Description
Human coronaviruses, first characterized in the 1960s, are named based on the spiked proteins located on their surface.  As of 2020, seven coronaviruses are known to infect humans. Four, of which — 229E, NL63, OC43, and HKU1 — are associated with the common cold. MERS-CoV is the coronavirus that causes Middle East Respiratory Syndrome, or MERS. SARS-CoV is the causative agent of Severe Acute Respiratory Syndrome (SARS), and SARS-CoV-2 is the virus that causes coronavirus disease 2019, or COVID-19 (CDC, 2020b). As of November 11, 2022, the United States had reported more than 634,709,332 confirmed cases of COVID-19 and over 6,609,119 reported COVID-19 deaths (JHU, 2022). Testing for a possible coronavirus infection can include molecular tests, such as nucleic acid-based testing like reverse transcription polymerase chain reaction (RT-PCR); host antibody testing; and antigen testing.

Policy

1. Targeted nucleic acid testing, such as RT-PCR, for COVID-19 (SARS-CoV-2), including rapid molecular tests, is considered MEDICALLY NECESSARY in the following situations:
   1. For individuals displaying signs and symptoms of possible COVID-19 infection (See Note 1)
   2. For asymptomatic individuals with known exposure to COVID-19, EXCEPT when the individual has had a previous COVID-19 infection within the last 90 days
   3. For asymptomatic individuals prior to undergoing immunosuppressive or aerosol-producing procedures
2. For individuals with signs or symptoms of SARS who have traveled to endemic areas or who have been exposed to persons with SARS, targeted nucleic acid testing, such as RT-PCR, for the detection of severe acute respiratory syndrome (SARS) coronavirus RNA is considered MEDICALLY NECESSARY. 
3. For individuals with signs or symptoms of Middle East respiratory syndrome (MERS) who have traveled to endemic areas or who have been exposed to persons with MERS, targeted nucleic acid testing, such as RT-PCR, for the detection of MERS coronavirus RNA is considered MEDICALLY NECESSARY. 
4. Host antibody serology testing to support a diagnosis of multisystem inflammatory syndrome in children (MIS-C) (see Note 2), multisystem inflammatory syndrome in adults (MIS-A) (see Note 3), or post-acute sequelae of SARS-CoV-2 infection (PASC) is considered MEDICALLY NECESSARY.
5. For symptomatic individuals, the use of an antigen-detecting diagnostic test for SARS-CoV-2, including antigen rapid tests, is considered MEDICALLY NECESSARY. 
6. For individuals with signs and symptoms of a respiratory tract infection (see Note 4), multiplex PCR-based panel testing of up to 5 respiratory pathogens is considered MEDICALLY NECESSARY.
7. For individuals with signs and symptoms of a respiratory tract infection (see Note 4), antigen panel testing of up to 5 antigens is considered MEDICALLY NECESSARY.
8. For the diagnosis of SARS-CoV-2 reinfection, whole genome sequencing of paired specimens from distinct lineages (as defined in Nextstrain or GISAID) is considered NOT MEDICALLY NECESSARY.
9. Antigen panel testing of 6 or more antigens is considered NOT MEDICALLY NECESSARY. 
10. Multiplex PCR-based panel testing of 6 or more respiratory pathogens is considered NOT MEDICALLY NECESSARY. 
11. For all other situations not described above, host antibody serology testing is considered NOT MEDICALLY NECESSARY.

The following does not meet coverage criteria due to a lack of available published scientific literature confirming that the test(s) is/are required and beneficial for the diagnosis and treatment of a patient’s illness.

12. For all situations, neutralization antibody testing for SARS-CoV-2 is considered NOT MEDICALLY NECESSARY.
13. Testing for other endemic coronaviruses, such as 229E, NL63, OC43, and HKU1, is considered NOT MEDICALLY NECESSARY.

NOTES:

Note 1: Signs and symptoms associated with a possible COVID-19 infection can include a fever, cough, fatigue, shortness of breath or difficulty breathing, congestion or runny nose, chills, muscle pain, headache, sore throat, new loss of taste or smell, nausea, vomiting, diarrhea, conjunctivitis, rash on skin or discoloration of fingers or toes (CDC, 2020f; WHO, 2020d).

Note 2: According to the CDC, evidence of possible MIS-C includes (CDC, 2021d):

• Fever of at least 38.0˙C for at least 24 hours
• Multisystem (2 or more) organ involvement
• Laboratory evidence of inflammation, “including, but not limited to, one or more of the following: an elevated C-reactive protein (CRP), erythrocyte sedimentation rate (ESR), fibrinogen, procalcitonin, d-dimer, ferritin, lactic acid dehydrogenase (LDH), d-dimer, ferritin, lactic acid dehydrogenase (LDH), or interleukin 6 (IL-6), elevated neutrophils, reduced lymphocytes and low albumin (CDC, 2020d)”

Some children may fulfill full or partial criteria for Kawasaki disease

Note 3: According to the CDC, evidence of possible MIS-A includes (Morris et al., 2020; Patel et al., 2021):

• A severe illness requiring hospitalization in a person aged ≥ 21 years; 
• A positive test result for current or previous SARS-CoV-2 infection (nucleic acid, antigen, or antibody) during admission or in the previous 12 weeks;
• Severe dysfunction of one or more extrapulmonary organ systems (e.g., hypotension or shock, cardiac dysfunction, arterial or venous thrombosis or thromboembolism, or acute liver injury); 
• Laboratory evidence of severe inflammation (e.g., elevated CRP, ferritin, D-dimer, or interleukin-6);
• Absence of severe respiratory illness (to exclude patients in which inflammation and organ dysfunction might be attributable simply to tissue hypoxia).

Note 4: Signs and symptoms of a respiratory tract infection:

• A temperature greater than 102°F  
• Pronounced dyspnea
• Tachypnea
• Tachycardia

Rationale
On March 11, 2020, the World Health Organization (WHO) declared the novel coronavirus SARS-CoV-2, or COVID-19, a global pandemic (Cucinotta & Vanelli, 2020). COVID-19 is the third recent human coronavirus to be declared an emergency. SARS (Severe Acute Respiratory Syndrome) was recognized as an emergency by the WHO in February 2003 (WHO, 2022c). This outbreak in 2003 resulted in over 8000 cases in 26 different countries.  Since 2003, only four limited reoccurrences have been reported according to the WHO—three incidences are due to laboratory accidents (in Taipei and Singapore) and one incident of undetermined source in China (WHO, 2022c). As early as September 2012, another human coronavirus, MERS-CoV, began to spread in the Middle East, causing Middle East Respiratory Syndrome (MERS). Although the WHO did not initially declare MERS an emergency, they have since added MERS to their list of pandemic/epidemic diseases. Since September 2012 and as of the end of October 2021, the WHO reports 2574 laboratory-confirmed cases of MERS with 858 MERS-associated deaths (34.4% fatality rate) in 27 countries (WHO, 2022b). 

Unlike the initial SARS and MERS outbreaks that were predominantly regionally contained, COVID-19 became a global pandemic. According to the WHO, as of November 11,2022, there were more than 630 million confirmed cases of COVID-19 with over 6,584,104 confirmed deaths worldwide (WHO, 2022a). Infection from the novel human coronavirus SARS-CoV-2 can result in coronavirus disease 2019 (COVID-19). The WHO reports approximately 15% of individuals with COVID-19 develop severe disease requiring oxygen support while 5% develop “critical disease” with complications such as respiratory failure or multiorgan failure (WHO, 2021b). Older individuals and patients with comorbidities — such as cardiovascular disease, diabetes mellitus, hypertension, chronic lung disease, cancer, chronic kidney disease, obesity, and smoking — have an increased likelihood of poor outcomes (McIntosh, 2021). Sepsis, multiorgan failure (including the kidney, liver, and heart), pneumonia, and acute respiratory distress syndrome (ARDS) can also occur (WHO, 2021b; Yang et al., 2020). Severe outcomes have been associated with the following laboratory features: lymphopenia, elevated liver enzymes, elevated lactate dehydrogenase (LDH), elevated inflammatory markers (such as CRP and ferritin), elevated D-dimer, elevated prothrombin time (PT), elevated troponin, elevated creatine phosphokinase (CPK), and acute kidney injury (McIntosh, 2021). 

Much of what has generated this global pandemic is attributed to the different levels of transmissibility of the SARS-CoV-2 virus compared to SARS-CoV-1 and MERS, which can arise from the viral load. Simply put, viral load is the number of viral particles/virions in a milliliter of blood (Ryding, 2020). The viral load of SARS-CoV-2 “peaks around the time of symptom onset, followed by a gradual decrease to a low level after about 10 days. Regarding the period of high infectiousness, a recent study reported that exposure to an index case within 5 days of symptom onset confers a high risk of secondary transmission” (Kawasuji et al., 2020). This finding was corroborated by other studies, which found that “SARS-CoV-2 viral load in the upper respiratory tract appeared to peak in the first week of illness, whereas that of SARS-CoV peaked at days 10 – 14 and that of MERS-CoV peaked at days 7 – 10;” because SARS-CoV-2 viral load peaks faster, it can be more transmissible earlier in the disease course (Cevik et al., 2021). However, after reaching its peak during symptom onset, the viral load decreases “monotonically” (Kawasuji et al., 2020). If viral loads do not decrease, patients will be more likely to suffer worse outcomes and require hospitalization (Griffin, 2020). Viral load has been found to be either similar among symptomatic and asymptomatic COVID-19 positive individuals, or higher among symptomatic individuals (Kawasuji et al., 2020; Nackerdien, 2020). Infectiousness of COVID-19 also correlates with shedding, meaning that the viral particles can replicate in an individual and spread in the environment to others. The mean duration of SARS-CoV-2 RNA shedding “was 17.0 days (95% CI 15·5 – 18·6; 43 studies, 3229 individuals) in upper respiratory tract, 14.6 days (9·3 – 20·0; seven studies, 260 individuals) in lower respiratory tract, 17.2 days (14·4 – 20·1; 13 studies, 586 individuals) in stool, and 16.6 days (3·6 – 29·7; two studies, 108 individuals) in serum samples,” with maximum shedding duration reaching “83 days in the upper respiratory tract, 59 days in the lower respiratory tract, 126 days in stools, and 60 days in serum”(Cevik et al., 2021). 

In children and adolescents, reports of a multisystem inflammatory syndrome (MIS-C) with similarities to Kawasaki disease and toxic shock syndrome have been linked to COVID-19 (CDC, 2020d; DeBiasi et al., 2020; Jones et al., 2020; Verdoni et al., 2020; WHO, 2020c). Multisystem inflammatory syndrome has also been reported in adults (MIS-A). From June to October 2020, researchers reported 27 cases of MIS-A in the US and UK (Baum, 2020). The case definition of MIS-A includes “(1) hospitalization without evidence of severe respiratory illness (to exclude hypoxia as the cause of the signs and symptoms), (2) extrapulmonary organ system involvement (including hypotension or shock, cardiac dysfunction, arterial or venous thromboembolism, acute liver injury, or dermatologic abnormalities), and (3) laboratory evidence of acute inflammation (e.g., highly elevated C-reactive protein, ferritin, D-dimer, or interleukin-6)” (Baum, 2020). Most patients present with a fever > 100.4 °F, cardiac abnormalities (arrhythmias, elevated troponin levels, or left or right ventricular dysfunction), and gastrointestinal symptoms. Rare symptoms include dermatological manifestations or respiratory symptoms such as pleural effusion. Patients may have elevated laboratory markers of inflammation including CRP, ferritin, and markers of coagulopathy including D-dimer (Morris et al., 2020). 
On September 20, 2020, the first case of a new COVID-19 variant, B.1.1.7 (Alpha) or VUI-202012/01 (VUI for “variant under investigation”) was recorded in the UK. As of August 11, 2021, the Beta and Gamma variants are not classified as “variants of concern” (CDC, 2022r). The Gamma variant is now considered a “variant being monitored” or VBM (CDC, 2022c). 

The B.1.617.2 (Delta) variant was first detected in the U.S. in March 2021, but had its first case in India in December 2020 (CDC, 2022r). The delta variant is classified as a “variant of concern.” Individuals with the delta variant also have viral loads “up to 1,260 times higher” than compared to those infected by the original virus, and as of July 2021, has been detected in more than 130 countries (GVN, 2021). It is also more contagious than other illnesses such as “MERS, SARS, Ebola, the common cold, the seasonal flu, and smallpox, and it is as contagious as chickenpox” (Mandavilli, 2021). It has been associated with different symptoms than other strains; “fever, headache, sore throat, and runny nose are common, while cough and loss of smell are not. Other reports link Delta to more serious symptoms, including hearing impairment, severe gastrointestinal issues and blood clots leading to tissue death and gangrene” (ASM, 2021). However, between August 7, 2022 and October 22, 2022, the Delta variant accounted for 0.0% of COVID-19 cases; Omicron accounted for the majority of cases (CDC, 2022c). 

B.1.1.529 (Omicron) variant was first reported to WHO in South Africa on November 24, 2021 and on December 1, 2021, the first case of Omicron was confirmed in the US (CDC, 2022l). Omicron variant is comprised of four different lineages, which include BA 1.1, B.1.1.529, BA.2, BA.2.12.1, and BA.3 (CDC, 2022k, 2022p). The Omicron variant was classified as a “variant of concern” (CDC, 2022c). At first, Omicron variant was detected in travel-related cases in several European countries, as well as Australia, Brazil, Canada, Hong Kong, Israel, Japan, Nigeria, Norway, Sweden, and the United Kingdom. However, cases doubled every 2 – 3 days in the UK thereafter and rose from 1% to 99% of infections nationally in 6 weeks, compared with 18 weeks for Delta. Omicron may be over 10 times more contagious than the original virus or about 2.8 times as infectious as the Delta variant due to mutations in the spike protein (CDC, 2022f). Symptoms are similar to previous variants but has shown to cause less severe disease which depends on COVID-19 vaccination status, the presence of other health conditions, age, and history of prior infection (CDC, 2022l). According to Our World in Data, by November 11, 2022, 68.2% of the US population had received at least one dose of a COVID-19 vaccine (Mathieu et al., 2022). 

The CDC has listed the Pfizer-BioNTech COVID-19 Vaccine, Moderna COVID-19 Vaccine, and Johnson & Johnson’s (J&J) Janssen COVID-19 Vaccine as the only three authorized and recommended vaccines to prevent COVID-19 in the US (CDC, 2022d). The Pfizer-BioNTech and Moderna COVID-19 vaccines are mRNA vaccines, which instruct B and T lymphocytes to fight off that specific mRNA-encoded protein from COVID-19 in the event of future exposure. They both require two doses to obtain optimal effectiveness (CDC, 2022o). After the two doses, the Pfizer vaccine and Moderna vaccine are 95% and 94.1% effective, respectively, in preventing COVID-19 (Branswell, 2020). The Johnson & Johnson/Janssen COVID-19 vaccine is a viral vector vaccine, which uses a “modified version of a different virus (the vector) to deliver important instructions to our cells” (CDC, 2022q). The Janssen COVID-19 vaccine only requires one dose, and in clinical trials, the vaccine was 66.3% effective at “preventing laboratory-confirmed COVID-19 illness in people who had no evidence of prior infection 2 weeks after receiving the vaccine” (CDC, 2021c).  For primary and booster vaccination, CDC notes a preference for mRNA vaccines (Pfizer-BioNTech or Moderna) over the Janssen COVID-19 Vaccine. However, the Janssen vaccine is preferable in instances where there are no other vaccines available (CDC, 2022d). 

As of August 31, 2022, there are 42 vaccines in Phase 3 trials and 10 in combination phase 2/3 trials (Zimmer et al., 2022). AstraZeneca and Novavax are approved for usage in the EU (AstraZeneca, 2021; Zimmer et al., 2022). AstraZeneca is a vector vaccine, which contains weakened versions of a virus that has the same entryway as COVID-19 into a cell, and Novavax is a protein subunit vaccine, which contains stabilized but immunogenic pieces of the SARS-CoV-2 virus, namely the spike protein (AstraZeneca, 2021; CDC, 2022o; NIH, 2020). AstraZeneca and Novavax COVID-19 vaccines require two doses (AstraZeneca, 2021; Roberts, 2021). The AstraZeneca and Novavax COVID-19 vaccines have been found to be up to 90% effective in UK trials (Roberts, 2021; Van Beusekom, 2020). 

With the emergency of the Omicron variant in November 2021, further research suggested that vaccine effectiveness at preventing infection wanes over time, especially in those aged 65 years and older. Data from clinical trials showed that a booster shot increased the immune response in trial participants who finished a Pfizer-BioNTech or Moderna primary series 6 months earlier or who received a J&J/Janssen single-dose vaccine 2 months earlier. For Pfizer-BioNTech and J&J/Janssen, clinical trials also showed that a booster shot helped prevent severe symptoms of COVID-19 (CDC, 2022b). 

Vaccines in clinical trials have focused on targeting the spike (S) protein (based on experience with SARS-CoV-1), which contains a receptor binding domain (RBD) responsible for allowing entrance into host cells and is the current target for neutralizing antibodies. Other targets could potentially include the nucleocapsid (N) protein, likely for inactivated virus or live attenuated approaches, or T cell epitopes, which may provide additional protection. Knowing that T-cell responses against structural proteins in SARS-CoV-1 were more immunogenic than non-structural proteins could be key in manufacturing more effective vaccines for SARS-CoV-2 as well (Tregoning et al., 2020).

Besides the viruses associated with SARS, MERS, and COVID-19, four other human coronaviruses (HCoVs) are currently known — 229E, NL63, OC43, and HKU1. These four viruses are considered endemic to the human population, and they typically cause mild respiratory tract infections associated with the common cold; in fact, it is approximated that up to one-third of all “common colds” may be due to one of these four endemic human coronaviruses. These HCoVs can cause both upper and lower respiratory infections, but they typically result in relatively mild, or even asymptomatic, cases. In immunosuppressed individuals, including those with pre-existing pulmonary diseases, progression to acute respiratory failure can occur in some cases (Corman et al., 2019; Ludwig & Zarbock, 2020). 

Nucleic Acid Testing for Human Coronavirus Infections
Coronaviruses are a family of enveloped, single-stranded positive-sense RNA viruses. During the initial phase of infection, the virus can be detected in respiratory specimen due to high concentrations of viral RNA (Figure 1). RT-PCR is a powerful molecular technique that synthesizes complimentary DNA (cDNA) from the initial RNA template and uses primers to manufacture multiple cDNA copies for analysis. RT-PCR, when used with appropriate primers targeting the SARS-CoV-2 RNA, is used to diagnose an acute infection. The CDC RT-PCR Diagnostic Panel detects SARS-CoV-2 virus in the upper and lower respiratory specimen. The CDC has released standard primers to detect SARS-CoV-2 RNA, but any primers or probes that receive an Emergency Use Authorization (EUA) label may also be used with the CDC’s RT-PCR Diagnostic Panel (CDC, 2020e). As depicted in Figure 1, the concentration of viral RNA decreases as the immune system fights the infection, and very low or undetectable viral RNA levels are typically present after an individual has recovered. Consequently, RT-PCR cannot be used to screen for a past infection. Another limitation to RT-PCR is that it does require specific instrumentation, and, therefore, is less amenable as a rapid, point-of-care test. RT-PCR results of SARS-CoV-2 may fluctuate and become unstable over time, thus requiring other clinical diagnostic measures, such as computerized tomography (CT imaging) to supplement isolation, discharge, and any transfers during this epi demic (Li et al., 2020).

