Rotorua, New Zealand, and as part of his research in Alaska. During 1982-1983 he had a 6-month assignment with the Pa¬ thology Branch of the British Forestry Commission, where he was able to establish a worldwide culture col¬ lection of various species of Armillaria. He currently is involved with root disease research in the Southwest¬ ern United States, and with efforts to model the dy¬ namics, behavior, and impact of various forest pests, including Armillaria root disease. Address: USDA For¬ est Service, Rocky Mountain Forest and Range Experi¬ ment Station, 240 West Prospect, Fort Collins, CO 80526, USA. Glen A. Kile is a Senior Principal Research Scientist and Program Leader, CSIRO Division of Forestry and Forest Products. A Ph.D. graduate from the University of Tasmania (1972), his research with CSIRO has been concerned with etiology and epidemiology of diseases in native forests — Armillaria and stem rot fungi in eucalypts, and Chalara australis in Nothofagus rainforest. He discovered Armillaria luteobubalina is a major patho¬ gen in dry sclerophyll eucalypt forests, amenity tree plantings, and in horticulture in Australia, and has un¬ dertaken major taxonomic and population studies of Armillaria. He was Deputy Chairman and Chairman of the IUFRO Working Party on Root and Butt Rots of Forest Trees 1978-88. Address: CSIRO, Division of For¬ estry and Forest Products, Stowell Avenue, Battery Point, Tasmania 7004, Australia James B. Anderson is a Professor of Botany at the Uni¬ versity of Toronto, Erindale Campus. He obtained his B.A. in biology from the University of Rhode Island in 1975 and his Ph.D. in botany from the University of Vermont in 1980 shortly before he moved to his present appointment in Toronto. Dr. Anderson's research is concerned with the genetic control of mating and the process of speciation in higher fungi. His Ph.D. work was on sexual incompatibility and intersterility in Armillaria. Recently he has used molecular-genetic markers to examine phylogenetic relationships among species, and dispersal patterns in local environments. His current objective is to clone the mating-type genes of Armillaria and to compare their organization and ex¬ pression among different strains and species of Armillaria. Address: University of Toronto, Department of Botany, Erindale College, Mississuaga, Ontario, L5L 1C6 Canada . Harold H. Burdsall, Jr. is presently investigating the taxonomy and ecology of species of Armillaria and other forest fungi at the Center for Forest Mycology Re¬ search, USDA Forest Service, Madison. He received his B.A. degree in botany (1962) from Miami University and Ph.D. degree in mycology and plant pathology (1967) from Cornell University. He held a position with the Beltsville Forest Disease Laboratory (1967-71), then joined the Center for Forest Mycology Research in 1971. He became Project Leader of the Center in 1982. He has authored numerous papers on the taxonomy, nomenclature, and ecology of wood-rotting fungi. Cur¬ rent specific research interests concern species recogni¬ tion in Armillaria using immunological and biochemical techniques. He is a past President of the Mycological Society of America. Address: USDA Forest Service, Forest Products Laboratory, One Gifford Pinchot Drive, Madison, WI 53705-2398, USA. James W. Byler, Ph.D. University of California, Berke¬ ley (1970), is currently Pathology Supervisor with the USDA Forest Service Northern Region at Missoula, Montana. His experience with Armillaria root disease About the Authors 227 dates back to his M.S. research on yellow poplar root and butt rot at West Virginia University (1964). He and his staff are responsible for survey, evaluation, train¬ ing, and technical advice on Armillaria and other root diseases in northern Rocky Mountains. Address: USDA Forest Service, Cooperative Forestry and Pest Manage¬ ment, R-l, Federal Building, PO Box 7669, Missoula, MT 59807, USA. Gregory M. Filip is an Associate Professor of Forestry at Oregon State University in Corvallis, Oregon. Before joining Oregon State University in 1990, Dr. Filip was a Research Plant Pathologist with the USDA Forest Ser¬ vice, Pacific Northwest Research Station, in La Grande, Oregon. He completed a Ph. D at Oregon State Univer¬ sity in 1976 on chemical applications for