Created by: Leena Dhaliwal, Adnan Hassanali, Tristen Nimojan, Abdullah Syeddan

Figure 1: Illustration of SARS-CoV-2. Adapted from Eckert & Higgins (2020).

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) is the virus that caused a disease outbreak which originated in China in late 2019 (Sheposh, 2021). The name of the disease caused by this virus is coronavirus disease 2019, more commonly known as COVID-19 (Sheposh, 2021). This outbreak quickly spread across the globe and was declared a global pandemic by the World Health Organization (WHO) on March 11, 2020. As of January 27, 2021, the number of confirmed cases is 99,864,391 and the number of confirmed deaths is 2,149,700 globally (WHO, 2021).

The incubation period of SARS-CoV-2 is typically around five days and infected individuals tend to develop symptoms within approximately twelve days of infection (Lauer et al., 2020). During the time where the infected individual has not yet begun to develop symptoms, they are still able to transmit the virus (He et al., 2020). In fact, a sizable portion of person to person transmission may occur during this time period (He et al., 2020). Additionally, some individuals may be asymptomatic, in other words they never develop any symptoms (Li et al., 2020). These individuals may substantially contribute to disease transmission (Li et al., 2020). Transmission of this virus can occur via direct physical contact with an infected person or a contaminated surface as well as through respiratory droplets formed when an infectious individual sneezes, breathes, coughs, or speaks (Sheposh, 2021). Symptoms of COVID-19 tend to resemble that of influenza with the common symptoms being fever, dry coughs, shortness of breath, and fatigue (Sheposh, 2021). Other symptoms may include muscle pain, nausea, vomiting, diarrhea, runny nose, nasal congestion, sore throat, chills, headache, and loss of smell and/or taste (Sheposh, 2021). According to the CDC (2021), individuals infected with COVID-19 are the most contagious when they are presenting symptoms. Although most cases are mild, severe cases of COVID-19 can result in pneumonia, kidney failure, and death (Sheposh, 2021).

COVID-19 diagnostic tests function to identify people who are infected with SARS-CoV-2. Covid-19 diagnostic tests include nucleic acid tests (e.g., RT-PCR, CRISPR, isothermal amplification) and serological tests (e.g., ELISA, LFA) among other diagnostic methods such as the use of computed tomography (CT) and artificial intelligence (AI). All these diagnostic tests play important roles in hospitals, point-of-care, or large-scale population testing (Weissleder et al., 2020).

The validity of these tests can be measured by their analytical and clinical sensitivities and specificities. (Chau et al., 2020). Analytical sensitivity is the ability of the assay to reliably detect the minimum amount of the target substance within a sample (referred to as the limit of detection) (Chau et al., 2020). Analytical specificity is the ability of an assay to detect only the analyte being measured without cross-reacting with other substances (Chau et al., 2020). Clinical sensitivity measures the ability of a test to correctly generate a positive test result for individuals who have the condition being tested for (“true positive” - a person is infected and tests positive) (Chau et al., 2020). For example, a test that has 95% sensitivity will correctly identify 95% of patients who have the condition while 5% of patients will be “false negatives” (patients who have the condition but the test produces a negative result). A test with high sensitivity will generate fewer false negatives results. Clinical specificity determines the test’s ability to accurately produce a negative test for individuals who do not have the condition being tested for (“true negative” - a person is not infected and receives a negative test result) (Chau et al., 2020). A test that has 95% specificity will correctly identify patients who do not have the condition, while 5% of patients will be “false positives” (patients who test positive but do not have the condition). A higher specificity would mean fewer false positives. In all of these cases, the condition being tested for would be whether or not the patient has COVID-19.

Diagnostic testing for COVID-19 is essential for controlling the pandemic (Halliday, 2020). Individuals presenting symptoms of COVID-19 are to be tested in an attempt to prevent further transmission (Halliday, 2020). Confirming a positive result can help to determine the next steps whether that be isolation, administer treatment, and/or hospitalization. Diagnostic testing is also important for contact tracing which involves identifying all the people that the covid positive patient has come into contact with during the last two weeks (CDC, 2021). These individuals can then monitor themselves for COVID-19 symptoms and self-isolate for 14 days after their last contact with the patient (CDC, 2021). If the contacts show any COVID-19 symptoms, then they should get tested (CDC, 2021). This measure is in place to slow down the spread of COVID-19.