Clinical Utility and Validity of Nucleic Acid Testing
Many studies have been performed to date to evaluate the analytical performance of RT-PCR. One study, using a high-throughput platform, for example, reported a limit of detection (LoD) of 689.3 copies/mL and 275.72 copies per reaction at 95% detection probability (Pfefferle et al., 2020). The WHO diagnostic RT-PCR test utilizes two genes--the E gene as the molecular target (where the limit is 3.9 copies per reaction) and the RdRp gene as the molecular target (limit of 3.6 copies per reaction) (Lippi et al., 2020). One recent study reported possible in vitro cross-reactivity between the RdRp-based method used predominantly in European labs with SARS-CoV in cell culture (Chan et al., 2020). SARS-CoV is the coronavirus that caused the initial SARS (Severe Acute Respiratory Syndrome) outbreak in 2003 (WHO, 2022c). The likelihood of either a co-infection of SARS-CoV and SARS-CoV-2 or a concurrent outbreak of both viruses is small. The CDC diagnostic panel test does not target the RdRp gene; it consists of two primer/probe sets of the N gene and one primer/probe set for human RNase P gene (RP) as the control.  The CDC diagnostic panel has a reported limit of 1.0 – 3.2 copies/µL (CDC, 2020a; Lippi et al., 2020). Reports of initial negative RT-PCR results in individuals who later develop symptomatic COVID-19 have been published, but this may occur if the sample was not properly collected or if it was taken from the patient early in the infection during the initial incubation period of SARS-CoV-2, which is approximately 6 days (interquartile range [IQR], 2 – 11 days) (Backer et al., 2020; Lippi et al., 2020). Consequently, it is important to remember that “Negative results do not preclude SARS-CoV-2 infection and should not be used as the sole basis for patient management decisions. Negative results must be combined with clinical observations, patient history, and epidemiological information (LabCorp, 2020b).”

To compare and analyze the diagnostic efficacy of two RT-PCR test kits for detection of SARS-CoV-2, Lu et al. (2020) studied throat swab samples from 18 hospitalized patients with a clinical COVID-19 diagnosis and 100 hospitalized patients without COVID-19 diagnosis. Two different RT-PCR tests from Sansure Biotech Inc (SansureBiotech, 2020) and Shanghai BioGerm Biotechnology Co., Ltd (BioGerm, 2020) were used. The table below (Lu et al., 2020) shows that the detection efficacy of BioGerm PCR kit was higher than that of the Sansure PCR kit. These two kits had the same specificity and positive predictive value, but the sensitivity of the Sansure PCR kit was 83.3%, whereas the sensitivity of the BioGerm PCR kit was 94.4%. For the Sansure PCR kit, 3 of the 18 samples were false-negative results, and for the BioGerm PCR kit, 1 of the 18 samples was a false-negative result. No false-positive results were detected in these tests. The author suggests that “these findings provide important information for the ongoing optimization of viral detection assays following the emergence of COVID-19” (Lu et al., 2020).

	COVID‐19 samples (n = 18)		None‐ COVID‐19 samples (n = 100)		Sensitivity (95%CI)	Specificity (95%CI)	PPV (95%CI)	NPV (95%CI)	Kappa (95%CI)
Test kits	Positive	Negative	Positive	Negative
Sansure	15	3	0	100	0.833(0.577‐0.956)	1.000(0.954‐1.000)	1.000(0.747‐1.000)	0.971(0.911‐0.992)	0.894(0.726‐1.000)
BioGerm	17	1	0	100	0.944(0.706‐0.997)	1.000(0.954‐1.000)	1.000(0.771‐1.000)	0.990(0.938‐0.999)	0.966(0.880‐1.000)

 

 

 

 

 

 

 

 

Table 2. Diagnosis efficacy of Sansure and BioGerm test kits for SARS‐CoV‐2 nucleic acid detection

In a case series study of multisystem inflammatory syndrome in adults (MIS-A) associated with SARS-CoV-2 infection, 16 patients ranging from 21 to 50 years old were enrolled and tested with PCR assay. 10 out of 16 patients had positive SARS-CoV-2 PCR test results at the time of admission. Two patients had positive SARS-CoV-2 PCR test results 14 and 37 days before admission and negative PCR results at the time of admission. Three patients had positive SARS-CoV-2 PCR test results 25 – 41 days before admission and continued positive PCR test results at the time of admission. “Given the high proportion of MIS-C patients with negative PCR testing, clinical guidelines recommend the use of both antibody and viral testing to assist with diagnosis” (Morris et al., 2020).

Li et al. (2021) conducted a cross-sectional analysis on 30 patients with COVID-19 diagnoses to compare the sensitivity of SARS-CoV-2 testing in anterior nasal vestibular swabs versus oropharyngeal swabs. After specimen collection, RT-PCR assays were used to test them for SARS-CoV-2. They found that 56.7% of the patients tested positive using oropharyngeal specimen, whereas 66.7% of patients tested positive with the nasal swab specimens. Ultimately, there is “adequate sensitivity” to use the less invasive anterior nasal vestibular swabs to detect COVID-19 infection confirmed by RT-PCR (Li et al., 2021).

Yau et al. (2021) evaluated the clinical utility of a rapid “on-demand” PCR-based testing service in an acute hospital setting. In an effort to increase hospital efficiency starting from July 2020, the researchers focused on moving patients quickly to isolation rooms and minimize potential risk of transmission in crowded areas. From their study, it was found that the “daily/monthly PCR positive test numbers approximately followed the local and national UK trend in COVID-19 case numbers, with the daily case numbers being reflective of the Nov and Dec 2020 surges.” It ultimately helped to reduce “unnecessary ‘length-of-stay’ in a busy acute respiratory ward.” Patients were able to be rapidly separated based on COVID-19 positive diagnosis and the system in place reduced exposure and nosocomial transmission (Yau et al., 2021).

Dighe studied a lateral flow strip-based RNA extraction and amplification-free nucleic acid test (NAT) for rapid diagnosis of COVID-19 at point of care which takes no longer than 30 minutes. This test uses highly specific 6-carboxyfluorescein (6-FAM) and biotin labeled antisense oligonucleotides (ASOs) as probes those are designed to target the N-gene sequence of COVID-19. This study evaluated 60 samples using the lateral flow assay and results were compared with the FDA-approved TaqPath RT-PCR kit. According to the results, the assay obtained almost 99.99% accuracy and specificity. The authors conclude that this new LFA method could be "expanded beyond COVID-19 detection, simply by altering its targeting antisense oligonucleotides, to become a global health technology that contributes to providing low-cost diagnostics" (Dighe et al., 2022).

Mawhorter et al. (2022) investigated the impact and cost of a routine pre-operative COVID-19 PCR testing algorithm for asymptomatic patients before elective surgery at a rural academic institution per recommendations by the American College of Surgeons. From 7579 pre-procedural tests that were completed since May 2020 using the protocol, the study yielded 31 (0.41%) positive results in asymptomatic patients. With these positive results, there were impacts on both the cost and delay of the procedure. The results showed that “20 procedures (62.5%) were delayed an average of 49 days, 8 were not performed, and 3 proceeded without delay,” with a prolonged delay for the 3 urological procedures of 59 days. They also identified that the number needed to test for one positive result was 244, with $11,573 as cost for each positive result. This analysis found that the hospital was able to be more cost-effective (each test was $34-54) with a standardized testing algorithm prior to procedure performance (Mawhorter et al., 2022). 

Host Antibody Testing
The COVID-19 illness begins with an initial infection by SARS-CoV-2. Viral invasion stimulates the host immune response to produce immunoglobulins, such as IgM, IgA, and IgG, that can target the invading virus. However, there is a delay between the time of initial infection and the production of immunoglobulins (Figure 1) (The_Native_Antigen_Company, 2020). Typically, several days after the initial onset of symptoms, the first IgM immunoglobulins are produced to combat the viral infection. IgA (not shown in Figure 1), immunoglobulins secreted to protect predominantly the mucosal linings of the gastrointestinal, respiratory, and genitourinary tracts (Woof & Kerr, 2006), typically have a half-life of 4 – 6 days (Morell et al., 1973). Finally, IgG, the long-term immunoglobulins found within body fluids that fight bacterial and viral infections, are produced and IgM production wanes. Some limited studies have indicated that some individuals may initially produce IgM and IgG antibodies concurrently, but additional research is needed (Padoan et al., 2020). 

Serological host antibody tests can detect the presence of IgM and IgG antibodies that an individual has developed in response to an infection—in this case, a SARS-CoV-2 viral infection. The test may report total antibodies present, meaning either it does not distinguish between IgG and IgM or that it is reporting the sum of IgG and IgM.  This is sometimes referred to as “total antibody testing”. On the other hand, the test may be specific for one antibody, such as IgG or IgM, or the test may claim to accurately distinguish between the antibodies. 

Another type of antibody testing is “neutralizing” antibody detection, as opposed to “binding” antibody detection described above. This process involves incubating serum with a live version of the virus. The analytes of interest are the antibodies that have the capability to prevent infection by the virus (i.e. neutralization). Identification of these antibodies may contain useful clinical information and are often reported in an aggregate titer, as opposed to specifying each individual antibody (CDC, 2022h; Espejo et al., 2020). Due to use of live virus, these tests may need to be performed in a higher biosafety laboratory, although some techniques using pseudoviruses may be performed in less restrictive laboratories (CDC, 2022i).

Clinical Utility and Validity of Host Antibody Testing
Antibody testing has many potential uses.  Ideally, the use of an accurate, reliable antibody test could possibly show whether someone has previously been exposed to the virus.  This could indicate possible immunity in an individual. Please note that the antibody test is not used as a diagnostic test, meaning it should not be used to diagnose an acute infection.  Within the FDA policy for diagnostic testing for COVID-19, issued on November 15, 2021 they state, “Results from antibody testing should not be used to diagnose or exclude SARS-CoV-2 infection” (FDA, 2022e).  

The FDA published a Frequently Asked Questions (FAQ) update on March 19, 2021. In it, they remarked that antibody tests cannot be used to diagnose COVID-19, and that molecular or antigen tests are available for diagnostic purposes (FDA, 2022a).

Since SARS-CoV-2 is a new, emerging virus, it is not known for certain how long it takes for the seroconversion to occur or when antibodies start to appear in the blood at high enough concentrations for accurate testing results.  A recent study published in Clinical Infectious Diseases reports an average of seroconversion time for IgM and IgG at 12 and 14 days, respectively (Zhao et al., 2020). A small study (n = 34 patients) reports the presence of IgG for at least seven weeks (the duration of the study) (Xiao et al., 2020).  Another study, however, reports that IgM testing has similar, if not better positive detection rate than PCR 5.5 days after initial onset of symptoms; however, the total window of antibody detection for IgM was only 5 days long (Guo et al., 2020) (See Figure 1).  If the patient was not tested during the detection window, then the individual would not necessarily have a “positive” result for IgM. The authors also report the detection of IgA antibodies (median onset at 5 days after initial symptoms [IQR 3 – 6 days]), and 92.7% of total samples report a positive result for IgA. This same study also reports that IgG detection occurs, on average, fourteen days after initial onset of symptoms (Guo et al., 2020). Another study reports that IgA-based ELISA testing has higher sensitivity than IgG-based ELISA testing, but the IgG-based ELISA testing has higher specificity. The authors recommend IgG-based testing over the IgA-based testing in immunosurveillance studies since IgG has a longer biological half-life (Okba et al., 2020). At least one published study to date has reported that as many as 6.9% of individuals who previously had tested positive with RT-PCR results did not show the presence of antibodies for the length of the study (at least 40 days after the initial onset of symptoms) (Zhao et al., 2020).  

Ideally, any rapid diagnostic test for the outpatient setting must be accurate and reliable. Current research indicates that the diagnostic window for IgA and IgM is very limited. Some data indicate that host antibody testing can also yield inaccuracies.  Also, for IgG testing, the significance of positive results is questionable at the current time. A positive result could indicate a previous infection, assuming the test did not cross-react with any other IgG the host produced in response to one of the four coronaviruses known to cause the common cold in humans, for example. It is not currently known, however, if the presence of IgG antibodies indicates immunity (or degree thereof) of the host against SARS-CoV-2. The duration of any conferred immunity, or the level of IgG antibodies required to effectively acquire such immunity, are also unknown. Additional research is needed and encouraged.

Lisboa Bastos et al. (2020) performed a meta-analysis to investigate the diagnostic accuracy of serological testing for COVID-19. The authors aimed to identify studies where serological testing was compared to the “reference standard of viral culture or reverse transcriptase polymerase chain reaction”. The authors identified a total of 40 studies for inclusion in the study. The authors found the pooled sensitivity of enzyme linked immunosorbent assays (ELISAs) measuring IgG or IgM to be 84.3% (with a 95% confidence interval [CI] of 75.6% – 90.9%). For lateral flow immunoassays (LFIAs), the pooled sensitivity was found to be 66% (95% CI: 49.3% – 79.3%), and for chemiluminescent immunoassays (CLIAs), the pooled sensitivity was found to be 97.8% (95% CI: 46.2% – 100%). Pooled specificities ranged from 96.6% – 99.7%. Sensitivity was also found to be higher at least three weeks from symptom onset (69.9% to 98.9%) compared to within the first week (13.4% to 50.3%) Of the samples used to calculate specificity, 83% were “from populations tested before the epidemic or not suspected of having COVID-19." The authors performed 49 bias risk assessments (one for methodology and one for patient selection) and identified 48 with a “high risk of patient selection bias” and 36 with “high or unclear risk of bias from performance or interpretation of the serological test”. The authors also noted that only four of the forty studies including outpatients and only two studies evaluated point-of-care testing. The authors concluded that “currently, available evidence does not support the continued use of existing point-of-care serological tests” but acknowledged that “higher quality clinical studies assessing the diagnostic accuracy of serological tests for covid-19 are urgently needed” (Lisboa Bastos et al., 2020).

Kontou et al. (2020) performed a meta-analysis investigating the use of antibody tests in detecting SARS-CoV-2. The authors focused on IgG and IgM tests based on enzyme-linked immunosorbent assays (ELISA), chemiluminescence enzyme immunoassays (CLIA), fluorescence immunoassays (FIA), and lateral flow immunoassays (LFIA). A total of 38 studies encompassing 7848 individuals (3522 COVID-19 cases, 4326 healthy controls) were included. Of the 38 studies, 21 included data for both COVID-19 cases and controls. Fourteen studies using ELISA were included, and the authors found that IgG and IgM perform “similarly” individually, but in combination, resulted in a sensitivity of 0.935. Thirteen studies using CLIA resulted in an IgG sensitivity of 0.944, an IgM sensitivity of 0.810, and a combined IgG/IgM sensitivity of 0.910. The specificities ranged from 0.954 to 0.984. Thirteen studies used LFIA and found the IgG and IgM sensitivities to range from 0.53 – 0.66. Combining IgG and IgM resulted in sensitivities of 0.78 – 0.83. The authors also attempted to analyze FIA-based studies but were unable to due to the paucity of studies (three identified). The authors concluded that ELISA- and CLIA-based testing performed better sensitivity-wise and that LFIA studies are “more attractive for large seroprevalence studies but show lower sensitivity”.  (Kontou et al., 2020)

Ko et al. (2020) investigated the differences in neutralizing antibody production between asymptomatic and “mild” symptomatic COVID-19 patients, compared to pneumonic COVID-19 patients. A total of 70 patients (15 asymptomatic, 49 mild symptomatic, and 6 pneumonic) were included. A microneutralization assay was performed, along with a FIA and ELISA. Neutralizing antibody production was observed in all the pneumonic patients, 93.9% of the mildly symptomatic patients, and 80% of the asymptomatic patients. Further, the entire pneumonic group showed “high” titer (defined as ≥ 1:80), while 36.7% of the mild group and 20% of the asymptomatic group showed high titer. Both the FIA (for IgG) and ELISA detected anti SARS-CoV-2 at a high sensitivity (98.8% and 97.6% respectively). The authors concluded that “Most asymptomatic and mild COVID-19 patients produced the neutralizing antibody, although the titers were lower than pneumonia patients” (Ko et al., 2020).

Wu et al. (2020) investigated the association between levels of neutralizing antibodies (NAbs) and clinical characteristics in recovered COVID-19 patients. A total of 175 patients with “mild” symptoms of COVID-19 were included. The authors found that NAbs were detected in patients starting in days 4-6 and reached peak levels in days 10 – 15. NAbs were also found not to cross-react with SARS-associated CoV,but correlated with “spike-binding antibodies targeting S1, receptor binding domain, and S2 regions. The authors also noted that NAbs titers were “significantly” higher in 56 “older” patients (1537 [IQR, 877-2427]) and 63 “middle-aged” patients (1291 [IRQ, 504-2126]) compared to 56 “younger patients” (459 [IQR, 225-998]). The authors concluded that “… NAb titers to SARS-CoV-2 appeared to vary substantially. Further research is needed to understand the clinical implications of differing NAb titers for protection against future infection” (Wu et al., 2020).