control of Armillaria root disease of ponderosa pine. For the past 20 years he has been involved in research concerning the effects of silvicultural systems on biology and con¬ trol of Armillaria root disease in the Pacific Northwest region of the USA. Address: Department of Forest Sci¬ ence, College of Forestry, Oregon State University, Corvallis, Oregon 97331, USA. Michael O. Garraway is a Professor of Plant Pathology and Botany at The Ohio State University in Columbus, Ohio. He received his Ph.D. in plant pathology from the University of California, Berkeley with a study of the nutrition and physiology of Armillaria. He is espe¬ cially interested in the effects of nutrition on the devel¬ opment of rhizomorphs of Armillaria and conidia of Bipolar is maydis. He has chaired the Mycology Commit¬ tee of the American Phytopathological Society, is co¬ author of the textbook, "Fungal Nutrition and Physiology" and is a Fellow of the Ohio Academy of Science. Address: The Ohio State University, Depart¬ ment of Plant Pathology, 201 Kottman Hall, 2021 Coffey Road, Columbus, OH 43210-1087, USA. Steve C. Gregory studied at Cambridge and Aberdeen Universities and joined the Pathology Branch of the British Forestry Commission Research Division in 1972. He is now a Principal Scientific Officer at the Forestry Commission, Northern Research Station, near Edinburgh, Scotland. His Armillaria research has in¬ cluded studies of pathogenicity testing and the distri¬ bution and pathogenicity of species in northern Britain. Dr. Gregory has also worked on wound rot in spruces, and is currently working on stump colonization by Heterobasidion annosum. In addition to his research in¬ terests, he has major commitments to forest damage monitoring and to the Forestry Commission diagnostic md advisory service for tree owners. Address: British orestry Commission, Northern Research Station, ir 1 hdlothian EH25 9SY, Scotland. Jean Jacques Guillaumin is Charge de recherche de l e classe in the Institute National de la Recherche Agronomique stationed at Clermont Ferrand, France. A graduate of the Institut National Agronomique of Paris (1965), he subsequently completed the these de 3 e cycle (1971) on the morphogenesis of Sphaerostilbe repens and a these d'Etat (1986) on the life cycle, taxonomy, distri¬ bution, and pathogenic behavior of European Armillaria species. Much of his research has concerned root diseases of fruit and forest trees caused by Rosellinia necatrix, S. repens , Collybia fusipes , and Armillaria species. Other aspects he has studied with the latter species include infection process, control (es¬ pecially by the use of tolerant fruit tree root stocks), the role of Armillaria in certain forest declines in France, and the taxonomy and distribution of non-European Armillaria species. Address: Station de Pathologie Vegetale, 12 Avenue de 1'Agriculture, F-63.100, Clermont Ferrand, France. Susan Hagle is a Plant Pathologist for the USDA Forest Service, State and Private Forestry, in the Northern Re¬ gion. She specializes in technology development and transfer for root disease management in the northern Rocky Mountains. She holds a Ph.D. degree in forest pathology from the University of Idaho, and is a fac¬ ulty affiliate at the University of Montana and Oregon State University. For the past 10 years she has assisted forest managers in developing comprehensive manage¬ ment plans to minimize root disease losses and reha¬ bilitate root disease-affected sites. Hazard and risk rating are of particular interest. Address: USDA Forest Service, R-l, Federal Building, PO Box 7669, Missoula, MT 59807, USA. Thomas C. Harrington received degrees in plant pa¬ thology from Colorado State University (B.S., 1977), Washington State University (M.S., 1980), and the Uni¬ versity of California at Berkeley (Ph.D., 1983). He is now Associate Professor of Plant Pathology at the Uni¬ versity of New Hampshire, where he conducts research on the fungal associates of bark beetles and root dis¬ eases on conifers. Current investigations include stud¬ ies on wind induced tree stress, the identification of Armillaria species in New Hampshire, and the use of genetics to analyze clonal spread of Armillaria ostoyae and competing species. Address: Department of Plant Biology, University