Diagnostic testing is also important for public health as it provides valuable information regarding the prevalence, spread, and contagiousness of the viral infection (Sanchez, 2020). Furthermore, diagnostic testing can provide information regarding the presence of COVID-19 specific antibodies within the blood plasma (Sanchez, 2020). This information helps public health officials get a sense of the level of immunity within the population (Sanchez, 2020). The more individuals within the population that have immunity, the greater the percentage of “herd immunity” (Sanchez, 2020). Herd immunity functions to protect the larger community as it limits the transmission of SARS-CoV-2 from person to person (Sanchez, 2020). However, it is not guaranteed that prior infection with COVID-19 will protect an individual from future infection (Mayo Clinic, 2020). There have been several accounts of patients contracting COVID-19 more than once, however the second infection is typically mild (Mayo Clinic, 2020). The ideal method of achieving herd immunity is through vaccinations (Mayo Clinic, 2020). As opposed to immunity achieved through natural infection, vaccinations provide a way to achieve similar results without risking serious complications or death that may result from COVID-19 (Mayo Clinic, 2020).

The decision on whether or not to test for COVID-19 should be based on the individual’s likelihood of contact with a COVID-19 carrier (WHO, 2020). In such a case, individuals that are considered suspect cases should be screened for the virus using nucleic acid amplification tests (NAAT) such as RT-PCR (WHO, 2020). The World Health Organization (WHO) recommends that suspect cases should also undergo additional screening for other respiratory diseases as co-infections of respiratory pathogens can occur (WHO, 2020).

Viral detection can occur in various samples including blood and stool but the greatest yield of viral specimens have been found to occur in respiratory samples (WHO, 2020). As a result, at minimum respiratory samples must be collected from the upper respiratory tract (via nasopharyngeal swabs or wash) and/or the lower respiratory tract (via sputum, endotracheal aspirate and/or bronchoalveolar lavage) (WHO, 2020). It is important to note that specimen collection from the lower respiratory tract offers a high risk of aerosolization and should be performed carefully and with proper procedure (WHO, 2020). In deceased patients, collection of lung tissue during autopsy is recommended (WHO, 2020). In patients who have survived and recovered from the virus, retrospective assays like serological testing can be useful for defining cases (WHO, 2020). Collected specimens should be transported to laboratories in 2-8°C and stored in temperatures below -20°C (ideally at -70°C) (WHO, 2020).

Laboratory testing for COVID-19 should strictly follow standard safety procedures to avoid further infection (WHO, 2020). Conformation of COVID-19 infection is routinely done via nucleic acid amplification tests (NAAT) such as real-time reverse transcriptase polymerase chain reaction (rRT-PCR) and should be considered the gold-standard for COVID-19 detection (WHO, 2020). The viral genes that have been targeted currently include the E, N, S and RdRp-coding genes as well as the RdRp Helicase domain (WHO, 2020). NAAT testing should be paired with high throughput sequencing if positive testing occurs in areas with no known COVID-19 viral circulation to confirm if the infection is truly caused by the SARS-CoV-2 pathogen (WHO, 2020). Serological samples can be collected to retroactively diagnose COVID-19 infection by measuring serum SARS-CoV-2 antibody levels (WHO, 2020). Serological samples should be stored to aid epidemiological studies on the outbreaks and spread of COVID-19 in different areas (WHO, 2020). Sequencing of the viral genome from collected samples should be done both to confirm the diagnosis of COVID-19 infection and to monitor for viral mutations that may affect currently available medical interventions (WHO, 2020).

Comparing the performance of three RT-PCR assays targeting SARS-CoV-2 RdRp Helicase, Spike and nucleocapsid genes with the RdRp-P2 assay (2020). The COVID-19 RdRp Helicase assay shows the lowest limit of detection in vitro and does not cross-react with other human coronaviruses or other respiratory pathogens in cell culture and clinical subjects (Chan et al., 2020).The COVID-19 RdRp Helicase assay may improve laboratory diagnosis of SARS-CoV-2 infection due to the assay’s high sensitivity and specificity (Chan et al., 2020).