Kweon et al. (2020) collected 97 samples from patients with COVID-19 to analyze the serologic profiles and time kinetics of IgG and IgM against SARS-CoV-2 using the AFIAS COVID-19 Ab (BodiTechMed, 2020) and the EDI™ Novel Coronavirus COVID-19 ELISA Kit (EpitopeDiagnostics, 2020). The AFIAS assay uses recombinant nucleocapsid protein as an antigen to determine IgG and IgM antibodies against SARS-CoV-2 within 20 minutes from whole blood, serum, or plasma. The EDI™ ELISA Kit uses the microplate-based enzyme immunoassay technique to detect antibodies by measuring the optical densities (ODs) of each well of immunocomplexes. To determine the kinetics of antibodies, studies were performed at different past symptom onset (PSO) periods and to determine diagnostic accuracy of serologic assays, diagnostic sensitivity and specificities were calculated by PSO of ≤ 14 days and > 14 days. Kinetic studies showed that “with both assays, IgM and IgG rapidly increased after 7 days post symptom onset (PSO). IgM antibody levels reached a peak at 15 – 35 d PSO and gradually decreased. IgG levels gradually increased and remained at similar levels after 22 – 35 d” (Kweon et al., 2020). The diagnostic accuracy of both serologic assays also differed based on PSO. “The sensitivity of IgG samples from ≤ 14 d PSO was as low as 35.7% – 57.1%, but it sharply increased for > 14 d PSO to 88.2% – 94.1%. This means that almost all patients with COVID-19 showed seroconversion after 14 d PSO, and IgG seronegative subjects in this period are considered less likely to be infected with SARS-CoV-2. In addition, both assays showed 94.2 – 96.4% of IgG specificities and increased IgG titers in COVID-19 patients were maintained. Thus, IgG serologic assays can be useful for ruling out SARS-CoV-2 infection after 14 d PSO, detecting past infection, and epidemiologic surveys” (Kweon et al., 2020). For IgM, the sensitivities were “as low as 21.4% (same in both assays) in the samples collected ≤ 14 d PSO and 41.2% – 52.9% in samples > 14 d PSO. These findings indicated that in patients infected with SARS-CoV-2, IgM seroconversion may not develop or might not be detected until the middle or late stages of infection. In other words, SARS-CoV-2 infection may be missed based on IgM seropositivity; thus, IgM tests must not be solely used in COVID-19 diagnosis and should be used only as a supportive tool in addition to molecular tests” (Kweon et al., 2020). In addition, IgM titers in COVID-19 patients showed a significant reduction after 35 d PSO; therefore, their utility in detecting past infection is limited. The author concludes that “testing for antibodies against SARS-CoV-2, especially IgG, has the potential for ruling out SARS-CoV-2 infection after 14 d PSO, detecting past infection, and epidemiologic surveys” (Kweon et al., 2020).

Caturegli et al. (2020) performed a case-control study to determine the clinical utility and validity of using SARS-CoV-2 antibodies, which were serum IgG and IgA antibodies formed against the SARS-CoV-2 spike protein detected by enzyme-linked immunosorbent assay (ELISA). When assays were formed 14 days or later after symptom onset, the researchers found that the sensitivity was 0.976 (95% CI, 0.928 to 0.995) and specificity was 0.988 (95% CI, 0.974 to 0.995), but the sensitivity decreased at earlier time points. Antibodies “predicted the odds of developing acute respiratory distress syndrome, which increased by 62% (CI, 48% to 81%; P < 0.001) for every 2-fold increase in IgG.” This demonstrates the linkage of antibodies used to measure clinical severity and for those who tested negative by NAAT but remained potentially COVID-positive. 

In a household cohort study, Churiwal assessed the utility of a rapid point of care test for COVID-19 antibodies by comparing the performance of BioMedomics COVID-19 IgM/IgG Rapid Antibody Test against an ELISA. The test was performed on 303 patients at study enrollment and 4 weeks later. According to the results, sensitivity was lower early in infection and those who never developed symptoms (74% sensitivity). Only two were detected among 499 tests early in infection due to false-positive IgM bands. When measured 4 weeks later after the onset of symptoms, it demonstrated robust sensitivity (90%) and complete specificity (100%). The authors conclude that "When used appropriately, rapid antibody tests offer a convenient way to detect symptomatic infections during convalescence” (Churiwal et al., 2021). 

Antigen Testing
Another possible diagnostic testing methodology is antigen detection testing, which relies upon the direct detection of parts of the virus called “antigens” — in this instance, proteins located on the outside of SARS-CoV-2, such as the spike protein (S) or nucleocapsid protein, that can cause an immune response in an individual.  What makes this method of testing distinct from antibody testing is that antigen testing directly measures the presence of the virus in a person whereas antibody testing is measuring the patient’s response to an infection. These antigen detection tests can be deployed as rapid antigen tests that decrease the turnaround time for results but usually lack specificity (Loeffelholz & Tang, 2020).

On May 8, 2020, the FDA issued the first EUA for antigen testing for COVID-19 to the Quidel Corporation for their Sofia^®2 SARS Antigen FIA lateral flow immunofluorescent sandwich assay for the qualitative detection of the nucleocapsid (N) protein antigen of SARS-CoV-2 for use in individuals suspected of COVID-19 by their healthcare provider (Quidel_Corporation, 2020).  This test has been approved as a point-of-care (POC) test (FDA, 2022c). This test functions by detecting the N protein of either the SARS-CoV or SARS-CoV-2 virus from an upper respiratory sample (either a nasal swab or nasopharyngeal swab). First, the sample is placed in a reagent tube so that any virus, if present, is broken apart to allow for the N proteins to be exposed. The sample then travels from the sample well, down a test strip — where the term “lateral flow” is derived —where the proprietary reagents will recognize any N proteins and trap them in place on the strip. The test requires at least 15 minutes to develop prior to analysis. The strip can then be read by the Sofia®2 system that measures the fluorescent signal from the proprietary reagents.  The Sofia^®2 system allows the user to have two different modes for analysis — “Walk Away” and “Read Now." For the “Walk Away” mode, the user will insert the test cassette strip into the system, and the results will be displayed in 15 minutes because the test will be developed while in the instrument. In “Read Now” mode, the user must have already allowed at least 15 minutes for the test to develop prior to inserting it into the instrument. Then, the Sofia^®2 system will display the result within one minute (Quidel_Corporation, 2020). On August 20, 2020, Quidel reported that the Sofia test’s labeling had been amended to include “either nasal or nasopharyngeal swabs” thereby allowing Quidel a second corresponding kit configuration (BioSpace, 2020).

On July 2, 2020, a second antigen test (BD Veritor System for Rapid Detection of SARS-CoV-2) from Becton, Dickinson, and Company was issued an EUA. This test is described as “a chromatographic digital immunoassay intended for the direct and qualitative detection of SARS-CoV-2 nucleocapsid antigens in nasal swabs from individuals who are suspected of COVID-19 by their healthcare provider within the first five days of the onset of symptoms”. The test is authorized for use in point-of-care (POC) settings. The test’s mechanism of action is as follows: if there are any antigens in the sample (in this case, the nucleocapsid of the virus), they will bind to antibodies conjugated to detector particles in the test strip. The new “conjugates” migrate to the “reaction area” and are captured by another line of antibodies. The test reads positive when the conjugate is found at both “Control” and “Test” positions on the device. BD Veritor reported the following values for the test (in comparison to RT-PCR): 84% positive predictive agreement, 100% negative predictive agreement, 98% overall percent agreement, 100% positive predictive value, and 97.5% negative predictive value. No cross-reactivity was reported (BD_Veritor, 2020).

On August 18, 2020, a third antigen test (LumiraDx SARS-CoV-2 Ag Test from LumiraDx UK Ltd.) was issued an EUA. The test is described as “a single use fluorescence immunoassay device designed to detect the presence of the nucleocapsid protein antigen directly from SARS-CoV-2 in nasal swab specimens, without transport media”. The mechanism of action is as follows: when a droplet of the specimen is added to the “Test Strip”, pre-made reagents on the strip react with any antigen in the specimen. The amount of fluorescence created is proportional to the amount of antigen detected. LumiraDx reported a limit of detection of 32 TCID50/mL [tissue-culture infectious dose], as well as a 97.6% positive percent agreement, 96.6% negative percent agreement, 93.1% positive predictive value, 98.8% negative predictive value, and 96.9% overall percent agreement (based on 257 total samples) (LumiraDx, 2020).

As of April 20, 2022, 50 antigen tests have Emergency Use Authorization (EUA) by the U.S. Food and Drug Administration (FDA) (FDA, 2022d). These testing methods include (among others): Bulk Acoustic Wave (BAW) Biosensors, Chemiluminescence Immunoassays, Chromatographic Digital Immunoassays, Digital Lateral Flow, Magnetic Force-assisted Electrochemical Sandwich Immunoassay (MESIA), Microfluidic Immunofluorescence Assay, and Paramagnetic Microbead-based Immunoassay (FDA, 2022d). 

Clinical Utility and Validity of Antigen Testing
To address the clinical performance, two primary studies were performed. Both studies only used frozen samples. The first study used 143 samples with 80% PPA or Positive Percent Agreement (47/59 of positive samples tested “positive”). They report 100% NPA or Negative Percent Agreement— all 84 negative samples tested “negative”. The second study used a total of 48 samples. Again, 80% of the positive samples tested “positive”; however, only a total of five positive samples were included within this second study. The remaining 43 samples were all negative samples. This study reports a sensitivity of 80.0%, but a 95% confidence interval range of 37.6% – 96.4%. A third supportive study was also performed.  In this study, thirty swabs were taken.  Twenty of these swabs were spiked with one lower concentration of the virus while the remaining ten swabs were spiked with a higher concentration of the virus.  Then, all 30 swabs were tested and compared to 47 control (“unspiked”) samples. In this study, none of the “unspiked” control samples tested “positive” while all 30 of the “spiked” samples, regardless of the concentration, tested positive.  Quidel also tested the limit of detection (LoD) of the Sofia®2 SARS Antigen FIA test. LoD is typically measured by determining the TCID50 (median tissue culture infective dose). The TCID50 is the amount where 50% of the cells within a sample are infected.(Wulff et al., 2012) For the Sofia®2 SARS Antigen FIA test, the LoD for a direct swab sample has a TCID50 of 113 mL whereas it is 850 mL if the initial sample is from a swab sample that has been diluted into 3 mL of reagent.  Finally, Quidel also checked this antigen test for possible cross-reactivity with a number of microorganisms and other viruses.  It shows no cross-reactivity with any of the microorganisms or viruses tests other than SARS-CoV. Of note, it does not cross-react with human coronavirus 229e, OC43, NL63, or MERS-CoV (heat-inactivated); however, they did not check for possible cross-reactivity with the other known human coronavirus (HKU1) due to a lack of availability at this time.  This is noteworthy since this coronavirus is associated with the common cold. Limitations of the Sofia^®2 SARS Antigen FIA test includes the following:

• This test must be performed using the Sofia^®2 system, and the test must be performed accurately following the test procedure.  Failure to do so can adversely affect the performance of the test and may invalidate the results.
• A positive test cannot distinguish between a SARS-CoV or a SARS-CoV-2 infection. SARS-CoV is the virus that caused the SARS outbreak of 2003. It should be noted that there is no current outbreak of SARS. 
• This test also does not distinguish between “live” (viable) virus and non-viable virus. Consequently, the test results do not necessarily correlate with viral culture results performed on the same sample.
• This test is only for the qualitative use on a sample from either a nasal swab or a nasopharyngeal swab.  It has not been approved for use, at this time, on any other sample, such as saliva.
• Negative test results can occur if the viral level is below the lower limit of the test. All negative results “should be treated as presumptive and confirmed with an FDA authorized molecular assay, if necessary, for clinical management, including infection control”(Quidel_Corporation, 2020) 
• Positive test results do not rule out co-infections, and negative results do not “rule in” other non-SARS viral or bacterial infections.
• The clinical performance assays submitted for FDA approval were performed using frozen samples; the test may have a different performance when used with a fresh sample (such as in a point-of-care setting).
• “If the differentiation of specific SARS viruses and strains is needed, additional testing, in consultation with state or local public health departments is required (Quidel_Corporation, 2020).”
•  As previously noted, the company did not check this test (as of publication date) for cross-reactivity with human coronavirus HKU1 due to a lack of availability of that strain.  This is notable since this particular virus is associated with upper respiratory conditions such as the common cold.

One multi-center study, currently a preprint at the time of publication, reports the development of another rapid antigen detection test (RADT) that screens for SARS-CoV-2 by targeting the nucleocapsid protein. This test, when using a nasopharyngeal swab sample, reports a 100% positive agreement with RT-PCR testing.  They also report 73.6% positive agreement when using a urine sample (Diao et al., 2020). This study is yet to be published in a peer-reviewed journal, and the test is not FDA-approved as of May 18, 2020. Another study published recently in ACS Nano reports on the development of a RADT using field-effect transistor (FET)-based biosensing where a graphene sheet for the FET is coated with a specific antibody against the SARS-CoV-2 spike protein. This method can detect the protein in concentrations as low as 1 fg/mL in buffer and has an LOD of 242 copies/mL for a clinical sample (versus 16/mL for a culture medium) (Seo et al., 2020). To date, the WHO states that “Ag-RDTs could play a significant role in guiding patient management, public health decision making and in surveillance of COVID-19. Currently, there is insufficient evidence on performance and operational use to recommend specific commercial products” (WHO, 2021a). 

Scohy et al. (2020) evaluated the Coris COVID-19 Ag [Antigen] Respi-Strip test in comparison to RT-PCR. The authors tested 148 nasopharyngeal swabs, with 106 testing positive by RT-PCR. The rapid antigen test detected 32 of these 106 positive results, for a sensitivity of 30.2%. All samples deemed positive by the antigen test were also deemed positive by RT-PCR. The authors noted that higher viral loads were associated with better detection by antigen tests but concluded that “the overall poor sensitivity of the COVID-19 Ag Respi-Strip does not allow using it alone as the frontline testing for COVID-19 diagnosis” (Scohy et al., 2020).

Mak et al. (2020) evaluated the BIOCREDIT COVID-19 Ag test in comparison to RT-PCR. The BIOCREDIT test’s limit of detection (LOD) was compared to RT-PCR and viral culture, and a total of 368 samples from confirmed COVID-19 cases were included. A sample volume of 100 μL was used. The authors found the LOD of BIOCREDIT to be 1000-fold less sensitive than viral culture (BIOCREDIT LOD: 10-2, viral culture: 10-5). RT-PCR’s LOD was measured to be 10-7. Further, BIOCREDIT detected between 11.1% and 45.7% of RT-PCR positive patients from COVID-19 patients. The authors concluded that “This study demonstrated that the RAD test serves only as adjunct to RT-PCR test because of potential for false-negative results” (Mak et al., 2020).

Lambert-Niclot et al. (2020) analyzed the COVID-19 Ag Respi-Strip test and compared its accuracy to RT-PCR. A total of 138 nasopharyngeal samples were included, with 94 testing positive by RT-PCR. The Respi-Strip test identified 47 of 94 positive specimens for a sensitivity of 50%, although the specificity was 100% for both tests. The authors also noted that the control lines were “barely” visible for 17 tests (9 positive and 8 negative). The authors acknowledged that due to the low prevalence in France (the country in which this study was performed), prospective studies should be undertaken(Lambert-Niclot et al., 2020).

Hirotsu et al. (2020) evaluated a new antigen test (LUMIPULSE) which is based on chemiluminescence enzyme immunoassay. A total of 313 nasopharyngeal swabs were included (82 serial samples from 7 COVID patients, 231 individual samples from 4 COVID patients and 215 healthy controls). These samples were tested by both LUMIPULSE and RT-PCR. Compared to RT-PCR, LUMIPULSE demonstrated a 91.4% overall agreement rate (286/313), with a 55.2% sensitivity and 99.6% specificity. At > 100 viral copies, LUMIPULSE agreed perfectly with RT-PCR, and at 10-100 viral copies, there was an 85% concordance rate (with concordance declining at lower viral loads). The authors concluded that “the LUMIPULSE antigen test can rapidly identify SARS-CoV-2-infected individuals with moderate to high viral loads and may be helpful for monitoring viral clearance in hospitalized patients” (Hirotsu et al., 2020).

Villaverde et al. (2021) conducted a multicenter study to compare the diagnostic accuracy of the Panbio coronavirus disease 2019 Antigen Rapid Test of nasopharyngeal samples in pediatric patients with COVID-19 symptoms ≤ 5 days. They demonstrated “limited accuracy in nasopharyngeal antigen testing: overall sensitivity was 45.4%, and 99.8% of specificity, positive-predictive value was 92.5%,” with moderate concordance between the RT-PCR and antigen test. They noted that a high proportion of false-negative results from the antigen tests (54.5%) may have public health implications in unknown spreading of the virus. But because this test has a good positive likelihood ratio, and is cheap, rapid, and widely distributed, it may be used as a first screening test in a pandemic situation, though its value as a diagnostic tool is questionable due to the low sensitivity and negative likelihood ratio.

Peacock studied the clinical utility of the BinaxNOW antigen test by Abbott Diagnostics, a lateral flow immunochromatographic point-of-care test which provides results in 15 minutes from a nasal swab. BinaxNOW was performed on 735 samples and results were compared to PCR. 623 of 735 (84.8%) had symptoms and 460 of 623 patients (62.6%) had symptoms for less than 7 days. Positive tests occurred in 173 (23.5%) for the PCR and 141 (19.2%) with the BinaxNOW test. Those with symptoms for more than 2 weeks had a positive test rate half of those with earlier onset." In patients with symptoms ≤ 7 days, the sensitivity, specificity, and negative and positive predictive values for the BinaxNOW test were 84.6%, 98.5%, 94.9%, and 95.2%, respectively" (Peacock et al., 2022). The authors conclude that BinaxNOW has good sensitivity and specificity and is recommended for patients with symptoms up to 2 weeks (Peacock et al., 2022).

Panel Testing
Multiple laboratories have developed panels to screen for possible microorganism infections from a single sample. For example, multiplex polymerase chain reaction (PCR) can simultaneously detect multiple pathogens rather than sequentially testing for each individual pathogen.  Such testing can be advantageous when different pathogens may manifest with similar clinical presentation; however, this testing can be costly and can also result in false-negatives if preferential amplification of one target over another occurs (Palavecino, 2015). As of May 4, 2022, the BioFire^® Respiratory Panel 2.1 (RP2.1), the QIAstat-Dx^® Respiratory SARS-CoV-2 Panel, ePlex Respiratory Pathogen Panel 2, cobas SARS-CoV-2 & Influenza A/B, Xpert Xpress SARS-CoV-2/Flu/RSV,  Quest Diagnostics RC COVID-19 +Flu RT-PCR, Sofia 2 Flu + SARS Antigen FIA, and the Influenza SARS-CoV-2 (Flu SC2) Multiplex Assay from the CDC received an EUA from the FDA for testing for COVID-19 (FDA, 2022c). The BioFire^® Respiratory Panel 2.1, the QIAstat-Dx^® Respiratory SARS-CoV-2 Panel, and ePlex Respiratory Pathogen Panel 2 use multiplex nucleic acid testing from a nasopharyngeal swab to detect and differentiate microorganisms listed in Table 1 (BioFire, 2020; GenMark_Diagnostics, 2020; Qiagen_GmbH, 2020), whereas the CDC Multiplex detects and differentiates influenzas A and B from SARS-CoV-2 (CDC, 2020c).
 

