key: cord-0823693-zl0otg0s authors: Rodino, Kyle G.; Smith, Kenneth P.; Pettengill, Matthew A. title: Novel assays for molecular detection of SARS-CoV-2 date: 2022-03-09 journal: Clin Lab Med DOI: 10.1016/j.cll.2022.02.004 sha: b0ac8a742e501ad5db226513aca0dad4acda6b56 doc_id: 823693 cord_uid: zl0otg0s From the onset of the SARS-CoV-2 / COVID-19 pandemic there has been a major emphasis on molecular laboratory tests for the virus. Shortages in various testing supplies, the desire to increase testing capacity, and a push to make point-of-care or home-based testing available has fostered considerable innovation for SARS-CoV-2 molecular diagnostics, advancements likely to be applicable to other diagnostic uses. We attempt to cover some of the most compelling novel types of molecular assays or novel approaches in adapting established molecular methodologies for SARS-CoV-2 detection or characterization. From the onset of the SARS-CoV-2 / COVID-19 pandemic there has been a major emphasis on molecular laboratory tests for the virus. Shortages in various testing supplies, the desire to increase testing capacity, and a push to make point-of-care or home-based testing available has fostered considerable innovation for SARS-CoV-2 molecular diagnostics, advancements likely to be applicable to other diagnostic uses. We attempt to cover some of the most compelling novel types of molecular assays or novel approaches in adapting established molecular methodologies for SARS-CoV-2 detection or characterization. The COVID-19 pandemic led to unprecedented demand for laboratory testing that far outpaced existing capacity. Shortages of supplies and labor exacerbated the problem, resulting in the need for improved efficiency of existing SARS-CoV-2 diagnostics, including specimen pooling and process enhancements. Novel diagnostic methodologies that increase efficiency or speed of testing will be discussed elsewhere. The simplest way to increase testing efficiency is to pool multiple patient specimens in a single RT-PCR reaction. If a pool tests negative, all patients are therefore negative. If the pool tests positive, all patient specimens that comprise that pool must be tested individually to identify the positive patient(s). However, pooling may reduce sensitivity due to sample dilution of weakly positive specimens 1 . It is also impractical to implement in high (>10%) prevalence settings where specimen pools are more likely to be positive, necessitating extensive re-testing of individual patients 2 . However, as SARS-CoV-2 prevalence decreases, specimen pooling may become an attractive option. Efficiency can also be enhanced by implementing changes in the typical RT-PCR testing workflow. One of the most significant rate-limiting steps in RT-PCR is the extraction step. This step purifies nucleic acids from the milieu of patient cells and proteins present in primary specimens and reduces inhibitory substances. However, swabs in VTM are a relatively simple sample matrix and potential inhibition can easily be assessed by monitoring internal controls, suggesting extraction may not be necessary. As such, labs have developed "extraction free" methods that rely on heat to inactivate virus and lyse cells but lack a traditional nucleic acid purification step. These methodologies are easy to implement and have been demonstrated to be faster than conventional methods while maintaining similar performance characteristics 3,4 . Viral replication is error prone, leading to mutations in the viral genome 5 . When these mutations confer a selective advantage, genomic variants can emerge and become dominant. From the beginning of the pandemic, variants with mutations deviating from the initial SARS-CoV-2 genomic sequence have be recognized and sorted into a variety of lineage classifications. Beyond these viral phylogenetic relationships, many health organizations have adapted variant classifications based on public health impact, with the United States Centers for Disease Control and Prevention (CDC) classifies SARS-CoV-2 variants into three groups: "variant of interest", "variant of concern", and "variant of high consequence". Each level of classification denotes more significant changes in overall prevalence, viral transmissibility, disease severity, antiviral resistance, and/or vaccine evasion. As such, detection of variants is of epidemiological, and in some cases clinical, interest. SARS-CoV-2 variants are first identified by genetic sequencing (often next generation sequencing, reviewed elsewhere in this edition). However, sequencing methodologies are time J o u r n a l P r e -p r o o f consuming, expensive, and difficult to deploy at large scale as a routine diagnostic. As such, most SARS-CoV-2 tests are performed by other methods, with RT-PCR remaining the gold standard. RT-PCR tests rely on primers and probes that hybridize to known sequences within the viral genome. Therefore, these assays detect sequences for which they were designed with high sensitivity and specificity. However, when faced with a variant containing changes in the assay's genetic target, the virus may evade detection 6 . Further, variants with no changes in the assay's target will be indistinguishable from any other positive result. In some cases, knowledge of these limitations can be leveraged to identify variants. For example, the assay used in the United Kingdom's national SARS-CoV-2 testing system contains targets for the nucleocapsid gene (N), the spike gene (S), a gene of unknown function (ORFab). In November 2020, a cluster of cases in Kent, England was identified in which the N and ORFab targets yielded positive results, but the S gene was consistently negative (https://assets.publishing.service.gov.uk/government/uploads/system/uploads/attachment_data/f ile/959360/Variant_of_Concern_VOC_202012_01_Technical_Briefing_3.pdf). Failure of one or more gene targets in an RT-PCR assay is referred to as gene dropout. This specific pattern of gene dropout was widely recognized and given the name S gene target failure (SGTF). Further investigation of this phenomenon led to the identification of the Alpha SARS-CoV-2 variant (also known as B.1.1.7) which contains a 