Based on information gathered from previous coronavirus epidemics (MERS-CoV and SARS-CoV-), it is understood that high sensitivity assays and diagnostics are essential for identifying cases, performing contact tracing and designing infection control measures (Chan et al., 2020). Standard viral culturing in Vero E6 cells is found to be impractical for acute diagnoses because of the three day period required for SARS-CoV-2 to show significant CPE (Chan et al., 2020). Additionally, serum testing as a technique for diagnosing SARS-CoV-2 can only be used to retroactively diagnose COVID-19 infection and is not useful for acute diagnoses (Chan et al., 2020). As a result of these limitations, RT-PCR maintains its status as the gold standard for acute COVID-19 detection and diagnosis (Chan et al., 2020).

RT-PCR allows the progress of the PCR reaction to be monitored in real time (ThermoFisher, 2020). This method allows the measurement of the amount of amplicon after each cycle which allows for highly accurate quantification (ThermoFisher, 2020). Additionally, Cross-contamination is eliminated by allowing amplification and detection to occur in a single test tube (ThermoFisher, 2020). Low experimental time allows the results of the experiment and subsequent diagnosis to be received within 3-5 hours of sample collection (ThermoFisher, 2020). After thorough testing against other respiratory pathogens it has been found that there is no cross-reactivity with other respiratory infections or coronavirus strains (ThermoFisher, 2020) allowing for confident results.

There are high costs for laboratory equipment and training (ThermoFisher, 2020). Additionally, This method is unable to retroactively diagnose patients with COVID-19 because the target material is not present after the acute phase of the infection has passed (ThermoFisher, 2020).

RT-LAMP is a diagnostic tool used in the detection of RNA viruses such as SARS-CoV-2. The reagents used in RT-LAMP include: forward and reverse, inner, outer, and loop primers, deoxynucleotide triphosphates (dNTPs), isothermal amplification buffer, MgSO4, heat-stable reverse transcriptase enzyme, and DNA polymerase (Lamb et al., 2020). Uracil-DNA glycosylase can also be added to reduce cross contamination from previous reactions (Lamb et al., 2020). The reaction involves adding the reagents to the samples collected from the COVID-19 patients (Lamb et al., 2020). Reactions are set up on ice, incubated at 63˚C for 30 minutes, and then subjected to 80˚C for 10 minutes for inactivation (Lamb et al., 2020). Then to visualize the results of the reaction, SYBR Green 1 can be added to the reaction tube where a change in colour (orange to yellow) indicates positive amplification (Lamb et al., 2020). The tubes can also be imaged under a UV light to check for the production of a fluorescent signal which would confirm a positive result (Lamb et al., 2020). Lastly, agarose gel electrophoresis can be used in the analysis of RT-Lamp as a laddering pattern on the gel would indicate positive amplification (Lamb et al., 2020).

Figure 2: Schematic illustrating the loop-mediated isothermal amplification (LAMP) primers. The inner primers (FIP, BIP), outer primers (F3, B3), and loop primers (LF, LB) are shown. Adapted from Mori & Notomi (2009).

Although RT-PCR is considered the gold standard for COVID-19 detection, RT-LAMP offers a faster and less expensive alternative (Figure 2) (Lamb et al., 2020). RT-LAMP is a one-step nucleic acid amplification method that can be completed in less than an hour, uses relatively inexpensive stable reagents, works at various temperature and pH ranges, and offers high sensitivity and specificity (Lamb et al., 2020). The high specificity of LAMP is a result of the amplification reaction occurring only when all six primers are bound to the target DNA sequence (Mori & Notomi, 2009). One of the main reasons that RT-PCR is more time efficient and cost effective in comparison to PCR is that it does not require thermocycling (Fu et al. 2011). Furthermore, this assay does not require special training or equipment to be performed (Lamb et al., 2020). These features make RT-LAMP a good candidate for point-of-care testing (POCT) (Lamb et al., 2020). POC tests are medical diagnostics tests that are performed outside the clinical laboratory (Schilling, 2015). POC tests can be performed at the time and place that the patient is being treated and results are produced in a matter of minutes as opposed to hours (Schilling, 2015). Therefore, POCT allows healthcare providers to make treatment decisions in a timely manner as to optimize patient care (Schilling, 2015).