Table 1: Respiratory Pathogen Panel Testing Containing SARS-CoV-2
BioFire® Respiratory Panel 2.1	QIAstat-Dx® Respiratory SARS-CoV-2 Panel	ePlex Respiratory Pathogen Panel 2
Adenovirus HCoV 229E HCoV HKU1 HCoVNL63 HCoV OC43 SARS-CoV-2 Human Metapneumovirus Human Rhinovirus/Enterovirus Influenza A Subtype H1 Subtype H3 Subtype H1-2009 Influenza B Parainfluenza Virus 1 Parainfluenza Virus 2 Parainfluenza Virus 3 Parainfluenza Virus 4 Respiratory Syncytial Virus Bordetella parapertussis Bordetella pertussis Chlamydia pneumoniae Mycoplasma pneumoniae	Adenovirus HCoV 229E HCoV HKU1 HCoVNL63 HCoV OC43 SARS-CoV-2 Human Metapneumovirus A+B Influenza A Subtype H1 Subtype H3 Subtype H1N1/pdm09 Influenza B Parainfluenza Virus 1 Parainfluenza Virus 2 Parainfluenza Virus 3 Parainfluenza Virus 4 Rhinovirus/Enterovirus Respiratory Syncytial Virus A+B Bordetella pertussis Chlamydia pneumoniae Mycoplasma pneumoniae	Adenovirus HCoV 229E HCoV HKU1 HCoVNL63 HCoV OC43 SARS-CoV-2 Human Metapneumovirus A+B Influenza A Subtype H1 Subtype H3 Subtype H1-2009 Influenza B Parainfluenza Virus 1 Parainfluenza Virus 2 Parainfluenza Virus 3 Parainfluenza Virus 4 Rhinovirus/Enterovirus Respiratory Syncytial Virus A+B Chlamydia pneumoniae Mycoplasma pneumoniae

Clinical Utility and Validity of Panel Testing
The BioFire RP2.1 panel must be used with either the BioFire FilmArray 2.0 or BioFire FilmArray Torch Systems, and it does not provide a quantitative value for any particular organism within the sample.  This panel “has not been established for specimens collected from individuals without signs or symptoms of respiratory infection (BioFire, 2020).” This panel has not been validated for the monitoring of treatment for any condition.  If a test result shows four or more organisms detected, then the sample should be retested. A negative result does not necessarily exclude an infection. “Negative test results may occur from the presence of sequence variants (or mutation) in the region targeted by the assay, the presence of inhibitors, technical error, sample mix-up, an infection caused by an organism not detected by the panel, or lower respiratory tract infection that is not detected by a nasopharyngeal swab specimen (BioFire, 2020).” 

The BioFire RP2.1 panel cannot necessarily distinguish between existing viral strains and new variants.  One example is the inability to distinguish between Influenza A H3N2v and seasonal Influenza A H3N2. This panel also cannot reliably differentiate between human rhinovirus and enterovirus due to genetic similarity. If detected, the “result should be followed-up using an alternate method (e.g. cell culture or sequence analysis) if differentiation between the viruses is required (BioFire, 2020).” The performance characteristics of several microorganisms detected by this panel, including HCoV 229E, were determined using retrospective clinical specimens due to the small number of positive specimens collected.  The BioFire RP2.1 panel should not be used if B. pertussis is suspected because of its low sensitivity. “[A] B. pertussis molecular test that is FDA-cleared for use on patients suspected of having a respiratory tract infection attributable to B. pertussis only should be used instead (BioFire, 2020).” This is because the RP2.1 panel targets a single-copy promoter target (ptxP) whereas more sensitive tests target the multi-copy IS481 insertion sequence. The BioFire RP2.1 panel also shows cross-reactivity with B. bronchiseptica and B. parapertussis at higher concentrations.

The primers used in the BioFire RP2.1 panel to detect COVID-19 may cross-react with coronaviruses from other species due to high sequence homology. BioFire reports predicted cross-reactivity with up to three bat coronaviruses (accession: MN996532, MG772933, and MG772934) and one pangolin coronavirus (accession: MT084071). However, “[i]t is unlikely that these viruses would be found in a human clinical nasopharyngeal swab; but if present, the cross-reactive product(s) produced by the BioFire RP2.1 will be detected as Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2) (BioFire, 2020).”

The difference between the BioFire RP2 panel and the BioFire RP2.1 panel is the ability to detect SARS-CoV-2. Consequently, within the Instructions for Use (IFU) for the RP2.1 panel, BioFire reports on the data of the RP2 panel.  The clinical performance of the RP2 panel was determined using both fresh and frozen samples. The clinical performance values for the four endemic HCoVs are listed in Table 2 (BioFire, 2020). They note a cross-reactivity between HCoV-OC43 and HCoV-HKU1.

Table 2: Clinical Performance of BioFire RP2/RP2.1 Panel for Endemic HCoVs
Analyte	PPA	PPA 95% CI	NPA	NPA 95%CI
HCoV-229E	11/12 (91.7%)	64.6 – 98.5	1595/1600 (99.7%)	99.3 – 99.9
HCoV-HKU1	43/43 (100%)	91.8 – 100	1557/1569 (99.2%)	98.7 – 99.6
HCoV-NL63	40/40 (100%)	91.2 – 100	1562/1572 (99.4%)	98.8 – 99.7
HCoV-OC43	33/41 (80.5%)	66.0 – 89.8	1566/1571 (99.7%)	99.3 – 99.9
Notes: Abbreviations used—PPA (Positive Percent Agreement); NPA (Negative Percent Agreement).

Concerning the detection of SARS-CoV-2, the BioFire RP2.1 panel reports a limit of detection (LoD), using the USA-WA1/2020 isolate, of 500 copies/mL when using a heat-inactivated virus. They report a 100% detection rate (20/20).  This equates to 6.9 X 10-2 TCID50/mL.   They also tested the LoD using an infectious virus isolate obtained from the World Reference Center for Emerging Viruses and Arboviruses (WRCEVA), contributed by the CDC. With this infectious sample, the LoD was determined to be 160 copies/mL (or 1.1 X 10-2 TCID50/mL). Again, they report a 100% detection rate (20/20) (BioFire, 2020).

Similar to the BioFire panel test, the QIAstat-Dx Respiratory SARS-CoV-2 panel test by Qiagen is for use on a proprietary system, the QIAstat Dx Analyzer System. It is also a qualitative test approved for testing in “patients suspected of COVID-19 by their healthcare provider”. It is also “not intended to be used as the sole basis for diagnosis, treatment, or other patient management decisions” (Qiagen_GmbH, 2020). It is important to note that the test performance in either immunocompromised individuals or asymptomatic individuals has not been established as of publication date. A positive test result cannot rule out a co-infection; an erroneous negative test result can be due to erroneous sample handling as well as variations in the target sequences, organism levels below the limits of detection, and/or use of an interfering reagent (such as certain medications or therapies). Since the QIAstat-Dx test targets the E gene of SARS-CoV-2, which is homologous to sequences in multiple bat SARS viruses, it is possible to cross-react with these bat SARS viruses; however, the likelihood of infection of these viruses in humans is unlikely since none have been reported to date (Qiagen_GmbH, 2020). 

Also, like the BioFire RP2/RP2.1 panel tests, the QIAstat-Dx test may not distinguish between existing viral strains and emerging viral strains, such as influenza A. However, unlike the BioFire RP2/RP2.1 panel tests, the QIAstat-Dx test does detect the IS481 multi-copy insertion sequence present in multiple Bordetella species.  This does increase the sensitivity of the test, but it can increase the possibility of false-positive results if the specimen is contaminated with a non-pertussis Bordetella species (Qiagen_GmbH, 2020).

In addressing the clinical performance of the QIAstat-Dx test for detecting SARS-CoV-2, Qiagen set up two positive trials (one at a higher concentration sample [n = 10] and one at a low positive contrived sample [n = 20), and they report a positive percent agreement (PPA) of 100% (30/30) (95% CI: 85.8 – 100%). Likewise, they did a negative control (n = 30) and report a negative percent agreement (NPA) of 100% (30/30) (95% CI: 85.8 – 100%). In reporting the limit of detection (LoD), they used 20 replicates with a detection rate of at least 95% (or 19/20) to generate a ‘positive’ signal. Using source material obtained from the clinical sample strain of the Hospital of Barcelona (Spain), Qiagen reports an LoD of 500 copies/mL.

The performance of the other targets within the panel were assessed in a multi-center study conducted at six geographically diverse study sites—Copenhagen, Denmark; Minneapolis, MN; Indianapolis, IN; Liverpool, NY; Columbus, OH; and Albuquerque, NM. The performance was determined using both frozen and fresh samples. The clinical performance values for the four endemic HCoVs are listed in Table 3 (Qiagen_GmbH, 2020).

Table 3: Clinical Performance of QIAstat-Dx Panel for Endemic HCoVs
Analyte	PPA	PPA 95% CI	NPA	NPA 95%CI
HCoV-229E	8/9 (88.9%)	56.5 – 98.0	1975/1975 (100%)	99.8 – 100.0
HCoV-HKU1	51/52 (98.1%)	89.9 – 99.7	1925/1932 (99.6%)	99.3 – 99.8
HCoV-NL63	40/47 (85.1%)	72.3 – 92.6	1936/1938 (99.9%)	99.6 – 100.0
HCoV-OC43	26/29 (89.7%)	73.6 – 96.4	1951/1955 (99.8%)	99.5 – 99.9
Notes: Abbreviations used—PPA (Positive Percent Agreement); NPA (Negative Percent Agreement).

As with the other two tests, the ePlex RP2 Panel “should not be used as the sole basis for diagnosis, treatment, or other patient management decisions. Positive results are indicative of active infection with the identified respiratory pathogen but do not rule out infection or co-infection with non-panel organisms. The agent detected by the ePlex RP2 Panel may not be the definite cause of disease. Negative results for SARS-CoV-2 and other organisms on the ePlex RP2 Panel may be due to infection with pathogens that are not detected by this test, or lower respiratory tract infection that may not be detected by a nasopharyngeal swab specimen. Negative results do not preclude infection with SARSCoV-2 or other organisms on the ePlex RP2 Panel and should not be used as the sole basis for patient management decisions. Negative results must be combined with clinical observations, patient history, and epidemiological information (GenMark_Diagnostics, 2020).” A limitation of ePlex RP2 Panel is its unpredictability in differentiating human rhinovirus and enterovirus due to genetic similarity. If differentiation is required, an ePlex RP2 Panel positive human rhinovirus/enterovirus result should be followed-up using an alternative method, such as cell culture or sequence analysis. Cross-reactivity with SARS-CoV-1 is also observed at high titers.

To test the performance characteristics of ePlex RP2 Panel for SARS-CoV-2 detection, 170 nasopharyngeal previously frozen swab samples were collected (59 known SARS-CoV-2 positive and 111 presumed SARSCoV-2 negative samples). “Positive percent agreement (PPA) was calculated by dividing the number of true positive (TP) results by the sum of TP and false negative (FN) results, while negative percent agreement (NPA) was calculated by dividing the number of true negative (TN) results by the sum of TN and false positive (FP) results” (GenMark_Diagnostics, 2020). The ePlex RP2 Panel detected SARS-CoV-2 in 59/59 positive specimens (100% positive percent agreement) and confirmed 111/111 negative specimens (100% negative percent agreement). To determine the limit of detection (LoD), the lowest concentration at which SARS-CoV-2 is detected at least 95% of the time, serial dilutions were prepared in a natural clinical matrix and at least 20 replicates per concentration were tested in the study. “The LoD concentration for detection of SARS-CoV-2 was determined to be 0.01 TCID₅₀/mL, which corresponds to 250 genomic copies per milliliter, as determined by digital droplet PCR (GenMark_Diagnostics, 2020).”

Regarding the “Influenza SARS-CoV-2 (Flu SC2) Multiplex Assay” from the CDC, the FDA reported a limit of detection (LOD) of 1.01 x 10^-2 (at ID­₅₀ [infective dose] / reaction). The panel was evaluated using 104 samples (33 positive for SARS-CoV-2, 30 positives for influenza A, and 30 positive for influenza B, 11 negative samples), and compared to an RT-PCR assay. There was a 100% concordance rate between the two tests. Additionally, cross-reactivity between the three analytes and 35 common respiratory pathogens (16 viruses, 18 bacterial species, 1 yeast) was evaluated, and no cross-reactivity was identified (FDA, 2020b).

The cobas SARS-CoV-2 & Influenza A/B panel is approved for emergency use authorization by the FDA; the panel uses qualitative detection of nucleic acids from SARS-CoV-2 in pooled samples. Six cultured viruses are tested for, two each of influenza A and influenza B strains as well as SARS-CoV-2. In an independent study, Poljak et al. (2020) performed a clinical evaluation of the cobas SARS-Cov-2 test (non-inclusive of influenza A/B panel). The cobas SARS-CoV-2 test was evaluated against an in-house and well-characterized comparator using 217 samples. cobas and the comparator showed overall agreement of 98.1%. Another comparative evaluation on 502 samples showed agreement of 99.6%. The authors concluded that cobas “is a reliable assay for qualitative detection of SARS-CoV-2 in nasopharyngeal swab samples collected in the Universal Transport Medium System (UTM-RT)(Poljak et al., 2020).”

There are other panels that are not yet FDA approved such as the AMPLIQUICK® Respiratory Triplex assay that detects and differentiates between SARS-CoV-2, influenza A/B and respiratory syncytial viruses in respiratory specimens. Results from AMPLIQUICK® were compared to the Allplex™ Respiratory Panel 1 and 2019-nCoV assays. 359 predetermined respiratory samples with diagnosed SARS-CoV-2, influenza A, influenza B and RSV were included in the study. The AMPLIQUICK^® Respiratory Triplex “showed high concordance with the reference assays, with an overall agreement for SARS-CoV-2, influenza A, influenza B, and RSV at 97.6%, 98.8%, 98.3% and 100.0%, respectively.” The authors conclude that the "AMPLIQUICK® Respiratory Triplex is a reliable assay for the qualitative detection and differentiation of SARS-CoV-2, influenza A, influenza B, and RSV in respiratory specimens, which may prove useful for streamlining diagnostics during the winter influenza-seasons" (Mboumba Bouassa et al., 2022).

Miscellaneous Testing
Other methodologies have been proposed to complement or even replace the standard tests described above. For example, a new “RT-LAMP” (reverse transcription loop-mediated isothermal amplification) application has started to see some use for the COVID-19 pandemic. This technique attempts to combine the speed of antigen testing and the accuracy of nucleic acid testing; RT-LAMP includes the traditional reverse transcriptase (RT), as well as a DNA polymerase with “strong strand displacement activity and tolerance for elevated temperatures and up to six DNA oligonucleotides of a certain architecture." These oligonucleotides act as primers for the RT, but additional oligonucleotides for the DNA polymerase are designed so that the DNA products loop back into their ends. This results in “self-priming templates” for the DNA polymerase, which allows the reaction [the nucleic acid amplification] to proceed as normal. Detection of the amplified DNA without specialized instrumentation is the key challenge; some tests use a pH indicator that changes the color of the solution the reaction is run in. Since the reaction does not require the use of a thermal cycler with real-time fluorescence measurement, the results can be delivered in a faster time frame than traditional RT-PCRs (Dao Thi et al., 2020).

Nagura-Ikeda et al. (2020) evaluated the “clinical performance of six molecular diagnostic tests and a rapid antigen test for severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2)." Self-collected saliva was the medium used for analysis. A total of 103 patients with COVID-19 were included (15 asymptomatic, 88 symptomatic). The six molecular diagnostic tests included three RT-PCR tests, an RT-qPCR test, a “cobas SARS-CoV-2 high-throughput system” and an RT-LAMP assay. The molecular diagnostic tests detected viral RNA in 50.5% – 81.6% of specimens and an antigen was detected in 11.7% of the specimens by the rapid antigen test. Viral RNA was also detected at a higher rate (65.6% – 93.4%) in specimens collected within 9 days of symptom onset compared to specimens collected after 10 days (22.2% – 66.7%). Viral RNA was detected in asymptomatic patients at a rate of 40% – 66.7%. The authors concluded “Self-collected saliva is an alternative specimen option for diagnosing COVID-19. LDT RT-qPCR … and RT-LAMP showed sufficient sensitivity in clinical use to be selectively used according to clinical settings and facilities. The rapid antigen test alone is not recommended for initial COVID-19 diagnosis because of its low sensitivity” (Nagura-Ikeda et al., 2020).

Dao Thi et al. (2020) performed a validation of a “two-color RT-LAMP assay protocol for detecting SARS-CoV-2 viral RNA using a primer set specific for the N gene." The authors wrote that a positive sample would be detected by a color change from red to yellow and tested their RT-LAMP assay on “surplus RNA samples isolated from 768 pharyngeal swab specimens collected from individuals being tested for COVID-19”. The results were compared to a traditional RT-qPCR assay. The specificity of the RT-LAMP assay was found to be 99.7%. Further, the RT-qPCR positive samples with a cycle threshold (CT) number of under 30 scored positive (agreeance) in the RT-LAMP assay at a 97.5% agreeance rate. Agreeance rate declined both at the 30-35 threshold and at the 35 – 40 threshold. The authors also developed a “swab-to-RT” LAMP protocol, which was measured at 86% sensitivity (for CT <30) and a 99.5% specificity. The authors concluded that “The RT-LAMP assay and LAMP-sequencing extend the range of available test methods and complement individual tests and pooled tests based on RT-qPCR with a faster, simpler, and potentially more cost-effective test method” (Dao Thi et al., 2020).

R. Wang et al. (2020) demonstrated a one-pot visual SARS-CoV-2 detection system named “opvCRISPR” by integrating reverse transcription loop-mediated isothermal amplification (RT-LAMP) and Cas12a cleavage in a single reaction system, which simplifies operations and avoids contamination. The opvCRISPR enables detection at every single molecular level in forty-five minutes. “The RT-LAMP reagents are incubated at the bottom of the tube, and CRISPR/Cas12a reaction reagents are added on the lid. SARS-CoV-2 RNA templates extracted from the respiratory swab are amplified by RT- LAMP, followed by mixing with the Cas12a reagents for cleavage. Once the Cas12a nuclease is activated by recognizing DNA target, it splits the quenched fluorescent single-stranded DNA (ssDNA) reporter (FAM- TTATT-BHQ1) indiscriminately, generating the fluorescence signal visible to the naked eye under blue light” (R. Wang et al., 2020). To investigate the diagnostic accuracy of opvCRISPR, 26 SARS-CoV-2 RT-PCR positive respiratory swab samples and 24 SARS-CoV-2 RT-PCR negative samples were tested. “All infected samples were determined to be SARS-CoV-2 positive while all uninfected samples tested to be negative by both opvCRISPR and RT- PCR. The opvCRISPR diagnostic results provide 100% agreement with the Centers for Disease Control and Prevention (CDC)-approved quantitative RT-PCR assay” (R. Wang et al., 2020). The author states that “the proposed method only requires minimal equipment, demonstrating great potential in enabling next-generation molecular diagnosis towards point-of- care diagnosis. However, the present method requires additional step to extract RNA. Further efforts need to be made to combine the RNA extraction module with the opvCRISPR to achieve from sampling to result nucleic acid detection” (R. Wang et al., 2020).