of New Hampshire, Durham, NH 03824, USA. Ian A. Hood (B.Sc., M.Sc., University of Auckland) has spent 21 years as a Forest Pathologist at New Zealand's Forest Research Institute, Rotorua, initiallv studving conifer needle diseases (Phaeocryptopus ganmannii in Douglas-fir; Cyclaneusma minus in radiata pine). In the last 10 years he has specialized in root diseases and About the Authors butt rots. He has a general interest in root and tree de¬ cay fungi in the Southeast Asia - Pacific region, and has carried out research in New Zealand, Fiji, and Canada. Address: Forest Research Institute, Pathology Section, Private Bag 3020, Rotorua, New Zealand. Aloys Hiittermann is Professor and Head of the Institut fur Forstbotanik, Universitat Gottingen. He re¬ ceived degrees (Diploma in Chemistry and Dr. rer. nat in Botany) at the University of Karlsruhe in 1964 and 1968, respectively. After post-doctoral work at the Uni¬ versity of Wisconsin, he moved to Gottingen, where he has worked since 1971 on the physiology and bio¬ chemical control of forest pathogens, particularly Heterobasidion annosum and Armillaria. Other research interests of his group include stress-physiology of for¬ est decline, soil microbiology, and ligninbiotechnology. He has been a visiting professor at the University of Tromso, Norway; Beijing, China; and Virginia Tech, Blackburg, USA. Address: Forstbotanisches Institut der Universitat Gottingen, D-3400 Gottingen-Weende, Busgenweg 2, West Germany. Kari Korhonen received a M.S. in biology from the University of Helsinki in 1970, and since then has worked as a Research Forest Pathologist at the Finnish Forest Research Institute at Helsinki and Vantaa. He has specialized in population and control studies of wood decay fungi, particularly Armillaria and Heterobasidion annosum. Following the recognition of bifactorial heterothallism in Armillaria, his research re¬ sulted in the recognition of the biological species of Armillaria in Europe. Address: Finnish Forest Research Institute, PO Box 18, SF-01301 Vantaa, Finland. Geral I. McDonald is Principal Plant Pathologist at the USDA Forest Service Intermountain Research Station, Moscow, Idaho. He received his B.Sc. degree in 1963 and Ph.D. degree in plant pathology in 1969 from Washington State University. Since joining the Intermountain Research Station in 1966, he has investi¬ gated the epidemiology and genetic interaction of Cronartium ribicola and its hosts, and genetic interac¬ tions between western spruce budworm and Douglas- fir and the relationship of this interaction to site quality. He is now investigating the ecological genetics of conifer interaction with Armillaria- caused root dis¬ eases. Address: USDA Forest Service, Forestry Sciences Laboratory, 1221 S. Main, Moscow, ID 83843, USA. Peter McNamee is a Partner and Senior Analyst with Environmental and Social Systems Analysts Ltd. (ESSA). He received his Ph.D. in systems ecology and forest pest management from the University of British Columbia in 1986. He currently leads ESSA's Forestry Group, which conducts projects for clients in Canada, the United States, and southeast Asia. He and his staff develop models, research, management, and monitor¬ ing plans, and decision support systems for forestry applications in the areas of pest management, land use, climate change, and integrated forest management planning. Current projects include development of an Annosus extension to the Western Root Disease Model and completing a decision support system for manage¬ ment of eastern hemlock looper in Newfoundland Ad¬ dress: Environmental and Social Systems Analysts Ltd., Box 12155 - Nelson Square, #705 - 808 Nelson Street, Vancouver, BC V6Z 2H2 Canada. Duncan J. Morrison is a Forest Pathologist with For¬ estry Canada at the Pacific Forestry Centre in Victoria, British Columbia. He received a B.Sc. in forestry and a M.Sc. in botany from the University of British Colum¬ bia in 1965 and 1968, respectively, and a Ph.D. in botany from the University of Cambridge in 1972. His Ph.D. thesis was concerned with factors affecting rhizomorph growth and variation in pathogenicity of Armillaria. Since joining Forestry Canada in 1966, Dr. Morrison has been involved in research on the biology and control of several root diseases, including