Researchers oftentimes have difficulty designing LAMP primers that are compatible with the target DNA sequence (Mori & Notomi, 2009). However, to aid in this process, there are softwares available that help generate candidate LAMP primers to the given target sequence (Mori & Notomi, 2009).

Figure 3: COVID-19 ELISA Test Protocol. Adapted from Epitope Diagnostics Inc. (2020).

The ELISA test is a highly specific and sensitive test that uses purified protein as antigen for binding of host antibodies (Serrano et al., 2020). Seroconversion typically occurs 1-3 weeks after exposure at which point IgG, IgM and IgA antibodies specific to SARS-COV-2 have the potential to be recognized (Serrano et al 2020). As a diagnostic tool for COVID-19, a blood sample is taken from the patient, diluted and added into microwells (Epitope Diagnostics Inc., 2020). Typically, the microwells contain purified SARS-COV-2 protein (such as nucleoprotein and spike protein) as antigen for binding of host antibodies, if present. After periods of incubation, Horseradish peroxidase (HRP)-labelled or alkaline phosphatase (AP)-labelled secondary antibody is added to the microwell (Epitope Diagnostics Inc., 2020). The addition of a secondary antibody qualifies the test as an indirect ELISA (Gibbs et al., 2020). There are many variants of ELISA testing which differ based on the addition of certain secondary antibodies or substrates. Substrates added to the wells can be either chromogenic, chemiluminescent or fluorescent and can change the method of detection. A chromogenic ELISA gives rise to coloured products, a chemiluminescent ELISA captures the emission of photons and a fluorescent ELISA which gives rise to fluorescent products after being excited by light (Gibbs et al., 2020). Fluorescent ELISAs are also slightly more sensitive than the other listed ELISAs (Gibbs et al., 2020). Substrates used for HRP are oxidized with the use of hydrogen peroxide while substrates for AP are dephosphorylated to produce the active product (Michigan Diagnostics, 2020). The results of the test are determined if a signal is found depending on the substrate used. If a SARS-COV-2 specific antibody is present, the substrate is converted into active form and can be assessed by measuring absorbance (Serrano et al., 2020).

Serological ELISA test has greater specificity and sensitivity after the seroconversion period while the accuracy of RT-PCR performed on nasopharyngeal swabs begins to decline at this point (Serrano et al., 2020). Thus, serological ELISA test is a good tool to assess if a patient has previously been infected by COVID-19, as SARS-COV-2 specific antibodies last for at least 8 months (Choe et al., 2021). Using only one antibody isotype will yield lower specificity and sensitivity, thus, ELISA tests utilize multiple, such as both IgA and IgG in order to garner a high specificity and sensitivity (Serrano et al., 2020). Antibody tests are advantageous as epidemiological tools to assess population seropositivity as well as for contact tracing for individuals who were asymptomatic or were not PCR tested during acute illness (Serrano et al., 2020).

Serological ELISA tests are very inaccurate during acute illness as the adaptive immune system has not had adequate time for proliferation of SARS-COV-2 specific B and plasma cells (Serrano et al., 2020). Thus, the patient lacks the necessary antibodies to produce a positive signal during ELISA testing. Serological ELISA tests also require sophisticated equipment and skilled operators while being time-consuming and expensive (Serrano et al., 2020).