Another methodology with potential application for COVID-19 testing is next-generation sequencing (NGS). The NGS procedure typically includes the following steps: first the patient’s DNA is prepared to serve as a template, then DNA fragments are isolated (on solid surfaces such as small beads) where sequence data is generated, then these results are compared against a reference genome. Any DNA sample may be used if the quality and quantity of that sample are sufficient, but the methods of library generation and data analysis often vary from panel to panel. NGS is often used to produce swift and high-volume sequencing (Hulick, 2020). The FDA issued an EUA to Illumina, Inc. for the Illumina COVIDSeq Test on June 10, 2020 but has since updated its indications on October 29, 2020 to be for the “qualitative detection of SARS-CoV-2 RNA from nasopharyngeal (NP) swabs, oropharyngeal (OP) swabs, anterior nasal swabs, mid-turbinate nasal swabs, nasopharyngeal wash/aspirates, nasal aspirates, and bronchoalveolar lavage (BAL) specimens from individuals suspected of COVID-19 by their healthcare provider” (FDA, 2021b). The FDA also issued an EUA to Helix OpCo LLC (dba Helix) for the Helix COVID-19 NGS Test on August 6, 2020. The test detects the gene for the SARS-CoV-2 spike protein, as well as one internal control (the human gene RPP30). The limit of detection was found to be 125 genetic copy equivalents / mL, and both the positive and negative percent agreements were measured to be 100% over 30 samples (Helix, 2020).

Furthermore, whole genome sequencing (WGS) has been demonstrated to have application for COVID-19 testing as well. WGS is conducted through four steps of DNA shearing, by using “molecular scissors” to cut DNA; then DNA bar-coding, for which “scientists add small pieces of DNA tags/bar codes to identify which piece of sheared DNA belongs to which [pathogen];” then the bar-coded DNA is put into the whole genome sequencer that identifies the bases; and finally, the data is analyzed to compare sequences and identify possible differences (CDC, 2016). In several countries, like the Netherlands, China, Vietnam, and the United States, particularly rapid WGS has been beneficial in informing outbreak response, general public health decision making, and infection risk in various facilities (Chau et al., 2021; Oude Munnink et al., 2020; Taylor et al., 2020; F. Wang et al., 2020). In the Netherlands, WGS with the first cases in February 2020 was able to confirm separate introductions of the virus into the country, and attribute increases in case prevalence to co-circulating virus variants following the spring holidays. WGS informed the sequence diversity that existed in Italy, which was where most COVID-positive individuals were returning from. The researchers concluded that “WGS in combination with epidemiological data strengthened the evidence base for public health decision-making in the Netherlands as it enabled a more precise understanding of the transmission patterns in various initial phases of the outbreaks. As such, we were able to understand the genetic diversity of the multiple introduction events in phase 1, the extent of local and regional clusters in phase 2 and the transmission patterns within the HCW [healthcare worker] groups in phase 3 (among which the absence or occurrence of very limited nosocomial transmission)” (Oude Munnink et al., 2020). In Vietnam, a similar application was made regarding a previously known strain responsible for a virus outbreak in the northern region. By whole genome sequencing, researchers were able to identify the first case of the B.1.1.7 variant from locally acquired infection. As the outbreak expanded, whole genome sequencing enabled enhanced surveillance in high risk groups, like those working in airports, who ended up being assigned another variant of A.23.1, as well as contact tracing and testing to detect more cases (Chau et al., 2021). In China, whole genome sequencing in this initial genomic study was able to provide insight towards the genotype-phenotype differences between COVID-19 positive patients. The researchers concluded, “Pedigree analysis suggested a potential monogenic effect of loss of function variants in GOLGA3 and DPP7 for critically ill and asymptomatic disease demonstration. Genome-wide association study suggests the most significant gene locus associated with severity were located in TMEM189–UBE2V1 that involved in the IL-1 signaling pathway…We identified that the HLA-A*11:01, B*51:01, and C*14:02 alleles significantly predispose the worst outcome of the patients” (F. Wang et al., 2020).

In the United States, a Morbidity and Mortality Weekly Report (MMWR) released in September 2020 utilized serial testing and virus whole genome sequencing at two skilled nursing facilities with COVID-19 outbreaks from April to June 2020 in Minnesota. From a total of 25 specimens from residents at the two different facilities, “strains from 17 residents and five HCP [health care personnel] were genetically similar, including one collected from a dietary worker with limited resident contact. Specimens from two HCP and one resident at facility A had distinctly different virus sequences from the first cluster and from each other. At facility B, 75 (66%) resident specimens and five (7%) HCP specimens were sequenced, all of which were genetically similar,” which suggested “intrafacility transmission.” However, the limited participation by HCPs in serial testing could have “have biased identification of infections and limited interpretation of genomic sequencing” and limited “the description of genetic diversity” (Taylor et al., 2020). Generally, whole-genome sequencing still seems to have some limitations, in that “it still presents practical difficulties such as high cost, shortage of available reagents in the global market, need of a specialized laboratorial infrastructure and well-trained staff” resulting in “SARS-CoV-2 surveillance blackouts across several countries” (Bezerra et al., 2021). As of May 4, 2022, there are no FDA approved tests specifically for WGS.

Other types of specimens or media have been proposed as viable for COVID-19 testing, such as saliva. Saliva’s primary advantages include its flexibility, its safety, and overall ease of use in testing. Santosh et al. also noted that To et al. found that saliva has a “high consistency rate of greater than 90% with nasopharyngeal specimens in the detection of respiratory viruses, including coronaviruses” (Sri Santosh et al., 2020; To et al., 2019). On August 15, 2020, the FDA issued an EUA to Yale School of Public Health for “SalivaDirect” which uses saliva samples for COVID-19 testing. Although this test still uses RT-PCR, the test still detects the nucleic acids in saliva, but does not require otherwise specialized or proprietary equipment for extraction of those nucleic acids. In the “Performance Evaluation” section of the official EUA, the FDA noted a positive agreement level between SalivaDirect and the ThermoFisher Scientific TaqPath COVID-19 combo kit to be 94.1% (32/34) and a negative agreement level to be 90.9% (30/33). (FDA, 2020a)

A third innovation in COVID-19 testing was published by the FDA on July 18, 2020. On this date, the FDA stated that they reissued an EUA to Quest Diagnostics to authorize Quest SARS-CoV-2 rRT-PCR test for use with “pooled” samples. This testing practice refers to testing multiple samples simultaneously, thereby allowing more efficient testing. The Quest SARS-CoV-2 rRT-PCR test was authorized to test up to 4 samples at once. The FDA notes that this strategy is most efficient in areas with low prevalence of COVID (i.e., most tests are expected to be negative). In the EUA, the FDA writes that if the “positivity rate” for any given individual to be tested is over 25%, the pooling strategy should not be used due to inefficiency (FDA, 2020c). Yelin et al. found that a single positive sample could be identified in pools of up to 32 samples (with a false negative rate of 10%) and noted that detection of a single positive sample in a pool of 64 samples may be possible with additional amplification cycles. (Yelin et al., 2020). Additional EUAs have been issued specifically for tests using pooled samples, such as the UCSD RC SARS-CoV-2 Assay (University of California San Diego Health, RT-PCR, 5 samples), the Poplar SARS-CoV-2 TMA Pooling assay (Poplar Healthcare, TMA [transcription-mediated amplification], 7 samples), and the “COVID-19 RT-PCR Test” (LabCorp, RT-PCR, 5 samples) (LabCorp, 2020a; Poplar, 2020; UCSD, 2020).

Hogan et al. (2020) performed an analysis of pooled sample analysis in a community setting. The authors analyzed samples in pools of 9 or 10, and the RT-PCR assay targeted the envelope (E) gene. When a positive pool was identified, each sample was tested individually for both the E gene and the RNA-dependent RNA polymerase (RdRp) gene for confirmation. The authors investigated 292 pools encompassing 2740 nasopharyngeal samples and 148 bronchoalveolar lavage samples. Two positive samples were identified (0.07%), which both showed detection of both genes. The authors identified one pool with a “positive E signal” that was not reproducible with testing individual samples of that pool. The authors did acknowledge that this methodology may miss individuals in which a COVID-19 risk has not been identified, but concluded that “strategies such as pooled screening may facilitate detection of early community transmission of SARS-CoV-2 and enable timely implementation of appropriate infection control measures to reduce spread (Hogan et al., 2020).

Another innovative technique in COVID-19 testing was announced on April 14, 2022 in an FDA press announcement. The InspectIR COVID-19 Breathalyzer is the first FDA Emergency Use Authorization-approved diagnostic test to use breath samples; the test detects chemical compounds in breath and provides results in less than three minutes. According to the FDA, a validation study of 2,409 individuals (both symptomatic and asymptomatic) showed 91.2% sensitivity and 99.3% specificity for detecting COVID-19. In addition, the study evidenced a negative predictive value of 99.6% in a population composed of only 4.2% of people who were positive for the virus. A follow-up study with the Omicron variant showed similar sensitivity values (FDA, 2022b).

World Health Organization 
The World Health Organization (WHO) published an interim guideline for the diagnostic testing of “2019 novel coronavirus [termed 2019-nCoV]” on September 11, 2020 (WHO, 2020a). First, they state that routine confirmation of COVID-19 cases is based on nucleic acid testing. Regarding serum testing, they remark that “if negative NAAT results are obtained from a patient in whom SARS-CoV-2 infection is strongly suspected, a paired serum specimen could be collected. One specimen taken in the acute phase and one in the convalescent phase 2 – 4 weeks later can be used to look for seroconversion or a rise in antibody titres.” Finally, they recommend against viral culture or isolation as a routine diagnostic procedure and WHO does not recommend the use of saliva as the sole specimen type for routine clinical diagnostics (WHO, 2020a). 

The WHO released a scientific brief with recommendations for the use of SARS-CoV-2 Ag-RDTs, and updated their interim guidance on October 6, 2021. Within the guidelines, “SARS-CoV-2 Ag-RDTs (antigen detecting rapid diagnostic tests) that meet the minimum performance requirements of ≥ 80% sensitivity and ≥ 97% specificity compared to a NAAT reference assay can be used to diagnose SARS-CoV-2 in suspected COVID-19 cases” (WHO, 2021a). Ag-RDTs should be conducted within 5 – 7 days after the onset of symptoms, as “patients who present more than 5 – 7 days after the onset of symptoms are more likely to have lower viral loads, and the likelihood of false negative results with Ag-RDTs is higher.” WHO recommends that Ag-RDTs be used in settings when they are most reliable — in areas “when there is ongoing community transmission (≥ 5% test positivity rate). When there is no transmission or low transmission, the positive predictive value of Ag-RDTs will be low (many false positives), and in this setting NAAT is preferable as the first-line testing method or for confirmation of positive Ag-RDTs.” 

WHO recommends that the following populations be tested with Ag-RDTS: 

• “Symptomatic individuals (suspected COVID-19 cases) in the first 5 – 7 days since onset of symptoms”
• For asymptomatic individuals, only “limited to contacts of confirmed or probable cases and to at-risk health workers until more evidence is available on the benefits and cost effectiveness of testing low-risk groups with no known exposure to SARS-CoV-2, particularly in settings where testing capacity is limited”
• “Suspected COVID-19 cases in outbreak investigations” 

The WHO also indicates the following as priority uses for the Ag-RDTs: 

• “Community testing of symptomatic individuals meeting the case definition of suspected COVID-19”
• “To detect and respond to suspected outbreaks of COVID-19 including in remote settings, institutions and semi-closed communities (e.g., schools, care-homes, cruise ships, prisons, workplaces and dormitories), especially where NAAT is not immediately available”
• “To screen asymptomatic individuals at high risk of COVID-19, including health workers, contacts of cases and other at-risk individuals” 

Overall, “Ag-RDT testing is recommended in settings likely to have the most impact on early detection of cases for care and contact tracing and where test results are most likely to be correct” (WHO, 2021a).

WHO released a second scientific brief with recommendations concerning immunity passports (WHO, 2020b) on April 24, 2020. Within the guidelines, WHO states that as of the publication date, “no study has evaluated whether the presence of antibodies to SARS-CoV-2 confers immunity to subsequent infection by this virus in humans.” They go on to note, “Laboratory tests that detect antibodies to SARS-CoV-2 in people, including rapid immunodiagnostic tests, need further validation to determine their accuracy and reliability. Inaccurate immunodiagnostic tests may falsely categorize people in two ways. The first is that they may falsely label people who have been infected as negative, and the second is that people who have not been infected are falsely labelled as positive. Both errors have serious consequences and will affect control efforts. These tests also need to accurately distinguish between past infections from SARS-CoV-2 and those caused by the known set of six human coronaviruses. Four of these viruses cause the common cold and circulate widely. The remaining two are the viruses that cause Middle East Respiratory Syndrome and Severe Acute Respiratory Syndrome. People infected by any one of these viruses may produce antibodies that cross-react with antibodies produced in response to infection with SARS-CoV-2 (WHO, 2020b)”.

In 2021, WHO released an update to the scientific brief concerning immunity passports within a document titled ‘COVID-19 natural immunity.’ Within this brief, WHO discusses the various testing methods available. WHO notes that “there are many available serologic assays that measure the antibody response to SARS-CoV-2 infection, but at the present time, the correlates of protection are not well understood”. The most measured immune response is the presence of antibodies in serum. Serologic assays to detect the antibody response are usually based on enzyme immunoassays, which detect the presence of virus-specific antibodies in the blood or by live or pseudo-virus neutralization assays, which detect functional NAb. While serologic testing has limited use in clinical management because it does not capture active infection, it can be very useful in determining the extent of infection or estimating attack rates in given populations.  Interpreting the results of serologic testing, however, is complex: there are several antibody types and subtypes and multiple antigenic determinants/epitopes that can be used to target these antibodies, and the results may differ substantially depending on the combinations chosen. The results will also depend on the manufacturing specifics of the assay used” (WHO, 2021c). Other frequently used assays are enzyme-linked immunosorbent tests, chemiluminescent tests, and lateral flow rapid diagnostic tests. To conclude, “available tests and current knowledge do not tell us about the duration of immunity and protection against reinfection, but recent evidence suggests that natural infection may provide similar protection against symptomatic disease as vaccination, at least for the available follow up period.”

WHO released a scientific brief on May 15, 2020, concerning multisystem inflammatory syndrome in children and adolescents with COVID-19. Within the guidelines, they recommend standardized data describing clinical presentations.  

• WHO gives a preliminary case definition for individuals ages 0 – 19 years with fever three or more days AND at least TWO of the following:
  • “Rash or bilateral non-purulent conjunctivitis or muco-cutaneous inflammation signs (oral, hands or feet)
  • Hypotension or shock
  • Features of myocardial dysfunction, pericarditis, valvulitis, or coronary abnormalities (including ECHO findings or elevated Troponin/NT-proBNP)
  • Evidence of coagulopathy (by PT, PTT, elevated d-Dimers)
  • Acute gastrointestinal problems (diarrhea, vomiting, or abdominal pain).
• AND
  • Elevated markers of inflammation such as ESR, C-reactive protein, or procalcitonin
• AND
  • No other obvious microbial cause of inflammation, including bacterial sepsis, staphylococcal or streptococcal shock syndromes
• AND
  • Evidence of COVID-19 (RT-PCR, antigen test or serology positive), or likely contact with patients with COVID-19 (WHO, 2020c)”

Centers for Disease Control and Prevention 
In the update of the CDC guidelines Overview of Testing for SARS-CoV-2, dated February 11, 2022, the CDC states that “viral tests, including nucleic acid amplification tests (NAATs, such as Reverse Transcription-Polymerase Chain Reaction), antigen tests and other tests (such as breath tests) are used as diagnostic tests to detect current infection with SARS-CoV-2 and to inform an individual’s medical care.” The CDC also stated, “Viral testing is recommended for individuals who are close contacts of persons with COVID-19” (CDC, 2022n). Regarding use of antibody testing, the CDC remarks: “Antibody testing is not currently recommended to assess a person’s protection against infection or severe COVID-19 following COVID-19 vaccination or prior infection, or to assess the need for vaccination in an unvaccinated person. To evaluate for evidence of previous infection in a vaccinated individual, an antibody test specifically evaluating IgM/IgG to the nucleocapsid protein should be used (e.g., for public health surveillance or the diagnosis of Multisystem Inflammatory Syndrome in Children (MIS-C) Multisystem Inflammatory Syndrome in Adults (MIS-A)” (CDC, 2022n). 

The interim guideline for COVID-19 testing in non-healthcare workplaces also describes individuals for which SARS-CoV-2 testing with viral tests (i.e., nucleic acid or antigen tests) is “appropriate”, which include “all unvaccinated close contacts (people who have been within 6 feet for a combined total of 15 minutes or more during a 24-hour period).” For these close contacts, “because of the potential for asymptomatic (not having symptoms) or pre-symptomatic (not yet showing symptoms) transmission of SARS-CoV-2, it is important that unvaccinated individuals exposed to people with known or suspected COVID-19 be quickly identified and quarantined.” Generally, for screening in the workplace, “given the incubation period for COVID-19 (up to 14 days), CDC recommends conducting screening testing of unvaccinated, asymptomatic workers without known or suspected exposures at least weekly.” Diagnostic testing should be done for the following situations and populations (CDC, 2021b):

• “Persons with signs or symptoms consistent with COVID-19”
• “Asymptomatic persons with recent known or suspected exposure to SARS-CoV-2”
• “Determining resolution of infection (e.g., ending isolation)”

Screening testing considerations should also be implemented for “testing asymptomatic persons without known or suspected exposure to SARS-CoV-2 for early identification, isolation, and disease prevention.” This guideline is currently inactive on the CDC website (CDC, 2021b). 

Regarding testing for asymptomatic patients with “known or suspected exposure to SARS-CoV-2”, the CDC recommends testing for all close contacts of persons with SARS-CoV-2 infection. “Viral testing is recommended for individuals who have been exposed persons with COVID-19. People who have had an exposure with someone known or suspected of having COVID-19 should be tested at least 5 days after the exposure. If symptoms develop before 5 days, they should get tested immediately” (CDC, 2022n). The CDC also notes the settings to prioritize for screening testing, including: 

• “High-risk congregate settings, such as assisted living facilities, correctional facilities, and homeless shelters, that have demonstrated high potential for rapid and widespread virus transmission to people at high risk for severe illness.”
• “Settings that involve close quarters and that are isolated from healthcare resources (e.g., fishing vessels, wildland firefighter camps, or offshore oil platforms)” (CDC, 2022n).