those caused by Armillaria. Address: Environment Canada, Pacific Forest Research Centre, 506 West Burnside Road, Victoria, BC V8Z 1M5 Canada. Derek B. Redfern was educated at Aberdeen and Cam¬ bridge Universities. He was a Research Scientist with the Canadian Forestry Service, Maritime Forest Re¬ search Centre, New Brunswick 1966-68. In 1968 he joined the British Forestry Commission and is now Head of the Pathology Section, Northern Forest Re¬ search Station, Edinburgh. His research on Armillaria has included studies of pathogenicity and rhizomorph growth in soils, and he has also undertaken extensive research on Heterobasidion annosum. In his present posi¬ tion he also is involved in disease diagnoses and advi¬ sory work, with a particular interest in climatic damage to trees. Address: British Forestry Commission, North¬ ern Research Station, Roslin, Midlothian EH25 9SY Scotland. John Rishbeth was educated at Cambridge University. He was a Plant Pathologist in the West Indian Banana Research Scheme (1950-52) and a Lecturer-Reader in Plant Pathology at Cambridge University from 1953 until his retirement in 1984. He undertook pioneering research on the biology and control of Heterobasidion annosum from 1946-1967. From 1958 onward, he under¬ took extensive research on British species of Armillaria, including identification of species involved in attacks in a wide range of situations, modes of establishment in plantations, rate and extent of spread, and tests for pathogenicity. He supervised three post-graduate stu¬ dents in research on Armillaria. Address: c/o Botany School, Downing Street, Cambridge CB2 3EA, England About the Authors 229 Albert R. Stage is Principal Mensurationist at the For¬ estry Sciences Laboratory, Intermountain Research Sta¬ tion, Moscow, Idaho. He holds a Ph.D. in forestry from the University of Michigan. His research has included studies of planning methods for forest management, forest inventory techniques, site evaluation, and meth¬ ods for modelling forest establishment and develop¬ ment that include pest effects. He is Project Leader of a research team studying problems of developing deci¬ sion-support systems for forest management in the northern Rocky Mountains. This team has participated in multi-disciplinary research leading to specific models for impacts of Armillaria and Phellinus root diseases, mountain pine beetle in lodgepole pine, Douglas-fir tussock moth, and western spruce budworm. Address: USDA Forest Service, Forestry Sciences Laboratory, 1221 S. Main, Moscow, ID 83843, USA. Philip M. Wargo is Principal Plant Pathologist and Leader of the USDA Forest Service's Stress-triggered Tree Disease Project at the Center for Biological Control of Insects and Diseases of Northeastern Forests, North¬ eastern Forest Experiment Station, Hamden, Connecti¬ cut. He joined the Forest Service in 1969 after completing his Ph.D. degree in plant pathology at Iowa State University, Ames. His research has focused mainly on physiological effects of stress, and how stress predisposes trees to pathogenic organisms in die- back and decline diseases. A continuing interest has been developing and evaluating techniques and proce¬ dures to measure the effects of stress on trees. Much of his work has been on root physiology and vitality and root disease organisms, especially Armillaria species. Address: USDA Forest Service, Northeastern Forest Experiment Station, 51 Mill Pond Road, Hamden CT 06514, USA. Roy Watling is a Senior Principal Scientific Officer at the Royal Botanic Garden, Edinburgh. His Ph.D. from Edinburgh University covered ecological studies and taxonomy of the Bolbitiaceae. He is the author of sev¬ eral publications on agaric taxonomy and ecology, including European and Australian members of the ge¬ nus Armillaria, and an editor and main contributor to the British Fungus Flora. He has been awarded a D.Sc. for his contributions, and is a former President of the British Mycological Society. Address: Royal Botanic Garden, Edinburgh EH3 5LR, Scotland. Roy D. Whitney is a Forest Pathologist with Forestry Canada at the Great Lakes Forestry Centre, Sault Ste. Marie, Ontario. He received a B.S.F. from University of British Columbia, a M.F. from Yale, and a Ph.D. from Queen's. His Ph.D. research for Forestrv