Rapid lateral flow assays have come a long way when compared to earlier time points in the COVID-19 pandemic. Previously, the first approved rapid LFA test by the WHO was found to be 20% less specific than the RT-PCR test (FIND, 2020). The specificity and sensitivity of these tests now almost match the gold standards of COVID-19 diagnostics, that being the RT-PCR test and serological antibody test (Wang et al., 2020). Rapid lateral flow immunoassays use DNA probes that are complementary to conserved genes in the SARS-COV-2 genome, these include: ORF1, envelope protein and nucleocapsid protein (Wang et al., 2020). A sample is taken by swab and placed at the start of the testing strip, in addition with rabbit IgG which is used as a control (Wang et al., 2020). Newer LFA utilizes a hybrid capture fluorescence immunoassay technique which begins with the lysing and release of SARS-COV-2 from the throat swab (Wang et al., 2020). This technique increases the processing time to one hour, however, greatly increases the sensitivity as compared to previous COVID-19 rapid LFA test (Wang et al., 2020). The SARS-COV-2 RNA hybridizes with a designed DNA probe to create a RNA-DNA hybrid which is then captured by fluorescent nanoparticle(FNP)-labelled monoclonal antibodies (Wang et al., 2020). As the sample travels down the strip due to capillary action and will reach a Test (T) line which is coated with S9.6 monoclonal antibodies (Wang et al., 2020). The S9.6 monoclonal antibody captures the SARS-COV-2:FNP-labelled monoclonal antibody complex and leads to fluorescence of the line (Wang et al., 2020). Further down the strip is also a Control (C) line with coated anti-rabbit IgG antibodies (Wang et al., 2020). Used as a control, the line should fluoresce in every use as rabbit IgG is added along with throat samples (Wang et al 2020). A positive test will be indicated by presence of two fluorescent lines (T and C) while a negative test will be indicated by the presence of one line (just C) (Wang et al., 2020). Illumination of the T line is judged by a fluorescent-intensity cut off value (Wang et al., 2020).

Figure 4: Steps in COVID-19 specific antigen Lateral Flow fluorescence immunoassay. Adapted from Wang et al. (2020).

Figure 5: Potential results of lateral flow strip. Adapted from Wang et al. (2020).

The main benefit of a Lateral Flow Immunoassay is the time to produce results, as it can be as short as 15 mins (Augustine et al., 2020). The main source of increased processing time comes from the lack of amplification required as compared to RT-PCR (Wang et al., 2020). Additionally, the LFA can be used at Point of Care, and thus, does not require sophisticated laboratory machinery and expertise (Augustine et al., 2020). Thereby, also greatly lowering the cost of administration. This technology is especially beneficial for impoverished nations where facilities lack the funds to administer RT-PCR testing (Augustine et al., 2020).

Speed of processing comes to the cost of accuracy, as the main downside of rapid LFA testing is the reduced specificity and sensitivity compared to RT-PCR. Albeit, the gap is closing as new technologies and strategies to improve accuracy are discovered (Wang et al., 2020).

First, a blood sample is taken from the patient and is diluted. Once placed at the start of the strip, the sample will travel down the strip due to capillary action (Augustine et al., 2020). The sample will then hit a conjugation pad, where a conjugate will bind to SARS-COV-2 specific antibodies to create an antibody:conjugate complex (Augustine et al., 2020). Further down the strip are antibody isotype testing lines (Augustine et al., 2020). Similar to ELISA tests, LFA typically tests for 2 different isotypes to increase the specificity and sensitivity. These testing lines contain anti-IgM and anti-IgG antibodies to capture conjugated SARS-COV-2 specific IgM and IgG (Augustine et al., 2020). Additionally, there is a third Control line which is coated by anti-rabbit IgG and binds to rabbit IgG which is supplemented on the conjugation pad and carried forward by capillary action (Augustine et al., 2020). The control line should always be coloured to indicate proper function of the test strip (Augustine et al., 2020). A positive test is indicated by the presence of at least one out of two coloured isotype antibody testing lines and control line (Augustine et al., 2020). A negative test is indicated by the presence of only the coloured control line (Augustine et al., 2020).

Figure 6: Steps in COVID-19 specific Lateral Flow Assay. Adapted from Augustine et al. (2020).

The advantages of LFA antibody test are the same as the advantages for LFA antigen tests, mainly the increased processing time, lowered amount of reagents to allow for point of care testing and subsequently being a cheaper and easier to use alternative (Augustine et al., 2020).