The CDC states it is "working with state, local, territorial, academic, and commercial partners” for surveillance testing and COVID-19 research in the US (CDC, 2022n).  In general, the CDC remarks that viral testing is “diagnostic” when conducted among symptomatic patients, asymptomatic patients with known or suspected exposure to SARS-CoV-2, and to determine infection resolution. Viral testing is considered “screening” when performed on asymptomatic patients without known exposure to SARS-CoV-2 (CDC, 2021b). 

Finally, the CDC still maintains, “Consequently, evidence supports a time-based and symptom-based strategy to determine when to discontinue isolation or other precautions rather than a test-based strategy. For persons who are severely immunocompromised, a test-based strategy could be considered in consultation with infectious disease experts. For all others, a test-based strategy is no longer recommended” (CDC, 2021b).  

The CDC also published Interim Guidelines for COVID-19 Antibody Testing in Clinical and Public Health Settings on August 1, 2020, and updated the guidelines on January 24, 2022. The CDC states that “Both SARS-CoV-2 IgM and IgG antibodies may be detected around the same time after infection. However, while IgM is most useful for determining recent infection, it usually becomes undetectable weeks to months following infection; in contrast, IgG may remain detectable for longer periods. IgA is important for mucosal immunity and can be detected in mucous secretions like saliva in addition to blood; although, its significance in this disease is still to be determined.” The CDC also acknowledges the potential application of neutralizing antibody detection (as opposed to binding antibody detection), and remarks that the FDA has now authorized one competitive neutralization test (cVNT), which is a “binding antibody tests designed to qualitatively detect potentially neutralizing antibodies, often those that prevent interaction of RBD with the ACE-2 receptor” (CDC, 2022i). 

Regarding testing for past infections, the CDC recommends that antibody tests should not be used to diagnose a current COVID-19 infection (CDC, 2022a).

Within the CDC’s Interim Guidelines for Collecting and Handling Clinical Specimens for COVID-19 Testing, they recommend collecting and testing upper respiratory samples for initial diagnostic testing for current SARS-CoV-2 infections. Within their recommendation, they list the following (without stating a preference) as acceptable specimens:

• “Nasopharyngeal (NP) specimen collection/oropharyngeal (OP) (throat) specimen collection (performed by a trained healthcare provider, only) 
• Nasal mid-turbinate (MT) swab (performed by a healthcare provider or the patient after reviewing and following collection instructions)
• Anterior nasal specimen (performed by a healthcare provider or the patient after reviewing and following collection instructions)
• Nasopharyngeal wash/aspirate or nasal wash/aspirate (NW) (performed by a trained healthcare provider)
• Saliva (collected by patient with or without supervision)
• Breath (performed by a qualified, trained operator under the supervision of a healthcare provider licensed or authorized by state law to prescribe tests)
• Bronchoalveolar lavage, tracheal aspirate, pleural fluid, lung biopsy (generally performed by a physician in the hospital setting)
• Sputum (collected under the guidance of a trained healthcare professional)” (CDC, 2022g). 

The CDC issued employer-based guidelines titled Guidance for Businesses and Employers Responding to Coronavirus Disease 2019 (COVID-19): Plan, Prepare, and Respond to Coronavirus Disease 2019. (CDC, 2021a). This guideline is now considered archived and made available for historical purposes, and the CDC now refers to the OSHA guideline titled Protecting Workers: Guidance on Mitigating and Preventing the Spread of COVID-19 in the Workplace for the latest information. 

The CDC, in a joint interim set of guidelines with OSHA, issued Meat and Poultry Processing Workers and Employers Interim Guidance from CDC and the Occupational Safety and Health Administration (OSHA) (CDC & OSHA, 2021). This guideline is now considered archived and made available for historical purposes, and the CDC now refers to the OSHA guideline titled Protecting Workers: Guidance on Mitigating and Preventing the Spread of COVID-19 in the Workplace.

In their August 31, 2022 update, the CDC recommends a test-based strategy “may be used to remove a mask sooner.” The CDC recommends a symptom-based strategy primarily for healthcare professionals. The recommendations are listed below:

• “For people who are mildly ill with SARS-COV-2 infection and not moderately or severely immunocompromised,
  • Isolation can be discontinued at least 5 days after symptom onset (day 0 is the day symptoms appeared, and day 1 is the next full day thereafter) if fever has resolved for at least 24 hours (without taking fever-reducing medications) and other symptoms are improving.
  • If symptoms recur or worsen, the isolation period should restart at day 0.
  • People who cannot wear a mask, including children < 2 years of age and people of any age with certain disabilities, should isolate for 10 days.
  • In certain high-risk congregate settings that have high risk of secondary transmission, CDC recommends a 10-day isolation period for residents.” 
• “For people who test positive, are asymptomatic (never develop symptoms) and not moderately or severely immune compromised, 
  • Isolation can be discontinued at least 5 days after the first positive viral test (day 0 is the date the specimen was collected for the positive test, and day 1 is the next full day thereafter).”
• “For people who are moderately ill and not moderately or severely immunocompromised, isolation and precautions can be discontinued 10 days after symptom onset (day 0 is the day symptoms appeared, and day 1 is the next full day thereafter).” 
• “For people who are severely ill and not moderately or severely immunocompromised, isolation and precautions can be discontinued 10 days after symptom onset (day 0 is the day symptoms appeared, and day 1 is the next full day thereafter). 
  • Some people with severe (e.g., requiring hospitalization, intensive care, or ventilation support) may remain infectious beyond 10 days. This may warrant extending the duration of isolation and precautions for up to 20 days after symptom onset (with day 0 being the day symptoms appeared) and after resolution of fever for at least 24 hours (without taking fever-reducing medications) and improvement of other symptoms.
  • Serial testing prior to ending isolation can be considered in consultation with infectious disease experts.”
• “For people who are moderately or severely immunocompromised (regardless of COVID-19 symptoms or severity), … CDC recommends an isolation period of at least 20 days, and ending isolation in conjunction with serial testing and consultation with an infectious disease specialist to determine the appropriate duration of isolation and precautions. 
• The criteria for serial testing to end isolation are:
• Results are negative from at least two consecutive respiratory specimens collected In certain high-risk congregate settings that have high risk of secondary transmission, CDC recommends a 10-day isolation period for residents.
• Also, if a moderately or severely immunocompromised patient with COVID-19 was symptomatic, there should be resolution of fever for at least 24 hours (without the taking fever-reducing medication) and improvement of other symptoms” (CDC, 2022e).

In terms of ending isolation for nonhealthcare personnel, the recommendations are below, utilizing a symptom-based strategy as well: 

• “If you had no symptoms, you may end isolation after day 5.”
• “If you had symptoms and:
  • Your symptoms are improving — you may end isolation after day 5 if you are fever-free for 24 hours (without the use of fever-reducing medication).
  • Your symptoms are not improving — continue to isolate until: you are fever-free for 24 hours (without the use of fever-reducing medication) [and] your symptoms are improving.
• “If you had symptoms and had:
  • Moderate illness (you experienced shortness of breath or had difficulty breathing):
    • You need to isolate through day 10.
  • Severe illness (you were hospitalized) or have a weakened immune system:
    • You need to isolate through day 10.
    • Consult your doctor before ending isolation” (CDC, 2022j).

The CDC also published a case series of “Multisystem Inflammatory Syndrome in Adults Associated with SARS-CoV-2 Infection” [MIS-A]. The CDC observes that a “hyperinflammatory syndrome resembling MIS-C” may also manifest in adult patients and remarks that “Clinicians and health departments should consider MIS-A in adults with signs and symptoms compatible with the current working MIS-A case definition. Antibody testing for SARS-CoV-2 might be needed to confirm previous COVID-19 infection in patients who do not have positive SARS-CoV-2 PCR or antigen test results.” The working case definition of MIS-A was defined by CDC as follows:

• “a severe illness requiring hospitalization in a person aged ≥ 21 years; 
• a positive test result for current or previous SARS-CoV-2 infection (nucleic acid, antigen, or antibody) during admission or in the previous 12 weeks;
• severe dysfunction of one or more extrapulmonary organ systems (e.g., hypotension or shock, cardiac dysfunction, arterial or venous thrombosis or thromboembolism, or acute liver injury); 
• laboratory evidence of severe inflammation (e.g., elevated CRP, ferritin, D-dimer, or interleukin-6); and 
• absence of severe respiratory illness (to exclude patients in which inflammation and organ dysfunction might be attributable simply to tissue hypoxia).”

Patients with mild respiratory symptoms who met these criteria were included. Patients were excluded if alternative diagnoses such as bacterial sepsis were identified.

The CDC does note three limitations of the case series report, which are as follows:

• “First, cases described here were voluntarily reported or published and therefore are not representative of the true clinical spectrum or racial/ethnic distribution of this emerging syndrome. Additional cases might not have been reported or published; others might have remained unrecognized because of absence of COVID-like symptoms, lack of antibody testing, or negative test results. 
• Second, the working case definition excludes patients with severe respiratory dysfunction to distinguish MIS-A from severe COVID-19; however, the two conditions might overlap in some cases. 
• Finally, the working case definition for this syndrome is potentially nonspecific, and some patients with other disease processes might have been misclassified as having MIS-A” (Morris et al., 2020).

On July 9, 2021, the CDC updated their research on “Post-COVID Conditions: Information for Healthcare Providers” (CDC, 2022m). According to the CDC, “Post-COVID conditions” can be referred to as “long COVID, post-acute COVID-19, long-term effects of COVID, post-acute COVID syndrome, chronic COVID, long-haul COVID, late sequelae, and the research term “post-acute sequelae of SARS-COV-2 infection (PASC).” Generally, “it can be considered a lack of return to a usual state of health following acute COVID-19 illness” and “might also include development of new or recurrent symptoms unmasking of a pre-existing condition that occurs after the symptoms of acute illness have resolved.” However, the CDC considers post-COVID conditions “to be present if recovery does not occur after the 4-week acute phase even though many patients continue to recover between 4 and 12 weeks.” The CDC also warns of the possibility that some patients with post-COVID conditions won’t have positive tests for COVID-19 because of “a lack of testing or inaccurate testing during the acute period, or because of waning antibody levels or false-negative antibody testing during follow up.” Alternatively, the term “long COVID” is defined as “symptoms lasting 3 or more months that were not present prior to having COVID-19.” The CDC cites the most common symptoms to include: “Dyspnea or increased respiratory effort, fatigue, post-exertional malaise and/or poor endurance, cognitive impairment or “brain fog”, cough, chest pain, headache, palpitations and tachycardia, arthralgia, myalgia, paresthesia, abdominal pain, diarrhea, insomnia and other sleep difficulties, fever, lightheadedness, impaired daily function and mobility, pain, rash (e.g., urticaria), mood changes, anosmia or dysgeusia, menstrual cycle irregularities, [and] erectile dysfunction.” “Post-exertional malaise (PEM)” is defined as “the worsening of symptoms following even minor physical or mental exertion, with symptoms typically worsening 12 to 48 hours after activity and lasting for days or even weeks” (CDC, 2022m).

National Institutes of Health 
The NIH released COVID-19 treatment guidelines, and within the September 26, 2022 update, they addressed clinical spectrum of SARS-CoV-2 infection, which includes those with asymptomatic or presymptomatic infection, mild illness, moderate illness, severe illness, and critical illness. For asymptomatic and presymptomatic individuals, the NIH states that “It is unclear what percentage of individuals who present with asymptomatic infection progress to clinical disease. Some asymptomatic individuals have been reported to have objective radiographic findings consistent with COVID-19 pneumonia.” Additionally, the guideline discusses infectious complications in patients with COVID-19, which can be categorized as “coinfections at presentation,” such as “concomitant viral infections, including influenza and respiratory viruses” and community-acquired bacterial pneumonia, and “reactivation of latent infections,” such as chronic hepatitis B virus and latent tuberculosis reactivation, “nosocomial infections,” such as hospital-acquired or ventilator-associated pneumonia and Clostridioides difficile-associated diarrhea, and “opportunistic fungal infections,” like aspergillosis and mucormycosis among hospitalized COVID-19 patients  (NIH, 2022a).

The NIH also released COVID-19 testing guidelines. The following recommendations were made from the COVID-19 Treatment Guidelines Panel: 

• “To diagnose acute infection of SARS-CoV-2, the panel recommends “using either a nucleic acid amplification test (NAAT) or an antigen test with a sample collected from the upper respiratory tract (e.g., nasopharyngeal, nasal mid-turbinate, or anterior nasal)  to diagnose acute infection of SARS-CoV-2 (AIII).
• A NAAT should not be repeated in an asymptomatic person (with the exception of health care workers) within 90 days of a previous SARS-CoV-2 infection, even if the person has had a significant exposure to SARS-CoV-2 (AIII).
• SARS-CoV-2 reinfection has been reported in people after an initial diagnosis of the infection; therefore, clinicians should consider using a NAAT for those who have recovered from a previous infection and who present with symptoms that are compatible with SARS-CoV-2 infection if there is no alternative diagnosis (BIII).
• The Panel recommends against diagnosing acute SARS-CoV-2 infection solely on the basis of serologic (i.e., antibody) test results (AIII).
• There is insufficient evidence for the Panel to recommend either for or against the use of SARS-CoV-2 serologic testing to assess for immunity or to guide clinical decisions about using COVID-19 vaccines or anti-SARS-CoV-2 monoclonal antibodies (NIH, 2022b).

American Medical Association 
The AMA released public health guidelines and recommendations concerning serological testing for SARS-CoV-2 antibodies on May 14, 2020.  They list the limitations of antibody testing to include the potential for false-positive results, potential cross-reactivity, and lack of knowledge concerning relationship between antibody testing and immune status. The AMA recommends the following (AMA, 2020):

• “Use of serology tests should currently be limited to population-level seroprevalence study, evaluation of recovered individuals for convalescent plasma donations, and in other situations where they are used as part of a well-defined testing plan and in concert with other clinical information by physicians well-versed in interpretation of serology test results.”
• “Serology tests should not be offered to individuals as a method of determining immune status.”
• “Serology tests should not currently be used as the basis for any “immunity certificates,” to inform decisions to return to work, or to otherwise inform physical distancing decisions. Doing so may put individuals, their household and their community at risk.”
• “Serology tests should not be used as the sole basis of diagnosis of COVID-19 infection.”

“Messaging on serological testing to medically underserved communities should explicitly take into consideration cultural and social features which may bear on their ability to make long-term choices on physical distancing and other COVID-19 precautions (AMA, 2020).”

Infectious Diseases Society of America 
The Infectious Diseases Society of America (IDSA) on May 6, 2020, released their guidelines on the diagnosis of COVID-19. At this time, they focus solely on the use of targeted nucleic acid testing, such as RT-PCR, because “[a]t the time of this review, there was little evidence to inform use of serologic testing” (IDSA, 2020b). The IDSA convened a multidisciplinary panel of experts to review the research and literature on the available diagnostic testing for COVID-19. The panel used the Grading of Recommendations Assessment, Development and Evaluation (GRADE) methodology to assess the evidence of the studies and to make their recommendations. A primary recommendation implies that diagnostic testing and specimen collection devices are available whereas a contingency recommendation is made for situations where testing and/or personal protective equipment (PPE) are limited. 

The panel made 17 recommendations concerning the use of nucleic acid testing as follows (IDSA, 2020b):

• They strongly recommend using a nucleic acid amplification test (NAAT), such as RT-PCR, in symptomatic patients even when clinical suspicion for COVID-19 is low.
• They suggest (conditional recommendation) using a nasopharyngeal, mid-turbinate, anterior nasal swab, saliva, or a combined anterior nasal/oropharyngeal swab rather than oropharyngeal swab or saliva sample for testing in symptomatic individuals suspected of COVID-19.
• They suggest (conditional recommendation) that either a patient or a healthcare provider can collect an anterior nasal or mid-turbinate sample in a symptomatic patient with upper respiratory tract infections or influenza-like illness suspected of COVID-19.
• They suggest (conditional recommendation) “initially obtaining an upper respiratory tract sample (e.g., nasopharyngeal swab) rather than a lower respiratory sample for SARS-CoV-2 RNA testing in hospitalized patients with suspected COVID-19 lower respiratory tract in infection. 
  • If the initial upper respiratory sample result is negative, and the suspicion for disease remains high, the IDSA panel suggests collecting a lower respiratory tract sample (e.g., sputum, bronchoalveolar lavage fluid, tracheal aspirate) rather than collecting another upper respiratory sample.”
• They suggest (conditional recommendation) performing only one test in a symptomatic individual and not repeat testing if low clinical suspicion of COVID-19.
• They suggest (conditional recommendation) repeat testing of an initial negative result in a symptomatic individual be performed only if there is an intermediate or high clinical suspicion of COVID-19.
• They suggest (conditional recommendation) “using either rapid RT-PCR or standard laboratory-based NAATs over rapid isothermal NAAT in symptomatic individuals suspected of having COVID-19).”
• They suggest (conditional recommendation) RNA testing in asymptomatic individuals who are either known or suspected to have been exposed to COVID-19.
• They suggest against (conditional recommendation) RNA testing in asymptomatic individuals with no known contact with COVID-19 who are being hospitalized in areas with low prevalence. They consider a low prevalence rate to be less than 2% of the community.
• They suggest (conditional recommendation) RNA testing in asymptomatic individuals with no known contact with COVID-19 who are being hospitalized in areas with high prevalence of the disease. They consider a high prevalence rate to be 10% or higher. The IDSA does note that if the prevalence rate is between 2% and 9% the decision to test should be dependent on the availability of testing resources.
• They strongly recommend RNA testing in immunocompromised asymptomatic individuals who are being admitted to the hospital regardless of exposure to COVID-19.
• They strongly recommend RNA testing (versus no testing) in asymptomatic individuals before immunosuppressive procedures, such as a hematopoietic stem cell (HSCT) or solid organ (SOT) transplant regardless of a known exposure to COVID-19.
• They make NO recommendations “for or against SARS-CoV-2 RNA testing before initiating immunosuppressive therapy in asymptomatic individuals with cancer,” citing an evidence gap. This recommendation does not apply to candidates or recipients of HSCT or SOT.
• They make NO recommendations “for or against SARS-CoV-2 RNA testing before the initiation of immunosuppressive therapy in asymptomatic individuals with autoimmune disease,” citing an evidence gap. 
• They suggest (conditional recommendation) RNA testing in asymptomatic individuals without known exposure to COVID-19 who are undergoing major time-sensitive surgeries.
• They suggest against (conditional recommendation) RNA testing in asymptomatic individuals without known exposure to COVID-19 who are undergoing time-sensitive aerosol-generating procedures, such as a bronchoscopy, when PPE is available.
• They suggest (conditional recommendation) RNA testing in asymptomatic individuals without known exposure to COVID-19 who are undergoing time-sensitive aerosol-generating procedures when PPE is limited and testing is available.  For this recommendation, the IDSA gives greater detail due to restrictions in availability of PPE. They also note that their recommendation does not address the need for repeat testing if patients require multiple procedures over time.