Canada in / Saskatchewan and Ontario has been on the biology and control of the conifer root disease caused by Inonotus tomentosus. More recently, his work has dealt with the root diseases, including Armillaria, that affect conifers in northern Ontario. Address: c/o Great Lakes Forest Research Centre, Environment Canada, PO Box 490, Sault Ste. Marie, Ontario P6A 5M7, Canada. Ralph E. Williams is Supervisory Plant Pathologist/ Field Representative for the USDA Forest Service, Intermountain Region, Boise, Idaho. A graduate of the University of Idaho and Washington State University, his research and management responsibilities since 1970 have included root diseases, nursery diseases, white pine blister rust, and insect damage. He has de¬ veloped remote sensing techniques for surveying root diseases including Armillaria and Phellinus zveirii, mod¬ eled disease center, site, and stand associations, and established studies to evaluate effects of various stand management practices on root disease development and expression. Address: USDA Forest Service, R-4, Forest Pest Management, 1750 Front Street, Boise, ID 83701, USA. About the Authors Index adventitious wot development—63,65,66,70 aeration, effect on growth—33 Africa—128-129 alkaloids, response to—43 alleles—10-13,15,18 antibiotics—37,38 Armillaria root disease Africa—128-129 Asia and the Pacific—129-130 Australasia—130-132 Central and South America—127,128 Europe and Soviet China—123-125 geographical distribution in planted crops—122-132 hosts in planted crops — 124,126-129,131,132,134, 135,137,139,140-149 in plantations—122-139 North America—125-127 Asia and the Pacific—129-130 Australasia—130-132 barriers to infection—41 basal cankers—106,114 basidiome ontogeny (development) — 21,22 basidiome development in vitro—21,22 basidiospore infection—48,49,52,54 biochemical changes in growth and development—28,34 biological species—4,9-10,15-19 bioluminescence—38 butt rot-103,106-110,113,115,119 CIN ratios—30 callus tissue—63-65 carbohydrate x alcohol interaction—30,31 carbon sources—28-30,44,46 cell wall composition—25-27,34,40 Central and South America—127,128 clones—117,118 compatible mating—11-15,18,19 conifer root rot—110-113 boreal and coastal forests, North America—110 mixed conifers, North America — 111-113 pines (Europe)—110,111 cojitrol methods (see management actions) avoidance of hazardous sites—157,160 biological control—168-172 chemical—166-168 fallowing—164,173 fungicides—166-168 genetic resistance—161,162,173 inoculum reduction—158,163,164,172 mycorrhizal—172 previous vegetation type—160,161 regeneration methods—162 resistant species—161,162 ring barking (girdling)—165,171 root collar exposure—165,166 rotation length—162 silviculture—158-160 soil factors—163 soil fumigation—166 species selection, species composition—160-163 stand density manipulation (thinning)—162,163 stump fumigation—171 systemic chemicals—167 thinning—162,163 trenching (barriers)—165 control of other pests and diseases—139 cord forming fungi as antagonists—170,171 DNA differences—4,8,9 decay—73,74 decomposer (saprotroph)—103,105 diploid—10-13 disease center dynamics—152,154,155 disease carryover—155,156 inoculum longevity—154 disease control—157-171 concepts—157-158 needs assessment—158 disease detection — 74,75 aerial—74 ground survey—74,75 disease diagnosis—68-75 dieback—69 foliage effects—68 growth reduction—68 stress induced reproduction—69 disease dynamics in plantations—132-136 disease establishment—132 distribution pattern—134 secondary disease spread—134 Index 231 disease in monocultures—132 disease epidemiology—118,119 disease losses (see control)—136 crop age and type—136 evaluation of impact—136 disease signs basidiomes—62,73 mycelial fans—64,66,71-73 rhizomorphs—62,71 distal vs proximal rate of spread—66 distribution and dispersal—117 basidiospores—116 rhizomorphs and root contacts—117 ecological strategies—105 ectotrophic growth—64,66 energy reserves—90-92,101 carbohyd rates—92 oleoresins—91 starch—90-92 enzyme systems—35,36 eucalypt root rot—105,107,110,114,117-119 Europe and Soviet China—123-125 facilitative necrotroph—103 fertilization—58,138,139 fire effects-58,105,115,118-121,159,169,171,172 food base—48,50,51,56,57,60,61 forest dieback and decline—106,114-116 forest management and