6-nucleotide deletion within the probe binding site, precluding detection of the S gene. Since this discovery, other assays, including commercially available assays, have used SGTF as a proxy for the Alpha strain 7 . Although positive predictive value of SGTF is good in high-prevalence settings, the Alpha variant's nucleotide deletion also occurs in other variants (notably the Beta variant, also known as B.1.351). Detection of SGTF regained value with the emergence of Omicron, which shared the same deletion in the spike coding region with Alpha. Conveniently, the preceding Delta variant did not contain the same deletion, making SGTF a reliable proxy for classification as Omicron (BA.1) 8, 9 . This lack of specificity highlights an important limitation of using gene dropout as a detection method and suggests need for variant targeting PCR tests. Additionally, relevant to this type of assay generally, the LoD of the assay can create a "false-dropout" when a particular gene is not detected due to low positive. RT-PCR tests can be adapted for variant detection by incorporating variant-specific probes to existing assays. Typically, these would be multiplexed in the same reaction to allow detection of the widest possible number of variants 10 . However, multiplex assays suffer from the same limitations as single-plex assays in that genetic targets must be known in advance. Shifting variant makeup can render a panel with limited mutation targets obsolete or lose specificity if multiple lineages emerge with overlapping mutation combinations, which can be particularly challenging in the clinical lab given the significant time and financial investment needed for assay validation. Therefore, unbiased methods such as genetic sequencing will likely remain major methods of variant detection. Current CDC guidelines suggest discontinuation of SARS-CoV-2 isolation precautions by a time and symptom-based strategy. While this policy, based on generalized viral kinetics and disease timeline, may be sufficient when applied broadly, data suggest prolonged disease and extended infectivity in severely immunocompromised populations 11 While CRISPR-Cas systems do not amplify the target nucleic acid, a potential advantage of CRISPR-Cas in the detection phase is that some Cas nucleases produce signal amplification in that they can produce multiple signal molecule events per sequence specific binding event. When coupled with a priming amplification phase it may be possible to modestly improve on the limit of detection relative to standard molecular assays for SARS-CoV-2 35, 37 , but in existing studies the limit of detection is still similar to PCR and thus not likely to offer a meaningful difference in clinical sensitivity. Most studies evaluating CRISPR-Cas diagnostic applications utilize a paired amplification assay as described above, but to move CRISPR-Cas assays to the point of care and potentially reduce assay expense it is possible to develop assays that use CRISPR-Cas detection directly on specimens with no nucleic acid extraction, and no pre-amplification 36 . Although there is some signal amplification, the lack of nucleic acid amplification does leave this J o u r n a l P r e -p r o o f approach with considerably higher limits-of-detection (>100x) relative to RT-PCR, which could considerably impact the clinical sensitivity of SARS-CoV-2 assays depending somewhat on the population tested and application 38 . Like some other molecular methodologies, CRISPR-Cas assays are amenable to scale up for high-throughput application and may perform well for SARS-CoV-2 without nucleic acid extraction procedures 39 . While CRISPR-Cas technology offers the potential for signal boosting to modestly improve the limit of detection for compatible SARS-CoV-2 testing methodologies, and is also amenable to use for direct detection when sufficient target sequence is expected in specimens, it does not appear at this time that these types of applications will lead to significant improvement in test performance characteristics relative to RT-PCR and other established methods. Microfluidic devices essentially take advantage of the physical properties of fluids to direct or even manipulate the movement of fluid specimens through engineered microchannels or material substrates. These processes may naturally separate or concentrate an analyte of interest, or may be made to do so by applying an electrical charge. Lateral flow assays are a very simplistic variety of microfluidic device, and commonly used and familiar to clinical microbiologists, but there are far more sophisticated device designs as well, and different microfluidic device designs have been evaluated for developing inexpensive diagnostic tests that require no, or less, equipment than standard types of diagnostic assays 40, 41 . We limit further discussion here to microfluidic devices evaluated with molecular SARS-CoV-2 diagnostic assays. Isotachophoresis (ITP), a method utilizing electrophoresis to separate and concentrate charged analytes, was used in early 2020 to develop a novel microfluidic SARS-CoV-2 detection assay 42 . ITP helped perform nucleic acid extraction and concentration with limited reagent requirements, J o u r n a l P r e -p r o o f followed by off chip LAMP and returning to the microfluidic device for small volume (0.2 microliter) CRISPR-based detection phase in less than 30 minutes from start to finish. In a limited sample set this test had relatively good positive percent agreement (94%) compared to a standard RT-PCR assay. While the transition on and off chip for different phases does not result in an assay that would be appealing in its current form, this study was a technical achievement and demonstrates the potential to use ITP in a microfluidic device to reduce sample handling and dramatically reduce reagent requirements. Assessment of Specimen Pooling to Conserve SARS CoV-2 Testing Resources Massive and rapid COVID-19 testing is feasible by extractionfree SARS-CoV-2 RT-PCR Implementation of an extraction-free COVID RT-PCR workflow in a pediatric hospital setting Why are RNA virus mutation rates so damn high? 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