LFA capturing antibodies is simpler than LFA capturing antigens. However, this simplicity leads to a lower specificity and sensitivity as compared to ELISA antibody testing as there is no technique to circumvent lowered sample amount and lack of incubation time (Hackner et al., 2020). Thus, the test has lower sensitivity and specificity when compared to an ELISA antibody test which is the gold standard for serological antibody testing (Serrano et al., 2020).

Figure 7: Presence of bilateral ground-glass opacities in upper lobe. Adapted from Bernheim et al. (2020).

The use of CT scans was the initial diagnostic tested used in Wuhan China in the advent of the COVID-19 pandemic, when PCR and ELISA test specific to SARS-COV-2 had not yet been developed. The diagnosis of COVID-19 through CT scans is discerned through observation of bilateral, peripheral ground-glass and consolidative pulmonary opacities (Bernheim et al., 2020). However, further advancement in illness can result in the observation of linear opacities, greater lung involvement and special patterns referred to as “crazy-paving” and “reverse halo” pattern (Bernheim et al., 2020). CT scans are reliant on physicians’ ability to assess the scans. It has been documented that physicians have a high specificity when it comes to diagnosis of COVID-19 through CT scans however they have a low sensitivity (Bai et al., 2020). The low sensitivity stems from the fact that many patients who have contracted COVID-19 appear to have normal CT scans as most cases are mild, and thus, the rate of false negatives is higher (Bai et al., 2020). The test is based more on qualitative results rather than quantitative results seen in other tests with strict cut off values.

CT scans present quick results in order to detect the presence of COVID-19, specifically in more severe cases (Bai et al., 2020).

CT scans are limited due to their low sensitivity and specificity when compared to gold standard diagnostics such as RT-PCR (Bai et al., 2020). Additionally, CT scans are expensive and require specialists to administer and diagnose (Bai et al., 2020).

AI technology is being utilized to enhance the diagnosis of COVID-19 through CT scans (Zhang et al., 2020). The AI is taught by feeding the system CT scans from known positive and negative patients as well as CT scans from other common lung disorders to help differentiate (Zhang et al., 2020). As noted, COVID-19 diagnosis based on CT scans have a low sensitivity and thus the AI is able to enhance physician decision by indicating important critical markers to improve accuracy of diagnosis (Zhang et al., 2020). Additionally, the AI is able to predict the progression of illness and assist in evaluating drugs for effective treatment (Zhang et al., 2020). Overall, AI is a quick and useful tool for diagnosis of COVID-19, especially when the healthcare system is overloaded, while also being a good tool to enhance the accuracy of CT scan diagnosis (Zhang et al., 2020). More studies need to be conducted in order to identify the usefulness and accuracy of AI, as currently the technology is still in its infancy (Chen & See, 2020).

Figure 8: SARS-CoV-2 DETECTR workflow. Conventional RNA extraction can be used as an input to DETECTR which is then visualized by a fluorescent reader or lateral flow strip (Broughton et al., 2020).

CRISPR–Cas12-based assays can be for the detection of SARS-CoV-2 from extracted sample RNA. One detection method using this type of assay is called SARS-CoV-2 DNA Endonuclease-Targeted CRISPR Trans Reporter (DETECTR) (Broughton et al., 2020). In the first step of this method, the samples are collected from the patients. The most general forms of the SARS- CoV-2 samples are nasopharyngeal and oropharyngeal swabs (Nouri et al., 2021). After sample collection, viral RNAs need to be extracted from the raw sample. RNA isolation procedures typically involve three general steps: lysis, separation of RNA from other macromolecules such as DNA, proteins, and lipids, followed by RNA elution (Nouri et al., 2021). This isolation can be performed by magnetic bead purification, spin column isolation, or organic extraction which is considered the best method for this situation. Once isolation is completed, the isolated sample is amplified using methods such as RT-PCR or RT-LAMP to boost the amount of RNA past the limit of detection (Nouri et al., 2021). Once simultaneous reverse transcription and isothermal amplification using RT–LAMP is performed, the sample is placed in the DETECTR system where Cas12 detection of predefined coronavirus sequences occurs (Broughton et al., 2020). Then a reporter molecule is cleaved to confirm the detection of the virus (Broughton et al., 2020). The extracted RNA from a sample is placed in the DETECTR system where LAMP preamplification and Cas12-based detection for envelope (E) gene, nucleoprotein (N) gene and RNase Polymerase occurs, which is visualized by a fluorescent reader or lateral flow strip (Broughton et al., 2020).