Besides the 17 recommendations, the IDSA panel also released their algorithm for SARS-CoV-2 Nucleic Acid Testing. This algorithm, as seen in Figure 2, separates individuals into symptomatic and asymptomatic groups.  The IDSA notes that testing should be prioritized for symptomatic patients first.  When resources are sufficient, then testing for selected asymptomatic individuals can be considered. Regardless, the preferred testing methodology is direct SARS-CoV-2 nucleic acid amplification testing, such as RT-PCR. 

Figure 2: IDSA Algorithm for SARS-CoV-2 Nucleic Acid Testing (IDSA, 2020a). The Infectious Diseases Society of America (IDSA) released their algorithm for nucleic acid testing for COVID-19. According to the IDSA guidelines, testing priority should first be given to symptomatic patients; if resources are available, then testing asymptomatic individuals can be considered. Regardless, patients undergoing time-sensitive immunosuppressive procedures should be tested (IDSA, 2020b).  

IDSA also published a guideline regarding serology testing on August 18, 2020. In it, they make the following recommendations:

• “The IDSA panel suggests against using serologic testing to diagnose SARS-CoV-2 infection during the first two weeks (14 days) following symptom onset (conditional recommendation, very low certainty of evidence).”
• “When SARS-CoV-2 infection requires laboratory confirmation for clinical or epidemiological purposes, the IDSA panel suggests testing for SARS-CoV-2 IgG or total antibody three to four weeks after symptom onset to detect evidence of past SARS-CoV-2 infection (conditional recommendation, very low certainty of evidence).”
• The IDSA panel makes no recommendation either for or against using IgM antibodies to detect evidence of past SARS-CoV-2 infection (conditional recommendation, very low certainty of evidence).
• “The IDSA panel suggests against using IgA antibodies to detect evidence of past SARS-CoV-2 infection (conditional recommendation, very low certainty of evidence).”
• “The IDSA panel suggests against using IgM or IgG antibody combination tests to detect evidence of past SARS-CoV-2 infection (conditional recommendation, very low certainty of evidence).”
• “The IDSA panel suggests using IgG antibody to provide evidence of COVID-19 infection in symptomatic patients with a high clinical suspicion and repeatedly negative NAAT testing (weak recommendation, very low certainty of evidence).”
• “In pediatric patients with multisystem inflammatory syndrome, the IDSA panel suggests using both IgG antibody and NAAT to provide evidence of current or past COVID-19 infection (strong recommendation, very low certainty of evidence).”
• “The IDSA panel makes no recommendation for or against using capillary versus venous blood for serologic testing to detect SARS-CoV-2 antibodies (knowledge gap).”

IDSA also includes several comments on testing methodologies that are currently under evaluation. The methodologies named in this section of the guideline are “neutralizing antibody and cellular immune responses”, detection of viral antigens aside from the S or N protein, and multi-test algorithms (IDSA, 2020c).

In the 2018 IDSA guidelines, released before the COVID-19 pandemic, IDSA notes, “Suspected cases of SARS coronavirus and MERS coronavirus require immediate notification to the laboratory. Guidance for testing can be found at [CDC websites for SARS and MERS].” For the four endemic human coronaviruses, they only state that they are associated with common cold “symptoms of rhinorrhea, congestion, sore throat, sneezing, and cough and may present with fever”. They do note that for children with asthma or otitis media, these viruses can cause exacerbation of the conditions.  IDSA notes, “Diagnostic tests include NAATs, which are now common in commercial respiratory panels.” Within their table for the laboratory diagnosis of bronchiolitis, bronchitis, and pertussis, the IDSA lists possible diagnostic procedures for the detection of coronavirus to include NAAT, rapid antigen detection tests, and virus culture; however, they do not list one methodology as a preferred or recommended method over another.  For the antigen testing, they do include a footnote stating, “Rapid antigen tests for respiratory virus detection lack sensitivity and depending upon the product, specificity” (Miller et al., 2018).

Occupational Safety and Health Administration 
In the OSHA guidelines for employers, they classify jobs based on exposure risk to COVID-19 (lower, medium, high, or very high). In general, they recommend following CDC and local health department guidelines. They do state that employers should not require documentation for employees to return to work since healthcare provider offices and medical facilities may be extremely busy during the crisis. For jobs classified at medium exposure risk, administrative controls do include the responsibility of the employer to “communicate the availability of medical screening or other worker health resources (e.g., on-site nurse; telemedicine services).” For jobs classified at high or very high exposure risk, administrative controls include that employers should “consider offering enhanced medical monitoring of workers during COVID-19 outbreaks” (OSHA, 2020). OSHA does not state what the term enhanced medical monitoring entails. This guideline is now considered an OSHA Archive Document, and is presented as “historical content, for research and review purposes only.”  

On June 10, 2021, OSHA updated its guidelines for “Protecting Workers: Guidance on Mitigating and Preventing the Spread of COVID-19 in the Workplace.” The guidance was published to “help employers and workers not covered by the Occupational Safety and Health Administration’s (OSHA’s) COVID-19 Emergency Temporary Standard (ETS)  to identify COVID-19 exposure risks to workers who are unvaccinated or otherwise at-risk even if they are fully vaccinated (e.g., if they are immunocompromised.” (OSHA, 2021). The OSHA guideline has been modified to focus “only on protecting unvaccinated or otherwise at-risk, including if they are immunocompromised, and also [on] implement[ing] new guidance involving workers who are fully vaccinated but located in areas of substantial or high community transmission.” 

For implementing the most effective COVID-19 prevention programs, OSHA mentions employers should have measures “such as telework and flexible schedules, engineering controls (especially ventilation), administrative policies (e.g., vaccination policies), personal protective equipment (PPE), face coverings, physical distancing, and enhanced cleaning programs with a focus on high-touch surfaces.” For protecting “unvaccinated or otherwise at-risk workers and mitigating the spread of COVID-19,” employers should:

1. “Facilitate employees getting vaccinated. Employers should grant paid time off for employees to get vaccinated and recover from any side effects.” 
2. “Instruct any workers who are infected, unvaccinated workers who have had close contact with someone who tested positive for SARS-CoV-2, and all workers with COVID-19 symptoms to stay home from work to prevent or reduce the risk of transmission of the virus that causes COVID-19.”
3. “Implement physical distancing in all communal work areas for unvaccinated and otherwise at-risk workers. 
   1. Employers could also limit the number of unvaccinated or otherwise at-risk workers in one place at any given time, for example by implementing flexible worksites (e.g., telework); implementing flexible work hours (e.g., rotate or stagger shifts to limit the number of such workers in the workplace at the same time); delivering services remotely (e.g., phone, video, or web); or implementing flexible meeting and travel options, all for such workers.
   2. At fixed workstations where unvaccinated or otherwise at-risk workers are not able to remain at least 6 feet away from other people, transparent shields or other solid barriers can separate these workers from other people.”
4. “Provide workers with face coverings or surgical masks, as appropriate, unless their work task requires a respirator or other PPE.
   1. Employers should provide face coverings to unvaccinated and otherwise at-risk workers at no cost (and make replacements available to workers when they request them). 
   2. Unless otherwise provided by federal, state, or local requirements, unvaccinated workers who are outdoors may opt not to wear face coverings unless they are at-risk, for example, if they are immunocompromised.
   3. When an employer determines that PPE is necessary to protect unvaccinated and otherwise at-risk workers from exposure to COVID-19, the employer must provide PPE in accordance with relevant mandatory OSHA standards and should consider providing PPE in accordance with other industry-specific guidance.”
5. “Educate and train workers on your COVID-19 policies and procedures using accessible formats and in languages they understand.
   1. Training should be directed at employees, contractors, and any other individuals on site, as appropriate, and should include:
      1. Basic facts about COVID-19, including how it is spread and the importance of physical distancing (including remote work), ventilation, vaccination, use of face coverings, and hand hygiene.
      2. Workplace policies and procedures implemented to protect workers from COVID-19 hazards.
   2. In addition, ensure that workers understand their rights to a safe and healthful work environment, whom to contact with questions or concerns about workplace safety and health, and their right to raise workplace safety and health concerns free from retaliation.”
6. “Suggest or require that unvaccinated customers, visitors, or guests wear face coverings, especially in public-facing workplaces such as retail establishments, and that all customers, visitors, or guests wear face coverings in public, indoor settings in areas of substantial or high transmission.”
7. “Maintain ventilation systems.”
8. “Perform routine cleaning and disinfection. 
   1. If someone who has been in the facility within 24 hours is suspected of having or confirmed to have COVID-19, follow the CDC cleaning and disinfection recommendations.”
9. “Record and report COVID-19 infections and deaths.
   1. Under mandatory OSHA rules in 29 CFR part 1904, employers are required to record work-related cases of COVID-19 illness on OSHA’s Form 300 logs if the following requirements are met: (1) the case is a confirmed case of COVID-19; (2) the case is work-related (as defined by 29 CFR 1904.5); and (3) the case involves one or more relevant recording criteria (set forth in 29 CFR 1904.7) (e.g., medical treatment, days away from work). Employers must follow the requirements in 29 CFR part 1904 when reporting COVID-19 fatalities and hospitalizations to OSHA. More information is available on OSHA's website. Employers should also report outbreaks to local health departments as required and support their contact tracing efforts.” 
10. “Implement protections from retaliation and setting up an anonymous process for workers to voice concerns about COVID-19-related hazards.”
11. “Follow other applicable mandatory OSHA standards. 
    1. All of OSHA's standards that apply to protecting workers from infection remain in place. These mandatory OSHA standards include: requirements for PPE (29 CFR part 1910, Subpart I (e.g., 1910.132 and 133)), respiratory protection (29 CFR 1910.134), sanitation (29 CFR 1910.141), protection from bloodborne pathogens: (29 CFR 1910.1030), and OSHA's requirements for employee access to medical and exposure records (29 CFR 1910.1020). Many healthcare workplaces will be covered by the mandatory OSHA COVID-19 Emergency Temporary Standard”(OSHA, 2021).

Within the workplace, OSHA recommends “maintaining ventilation systems, implementing physical distancing, and properly using face coverings (or other personal protective equipment (PPE) and respiratory protection such as N95 respirators when appropriate) and proper cleaning” (OSHA, 2021).

American Association for Clinical Chemistry (AACC) 
The AACC released a set of recommendations for “implementing and interpreting SARS-CoV-2 EUA and LDT [laboratory developed test] serologic testing in clinical laboratories.” Serologic testing is currently only used for serum, plasma, and “less frequently, whole-blood or dried blood spots,” but not for other sample types, like saliva and cerebrospinal fluid. Serologic testing is “not recommended as the primary approach for diagnosis of SARS-CoV-2 infection.” For the recommended use of serologic testing, the AACC stated the following: 

• “Serologic testing may be offered as an approach to support diagnosis of COVID -19 illness in symptomatic patients and late phase negative molecular testing or for patients presenting with late complications such as multisystem inflammatory syndrome in children (MIS -C).
• Serologic testing can help identify people who may have been infected with or have recovered from the SARS -CoV -2 infection. 
• Serologic testing can be used to screen potential convalescent plasma donors and in the manufacture of convalescent plasma. 
• Serologic testing can be used for epidemiology and seroprevalence studies. 
• Serologic testing can be used for vaccine response and efficacy studies.”

Regarding serologic testing limitations, the AACC stated the following: 

• “False positive results may occur.
• Negative results do not preclude acute SARS CoV-2 infection or viral shedding.
• Serologic tests may not differentiate between natural infection and vaccine response.
• Serologic results should not be used for 
  • Determining individual protective immunity
  • Return to work decisions
  • Cohorting individuals in congregate settings
  • Assessment of convalescent plasma recipients
  • Use of Personal Protective Equipment
  • Placement of high-risk job functions” (Zhang et al., 2021)

European Centre for Disease Prevention and Control 
The ECDC in their guidance for laboratory support in the EU/EEA recommends using WHO-recommended testing strategies for the diagnosis and confirmation of COVID-19 (ECDC, 2022).

In the ECDC’s guideline titled “COVID-19 testing strategies and objectives”, the ECDC recommends performing laboratory testing in accordance with the WHO case definition. The following populations should be tested (ECDC, 2020):

• “Ideally, all people with COVID-19 symptoms should be tested as soon as possible after symptom onset. This requires easy access to testing for all, including non-residents. Test result turnaround time should be minimized, people testing positive should isolate and timely contact tracing should be carried out, ensuring that all close contacts are tested, irrespective of symptoms.
• All patients with acute respiratory symptoms in hospitals and in other healthcare settings, and all specimens from sentinel primary care surveillance should be tested for both SARS-CoV-2 and influenza during the influenza season to monitor incidence and trends over time. 
• Healthcare and social care settings require intensive testing when there is documented community transmission. Periodic and comprehensive testing of all staff and residents/patients is recommended to prevent nosocomial transmission. Furthermore, all patients/residents should be tested upon or just prior to admission.
• Clusters or outbreaks may occur in certain settings, such as workplaces, educational facilities, prisons, and migrant detention centres. Testing policies and systems should be in place for rapid detection and control to protect the relevant populations in these settings and to protect the community from amplified transmission.
• Countries experiencing high SARS-CoV-2 transmission in a local community should consider testing the whole population of the affected area. This would enable identification of infectious COVID-19 cases and allow for their prompt isolation to interrupt chains of transmission. Depending on the epidemiological situation, size and population density of the affected area, such an approach could be less disruptive for society than having to introduce and ensure compliance with more stringent public health measures.
• To prevent re-introduction, countries or subnational areas that achieved sustained control of the circulation of SARS-CoV-2 should, in addition to quarantine measures, consider targeted testing and follow-up of individuals coming from other areas within the same country, or from other countries that have not yet achieved sustained control of the virus” (ECDC, 2020).

Finally, ECDC notes that “Genomic surveillance of SARS-CoV-2 is essential to detect, monitor and assess virus variants that can result in increased transmissibility, disease severity, or have other adverse effects on public health and social control measures. Obtaining timely and accurate information on the emergence and circulation of variants of concern (VOCs) and variants of interest (VOIs) requires robust surveillance systems, including integrated genome sequencing with a well-defined sampling and sequencing strategy to ensure representativeness and reliability of findings” (ECDC, 2020, 2021).

American Academy of Pediatrics 
The AAP lists the most common scenarios for testing as inclusive of: symptomatic patients, patients who are asymptomatic but had exposure to a person with confirmed or probable COVID-19 infection, and patients who required screening as part of local public health, school, or workplace requirement.  The AAP notes that a person’s vaccination status may be a factor in decision-making concerning the need for screening.

Additionally, the AAP says that for patients who have symptoms, both NAATs (such as PCR testing) and antigen tests can be used. A positive result indicates a SARS-CoV-2 infection on either PCR or antigen diagnostics. That said, for a patient with a negative antigen result, a provider may repeat the antigen test at 48 hours per FDA guidance.

For purposes of testing symptomatic children who have recently had confirmed infections within 3 months, the AAP says providers should consider the possibility of a false-positive result. Especially using PCR tests and other NAAT tests, as these may remain positive from deposited viral genetic material for several months after an active infection. The AAP notes, “In a child with known exposure and compatible symptoms, there may be situations in which it is reasonable to retest within the 90-day window. If testing is performed within that window, antigen testing is generally preferable to NAATs because of the potential for positive NAAT results attributable to prior infection” (AAP, 2022a).

Further, the AAP previously stated in 2020-2021 guidance that antibody (serologic) tests “can provide evidence of previous infection with SARS-CoV-2 but are not useful for the diagnosis of acute infection. A positive antibody test result does not prove that a patient has protection against SARS-CoV-2, although the FDA and vaccine companies use serologic testing as a marker for immunogenicity and protection from SARS-CoV-2 infection. Thus, these tests should not be used to make decisions on grouping people in classrooms or other facilities at this time, and individuals with positive antibody tests should continue to adhere to guidelines about masking, social distancing, and other preventive measures” (AAP, 2022a). 

The AAP has also included some comments and discussion on Multisystem Inflammatory Syndrome in Children (MIS-C). MIS-C has been observed to have some association with COVID-19, and patients with this syndrome have been observed to test positive “far more often” for past SARS-CoV-2 infection (i.e. antibody testing) than acute infection (RT-PCR or antigen test). The CDC defines an MIS-C case by the following criteria:

• “An individual aged < 21 years presenting with:
  • fever ( > 38.°C for ≥ 24 hours, or report of subjective fever lasting ≥ 24 hours),
  • laboratory evidence of inflammation (including, but not limited to, one or more of the following: an elevated C-reactive protein (CRP), erythrocyte sedimentation rate (ESR), fibrinogen, procalcitonin, D-dimer, ferritin, LDH, or IL-6, elevated neutrophils, reduced lymphocytes, low albumin),
  • evidence of clinically severe illness requiring hospitalization,
  • with multisystem (≥ 2) organ involvement (cardiac, renal, respiratory, hematologic, gastrointestinal, dermatologic or neurological); AND
• No alternative plausible diagnoses; AND
• Positive for current or recent SARS-CoV-2 infection by RT-PCR, serology, or antigen test; or COVID-19 exposure within the 4 weeks prior to the onset of symptoms.

The CDC delineates a testing algorithm for MIS-C as follows:

• “Evaluate a child who presents with a persistent fever (≥ 3 days) who is moderately to severely ill with clinical signs of organ dysfunction.”
• “Early consultation and coordination with the nearest pediatric infectious disease and rheumatology specialist and pediatric referral center for optimal testing and management should be considered ...”
• “Laboratory screening for systemic inflammation may be considered and initial lab screenings may include complete blood cell count (CBC) with differential, urine analysis, ESR, and CRP, with the addition of ferritin, LDH, comprehensive metabolic panel, pro-BNP, troponin, and fibrinogen depending on initial clinical suspicion and/or evidence of inflammation on initial lab screening.”
• “Any child sick enough to warrant admission for fever, abdominal pain, diarrhea, and/or organ dysfunction in whom MIS-C is suspected should be cared for in a hospital with tertiary pediatric/cardiac intensive care units.
  • Chest radiograph, EKG, and troponin. If any of these or physical examination is abnormal, then consult with pediatric cardiology and consider additional diagnostic testing for myocardial injury (echocardiogram and/or cardiac MRI).
  • Expanded laboratory tests including pro-BNP, triglycerides, creatine kinase, amylase, blood and urine culture, D-dimer, prothrombin time/partial thromboplastin time (PT/PTT), INR, CRP, ferritin, LDH, comprehensive metabolic panel, and fibrinogen, if not already conducted.
  • In all cases, COVID-19 testing should be performed with RT-PCR assay and serologic testing. Later serology may be needed if all are negative initially. Serologic tests must be sent prior to administration of intravenous immunoglobulin (IVIG).” (AAP, 2022b).