disease—119-121 forest succession (habitat type)—113,159,160 fungal persistence—52,53 generic characteristics—3 generic name—1,2 genotype identification—18-19 geographical distribution of species—102,103,112 growth inhibition—41 phenolics—41 growth stimulation—28,34,41 alcohols—29-31,34 auxins—30-32 fatty acids—31 lipids—29,31,35,43 haploid—10-13 hemicompatible mating—11,12 host induced lysis—26,46 host pathogen interaction—39,46 genetic control—39 metabolic control—39 host range—103,104 host specialization—78,84 host resistance—77-80 ; response to infection—63 '"^chemical—63,65,75 exudates—63,69 ■m nstematic—63 host i -88,89,94,95,97,101 incompatible mating—11,12 infection—51-61 effects of inoculum potential—46,48,58 in relation to inoculum distribution—58,59 in relation to wounds—60 rhizomorphs—59,60 root contact—57-59 infection process—62,64,75 inoculum—48-61 effect on infection—59 longevity (persistence)—52,53,154 rhizomorph production—60 inoculum (substrate) quality—49-52 inoculum potential—48,57 in plantations—132,134,137,138 inoculum persistence (carryover)—155,156 inorganic nutrients—23,30 interaction between Armillaria and fumigants—166, 168,169 isolation techniques—73 lethal primary disease—110-114 light, effect on growth—22,23,33 management actions (see control)—151,155 fertilization—58,138,139,163 inoculum removal—155 silvicultural treatment—58,155-156 thinning and pruning—138,162-163 weed control—138 management and disease—137 natural forests—126,132,134-136 plantations—132 (see Armillaria root disease in plantations) mating reactions—11-12,15,19 mycopa rasite—105,121 mycotrophic (mycorrhizal) associations—105,106,121 necrotroph—106 nitrogen sources—29 nomenclature—1,2,5,9 non-lethal pritnan / disease—106-110 North America—125-127 nutrition—28,30,44 partial cutting—120 pathogenicity—76-87 assessment of results—78 differences between species—77,84-87 field observations—83-87 indirect methods of assessment—83,84 inoculation methods—81,82 methods of assessment—83 plant species—84-87 role of rhizomorphs—82,83 size of inoculum—81,82 type of inoculum—81,82 pathogenicity (relative)—107,110,114,115,118,121 pH, effect on growth—33 232 Index ohenolics (resins), role in growth and development—30,40,41,43,45 ohenolics, use for species differentiation—41 physiology—28,30 predisposition — 88-90,92,93,95,96,99,100 primary pathogen — 102,106,114,115,119 primary pathogen—76,80,87 protease—37,36 pseudosclerotial plates—21,23 resistance—88-90,92,94,101 rhizomorphs branching pattern—53,54 distribution—56 factors affecting development (moisture, pH, etc.) — 53-55,58 growth in soil—53 infection process—62,63 networks—56-58,60 structure—28 transport of water and nutrients—27 rhizomorph differentiation—25-27 ring disease—110,111 root disease model (Western)—151,152,155,156 assumptions—150-153,156 identification of research needs—156 root disease modeling—150-156 root lesions — 64-66,70,106,107,113,115,118,119 saprophytic behavior—77 secondary pathogen—77,80,115,121 selective logging, effects on disease development—119-121,159 selective media—73 serological differences—6,8 sexual system—11-12 silvicultural control options—158-160 site factors—89 soil organic matter—34 somatic haploid—13 somatic segregants—13 spatial distributions of fungal populations—115,117,121 species choice—132 species list—5 species relationships—4 spread within roots—66 stress — 43,44,88-101 acute, chronic—88,90,93 insect damage—98 interaction—88 light—93,94 moisture—94,95 nutritional factors—95,96 other insects—99 other pathogens—99 other soil factors—95,96 partial cutting—97 pollutants—96,97 temperature—94 timing—93 stress management—101 stress and predisposition—124,132,135-139 structure and morphogenesis—21 suberinase—39 substrate specialization—51,52 symptoms of disease—68 basal cankers—69-71 butt rot—67,70,74 diameter—66,67 foliage—67,68,70,74 height growth (shoot)—68,71 resin—64,65,67,70,71 taxonomic characters—6 temperature, effect on growth—33,38 toxins—67,68 Trichoderma—269,2 72,2 72 typification—2,3 vascular disruption—67,68 vegetative diploidy—12,14 virulence—76-87 assessment—78-81,83-84 vitamins—23,30 white pine blister rust—121 wood as inoculum in trials—48 wood decay—23,70,73 wood decay fungi—170 wounds—90,91,99 zone lines—23 Index 233 30112619250049