Primers that target the envelope (E) and nucleoprotein (N) genes of SARS-CoV-2 were used by this method and are similar to the ones used in the WHO assay and US CDC assay are utilized with slight modifications to be optimally used in the DETECTR system (Broughton et al., 2020). The N1 and N3 regions used by the US CDC assay weren’t used, as these regions did not have compatible protospacer adjacent motif sites for the Cas12 guide RNAs (gRNAs) used to detect 3 sars like coronaviruses in the E gene and only SARS-CoV-2 in the N gene (Broughton et al., 2020).

A key advantage of the DETECTR system over the CDC qRT–PCR assay is that the DETECTR system is capable of getting its result at a rate that is 400% faster than the qRT-PCR assay (Broughton et al., 2020). Other key advantages of the DETECTR system over the CDC qRT–PCR assay include isothermal signal amplification without needing thermocycling, rapid turnaround time, single nucleotide target specificity, integration with accessible and easy-to-use reporting formats such as lateral flow strips, and no requirement for complex laboratory infrastructure (Broughton et al., 2020). Using synthetically derived SARS-CoV-2 RNA, DETECTR is also able to distinguish SARS-CoV-2 with no cross-reactivity to other strains of coronavirus when the N gene gRNA (Broughton et al., 2020).

When compared to other assays, the estimated limit of detection for the CDC qRT–PCR assay tested by the California Department of Public Health is 1 copy per µl reaction, versus 10 copies per µl reaction for the DETECTR assay making it more likely for this system to produce a false negative (Broughton et al., 2020).

Figure 9: P-BEST design and detection results obtained for a set of 384 samples with a carrier rate of ~1% (Shental et al., 2020).

P-Best uses a single-step group testing method to test multiple people for SARS-CoV-2 at a much faster rate (Shental et al., 2020). Instead of testing each sample separately, samples are pooled into groups in the P-Best system, and each pool is tested for SARS-CoV-2 using a PCR-based diagnostic assay. Each individual’s sample is a part of multiple groups in a combinatorial grouping method to determine the specific positive individual from a positive group test result without the need for additional testing (Shental et al., 2020).

The ratio of the number of samples to the number of pools is affected by the positive carrier rate of the population. As the carrier rate increases, the efficiency is reduced, since more pools are required to correctly identify all positive subjects (Shental et al., 2020). When the number of carriers tested is around 1% (ideal), this system can correctly identify all positive individuals at an 800% decrease in the number of diagnostic tests needed when compared to testing each individual sample separately (Shental et al., 2020). Since pooling a positive sample with additional negative samples will result in dilution of the viral RNA concentration within the pool which can lead to a loss of sensitivity (Shental et al., 2020). The effect of pool size to the ability of detecting SARS-CoV-2 was tested using many pools of varying size up to 48 samples in a pool with one sample being positive for SARS-CoV-2. All these varied pools were found to be positive for SARS-CoV-2 when tested using other diagnostic kits (Seegene COVID-19 diagnostic kit) but lower yields of viral RNA were detected when using larger pools (Shental et al., 2020). With these results, the P-Best system has been shown to be proficient in detecting SARS-CoV-2 with a 0.71% rate of false positives and a 0.09% rate of false negatives under ideal conditions (Shental et al., 2020).

The current implementation of P-BEST is designed for a carrier rate of about 1% (Shental et al., 2020). When the carrier rate of a population is higher than this ideal value, the efficiency of the system changes and can lead to errors (Shental et al., 2020). At higher carrier rates too many pools would be PCR positive, and the number of samples identified by the method will be much larger than the number of expected carriers. This will prevent direct identification of the actual positive carriers in a single testing round and the user will have to retest the samples from the positive pools either individually or by using an alternate pooling design (Shental et al., 2020).

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