American College of Rheumatology 
The ACR published guidance regarding MIS-C associated with COVID-19. In it, they list SARS-CoV-2 IgG, IgM, and IgA as part of the diagnostic pathway for MIS-C (Henderson, Canna, Friedman, Gorelik, Lapidus, Bassiri, Behrens, Ferris, Kernan, Schulert, Seo, MB, et al., 2020).

In a December 5, 2020 update of the above guidelines, the ACR states that ESR, CRP, and testing for SARS-CoV-2 (by PCR or serology) should be considered a “tier 1” (first-line evaluation) for MIS-C (Henderson, Canna, Friedman, Gorelik, Lapidus, Bassiri, Behrens, Ferris, Kernan, Schulert, Seo, Son, et al., 2020).    

In a February 3, 2022 update of the above guideline, the ACR added new information concerning immunomodulatory treatment in MIS-C, hyperinflammation in COVID-19, as well as statements on thrombotic risk and anticoagulation in MIS-C (Henderson et al., 2022).

Table of Terminology

Term	Definition
2019-nCoV	2019 novel coronavirus
AACC	American Association for Clinical Chemistry
AAP	American Academy of Pediatrics
ACE-2	Angiotensin converting enzyme-2
ACR	American College of Rheumatology
ACS	American Chemical Society
Ag-RDTs	Antigen-detecting rapid diagnostic tests
AMA	American Medical Association
ARDS	Acute respiratory distress syndrome
BAL	Bronchoalveolar lavage
BNP	B-type natriuretic peptide
CARES Act	Coronavirus Aid, Relief, And Economic Security Act
Cas12a	CRISPR associated protein 12a
CBC	Complete blood cell count
CDC	Centers for Disease Control and Prevention
cDNA	Complimentary DNA
CFR	Code of Federal Regulations
CI	Confidence interval
CLIA	Chemiluminescence enzyme immunoassay
CLIA ’88	Clinical Laboratory Improvement Amendments Of 1988
CMS	Centers for Medicare & Medicaid Services
COVID-19	Coronavirus disease 2019
CPK	Creatine phosphokinase
CRP	C-reactive protein
CSSE	Center for Systems Science and Engineering
CT	Computerized tomography
CT	Cycle threshold
cVNT	Competitive neutralization test
DNA	Deoxyribonucleic acid
DPP7	Dipeptidyl peptidase 7
ECDC	European Centre for Disease Prevention and Control
ECHO	Echocardiogram
EQA	External quality assessment
ESR	Erythrocyte sedimentation rate
ETS	Emergency temporary standard
EU/EEA	European Union / European Economic Area
EUA	Emergency use authorization
FAQ	Frequently asked questions
FDA	Food and Drug Administration
FET	Field-effect transistor
FIA	Fluorescence immunoassays
Flu SC2	Influenza SARS-CoV-2 (multiplex assay)
FN	False negative
FP	False positive
GISAID	Global initiative on sharing all influenza data
GOLGA3	Golgi autoantigen, golgin subfamily a, 3
GRADE	Grading Of Recommendations Assessment, Development, and Evaluation
HCoVs	Human coronaviruses
HCP	Health care personnel
HCPCS	Healthcare Common Procedure Coding System
HCW	Healthcare worker
HHS	Health And Human Services
HKU1	Human coronavirus
HLA	Human leukocyte antigen
HSCT	Hematopoietic stem cell transplant
ICMA	Immunochemiluminometric assay
ICR	Investigative criteria for suspected cases of SARS-CoV-2 reinfection
IDSA	Infectious Diseases Society of America
IFU	Instructions for use
IgA	Immunoglobulin A
IgG	Immunoglobulin G
IgM	Immunoglobulin M
IL-1	Interleukin 1
IL-6	Interleukin 6
INR	International normalized ratio
IQR	Interquartile range
IVIG	Intravenous immunoglobulin
JAMA	Journal of the American Medical Association
JHU	Johns Hopkins University
LDH	Lactic acid dehydrogenase
LDTs	Laboratory-developed tests
LFIAs	Lateral flow immunoassays
LoD	Limit of detection
MERS	Middle east respiratory syndrome
MERS-CoV	Middle east respiratory syndrome–related coronavirus
MHRA	Medicines & Healthcare Products Regulatory Agency
MIS	Multisystem inflammatory syndrome
MIS-A	Multisystem inflammatory syndrome in adults
MIS-C	Multisystem inflammatory syndrome in children
MMWR	Morbidity And Mortality Weekly Report
MT	Mid-turbinate
N	Nucleocapsid
NAATs	Nucleic acid amplification tests
NAbs	Neutralizing antibodies
NGS	Next-generation sequencing
NIH	National Institutes of Health
NP	Nasopharyngeal
NPA	Negative percent agreement
NT-proBNP	N-terminal pro hormone BNP
NW	Nasopharyngeal wash/aspirate or nasal wash/aspirate
ODs	Optical densities
OP	Oropharyngeal
opvCRISPR	One-pot visual SARS-CoV-2 detection system
OSHA	Occupational Safety and Health Administration
PASC	Post-Acute Sequelae Of SARS-CoV-2 Infection
PCR	Polymerase chain reaction
PEM	Post-exertional malaise
PHE	Public Health England
PHS Act	Public Health Service Act
POC	Point-of-care
POC/NP	Point of care/near person
PPA	Positive percent agreement
PPE	Personal protective equipment
pro-BNP	Pro hormone BNP
PSO	Past symptom onset
PT	Prothrombin time
PTT	Partial thromboplastin time
ptxP	Single-copy promoter target
RADT	Rapid antigen detection test
RBD	Receptor binding domain
RdRp	Ribonucleic acid-dependent ribonucleic acid polymerase
RNA	Ribonucleic acid
RP	Ribonuclease P gene
RP	Respiratory pathogen
RP2	Respiratory panel 2
RP2.1	Respiratory panel 2.1
RT	Reverse transcriptase
RT-LAMP	Reverse transcription loop-mediated isothermal amplification
RT-PCR	Reverse transcription polymerase chain reaction
SARC	Severe acute respiratory syndrome
SARS-CoV	Severe acute respiratory syndrome- coronavirus
SARS-CoV-2	Severe acute respiratory syndrome coronavirus 2
SF-12	Short form 12 Health Survey
SNPs	Single nucleotide polymorphisms
SOT	Solid organ transplant
ssDNA	Single-stranded deoxyribonucleic acid
sVNT	Surrogate viral neutralization test
TCID50	Median tissue culture infective dose
TMA	Transcription-mediated amplification
TMEM189–UBE2V1	PEDS1-UBE2V1 readthrough
TN	True negative
TP	True positive
UCSD	University of California San Diego
VOC	Variant of concern
VUI	Variant under investigation
WGS	Whole genome sequencing
WHO	World Health Organization
WRCEVA	World Reference Center for Emerging Viruses and Arboviruses

References

1. AAP. (2022a, September 7). COVID-19 Testing Guidance.
2. https://services.aap.org/en/pages/2019-novel-coronavirus-covid-19-infections/clinical-guidance/covid-19-testing-guidance/AAP. (2022b, June 10). Multisystem Inflammatory Syndrome in Children (MIS-C) Interim Guidance https://www.aap.org/en/pages/2019-novel-coronavirus-covid-19-infections/clinical-guidance/multisystem-inflammatory-syndrome-in-children-mis-c-interim-guidance/
3. AMA. (2020, 05/14/2020). Serological testing for SARS-CoV-2 antibodies. American Medical Association. Retrieved 05/19/2020 from https://www.ama-assn.org/delivering-care/public-health/serological-testing-sars-cov-2-antibodies
4. ASM. (2021, July 23). How Dangerous is the Delta Variant (B.1.617.2)? https://asm.org/Articles/2021/July/How-Dangerous-is-the-Delta-Variant-B-1-617-2
5. AstraZeneca. (2021, January 29). COVID-19 Vaccine AstraZeneca recommended for use in the EU. Retrieved January 30 from https://www.astrazeneca.com/media-centre/press-releases/2021/covid-19-vaccine-astrazeneca-recommended-for-use-in-the-eu.html
6. Backer, J. A., Klinkenberg, D., & Wallinga, J. (2020). Incubation period of 2019 novel coronavirus (2019-nCoV) infections among travellers from Wuhan, China, 20-28 January 2020. Euro Surveill, 25(5). https://doi.org/10.2807/1560-7917.Es.2020.25.5.2000062 
7. Baum, S. G. (2020). Adult Multisystem Inflammatory Syndrome Associated with COVID-19. NEJM. https://www.jwatch.org/na52622/2020/10/21/adult-multisystem-inflammatory-syndrome-associated-with 
8. BD_Veritor. (2020). Veritor™ System https://www.fda.gov/media/139755/download
9. Bezerra, M. F., Machado, L. C., De Carvalho, V., Docena, C., Brandão-Filho, S. P., Ayres, C. F. J., Paiva, M. H. S., & Wallau, G. L. (2021). A Sanger-based approach for scaling up screening of SARS-CoV-2 variants of interest and concern. Infect Genet Evol, 92, 104910. https://doi.org/10.1016/j.meegid.2021.104910 
10. BioFire. (2020, 05/2020). BioFire® Respiratory Panel 2.1 (RP2.1). FDA. Retrieved 05/04/2020 from https://www.fda.gov/media/137583/download
11. BioGerm. (2020). 2019-nCoV nucleic acid detection kit. http://www.bio-germ.com/zt.php?class_id=102 
12. BioSpace. (2020, 8/20/20). Quidel to Update Packaging of Point-of-Care Sofia® SARS Antigen Test for COVID-19 to Include Either Nasal or Nasopharyngeal Swabs. 
13. BodiTechMed. (2020). AFIAS COVID-19 Ab. http://www.boditech.co.kr/eng/board/news/board_view.asp?num=30109 
14. Branswell, H. (2020, December 19). A side-by-side comparison of the Pfizer/BioNTech and Moderna vaccines. Retrieved January 30 from https://www.statnews.com/2020/12/19/a-side-by-side-comparison-of-the-pfizer-biontech-and-moderna-vaccines/
15. Caturegli, G., Materi, J., Howard, B. M., & Caturegli, P. (2020). Clinical Validity of Serum Antibodies to SARS-CoV-2 : A Case-Control Study. Ann Intern Med, 173(8), 614-622. https://doi.org/10.7326/m20-2889 
16. CDC. (2016, February 11). What is whole genome sequencing (WGS)? https://www.cdc.gov/pulsenet/pathogens/wgs.html
17. CDC. (2020a, 03/15/2020). CDC 2019-Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel FDA. Retrieved 04/30/2020 from https://www.fda.gov/media/134922/download
18. CDC. (2020b, 02/15/2020). Human Coronavirus Types. CDC. Retrieved 05/15/2020 from https://www.cdc.gov/coronavirus/types.html
19. CDC. (2020c, 7/2/2020). Influenza SARS-CoV-2 (Flu SC2) Multiplex Assay. https://www.fda.gov/media/139744/download
20. CDC. (2020d, 05/14/2020). Multisystem Inflammatory Syndrome in Children (MIS-C) Associated with Coronavirus Disease 2019 (COVID-19). CDC. Retrieved 05/26/2020 from https://emergency.cdc.gov/han/2020/han00432.asp
21. CDC. (2020e, 06/06/2020). Research Use Only 2019-Novel Coronavirus (2019-nCoV) Real-time RT-PCR Primer and Probe Information. Centers for Disease Control and Prevention. Retrieved 08/21/20 from https://www.cdc.gov/coronavirus/2019-ncov/lab/rt-pcr-panel-primer-probes.html
22. CDC. (2020f, 12/22/20). Symptoms of Coronavirus. Retrieved 2/11/21 from https://www.cdc.gov/coronavirus/2019-ncov/symptoms-testing/symptoms.html
23. CDC. (2021a, March 8). Guidance for Businesses and Employers: Plan, Prepare, and Respond to Coronavirus Disease 2019. Centers for Disease Control and Prevention. Retrieved 8/21/20 from https://www.cdc.gov/coronavirus/2019-ncov/community/guidance-business-response.html
24. CDC. (2021b, October 7). Interim Guidance for SARS-CoV-2 Testing in Non-Healthcare Workplaces. https://www.cdc.gov/coronavirus/2019-ncov/community/organizations/testing-non-healthcare-workplaces.html
25. CDC. (2021c, May 11). Johnson & Johnson's Janssen COVID-19 Vaccine. https://www.cdc.gov/coronavirus/2019-ncov/vaccines/different-vaccines/janssen.html
26. CDC. (2021d, 5/20/21). Multisystem Inflammatory Syndrome. CDC. Retrieved 04/19/2022 from https://www.cdc.gov/mis/mis-c/hcp/index.html
27. CDC. (2022a, September 28). COVID-19 Testing: What You Need To Know. https://www.cdc.gov/coronavirus/2019-ncov/testing/serology-overview.html
28. CDC. (2022b). COVID-19 Vaccine Booster Shots. https://www.cdc.gov/coronavirus/2019-ncov/vaccines/booster-shot.html?s_cid=11706:cdc%20covid%20booster:sem.ga:p:RG:GM:gen:PTN:FY22 
29. CDC. (2022c, November 11). COVID Data Tracker Variant Proportions. https://covid.cdc.gov/covid-data-tracker/#variant-proportions
30. CDC. (2022d, March 30, 2022). Different COVID-19 Vaccines. Retrieved April 19 from https://www.cdc.gov/vaccines/covid-19/clinical-considerations/interim-considerations-us.html
31. CDC. (2022e, September 14). Ending Isolation and Precautions for People with COVID-19: Interim Guidance. CDC. Retrieved 08/18/2020 from https://www.cdc.gov/coronavirus/2019-ncov/hcp/disposition-in-home-patients.html
32. CDC. (2022f). Genomic Surveillance for SARS-CoV-2 Variants: Predominance of the Delta (B.1.617.2) and Omicron (B.1.1.529) Variants — United States, June 2021–January 2022. https://www.cdc.gov/mmwr/volumes/71/wr/mm7106a4.htm?s_cid=mm7106a4_w 
33. CDC. (2022g, July 15). Interim Guidelines for Collecting and Handling of Clinical Specimens for COVID-19 Testing. https://www.cdc.gov/coronavirus/2019-ncov/lab/guidelines-clinical-specimens.html
34. CDC. (2022h, Jan. 24, 2022). Interim Guidelines for COVID-19 Antibody Testing. Retrieved 04/20/2022 from https://www.cdc.gov/coronavirus/2019-ncov/lab/resources/antibody-tests-guidelines.html
35. CDC. (2022i, January 24). Interim Guidelines for COVID-19 Antibody Testing. https://www.cdc.gov/coronavirus/2019-ncov/lab/resources/antibody-tests-guidelines.html
36. CDC. (2022j, August 11). Isolations and Precautions for People with COVID-19. https://www.cdc.gov/coronavirus/2019-ncov/your-health/isolation.html
37. CDC. (2022k). Omicron Variant: What you need to know. CDC. Retrieved April 19, 2022 from https://www.cdc.gov/coronavirus/2019-ncov/variants/omicron-variant.html
38. CDC. (2022l). Omicron Variant: What You Need to Know. https://www.cdc.gov/coronavirus/2019-ncov/variants/omicron-variant.html 
39. CDC. (2022m, September 22). Post-COVID Conditions: Information for Healthcare Providers. https://www.cdc.gov/coronavirus/2019-ncov/hcp/clinical-care/post-covid-conditions.html
40. CDC. (2022n, September 28). Testing for SARS-CoV-2 (COVID-19) Overview. https://www.cdc.gov/coronavirus/2019-ncov/hcp/testing-overview.html
41. CDC. (2022o). Understanding How COVID-19 Vaccines Work. Retrieved April 19, 2022 from https://www.cdc.gov/coronavirus/2019-ncov/vaccines/different-vaccines/how-they-work.html
42. CDC. (2022p). Variant Proportions. https://covid.cdc.gov/covid-data-tracker/#variant-proportions 
43. CDC. (2022q, March 2022). Viral Vector Vaccines. https://www.cdc.gov/coronavirus/2019-ncov/vaccines/different-vaccines/viralvector.html
44. CDC. (2022r, February 5). What You Need to Know about Variants. Retrieved April 19, 2022 from https://www.cdc.gov/coronavirus/2019-ncov/variants/variant.html
45. CDC, & OSHA. (2021, June 11). Meat and Poultry Processing Workers and Employers: Interim Guidance from CDC and the Occupational Safety and Health Administration (OSHA). Retrieved 08/18/2020 from https://www.cdc.gov/coronavirus/2019-ncov/community/organizations/meat-poultry-processing-workers-employers.html
46. Cevik, M., Tate, M., Lloyd, O., Maraolo, A. E., Schafers, J., & Ho, A. (2021). SARS-CoV-2, SARS-CoV, and MERS-CoV viral load dynamics, duration of viral shedding, and infectiousness: a systematic review and meta-analysis. The Lancet Microbe, 2(1), E13-E22. https://doi.org/10.1016/S2666-5247(20)30172-5 
47. Chan, J. F., Yip, C. C., To, K. K., Tang, T. H., Wong, S. C., Leung, K. H., Fung, A. Y., Ng, A. C., Zou, Z., Tsoi, H. W., Choi, G. K., Tam, A. R., Cheng, V. C., Chan, K. H., Tsang, O. T., & Yuen, K. Y. (2020). Improved Molecular Diagnosis of COVID-19 by the Novel, Highly Sensitive and Specific COVID-19-RdRp/Hel Real-Time Reverse Transcription-PCR Assay Validated In Vitro and with Clinical Specimens. J Clin Microbiol, 58(5). https://doi.org/10.1128/jcm.00310-20 
48. Chau, N. V. V., Hong, N. T. T., Ngoc, N. M., Anh, N. T., Trieu, H. T., Nhu, L. N. T., Yen, L. M., Minh, N. N. Q., Phong, N. T., Truong, N. T., Huong, L. T. T., Tu, T. N. H., Hung, L. M., Thanh, T. T., Dung, N. T., Dung, N. T., Thwaites, G., Van Tan, L., & for, O. C.-r. g. (2021). Rapid whole-genome sequencing to inform COVID-19 outbreak response in Vietnam. The Journal of infection, 82(6), 276-316. https://doi.org/10.1016/j.jinf.2021.03.017 
49. Churiwal, M., Lin, K. D., Khan, S., Chhetri, S., Muller, M. S., Tompkins, K., Smith, J., Litel, C., Whittelsey, M., Basham, C., Rapp, T., Cerami, C., Premkumar, L., & Lin, J. T. (2021). Assessment of the Field Utility of a Rapid Point-of-Care Test for SARS-CoV-2 Antibodies in a Household Cohort. Am J Trop Med Hyg, 106(1), 156-159. https://doi.org/10.4269/ajtmh.21-0592 
50. CMS. (2021, October 21). Frequently Asked Questions: SARS-CoV-2 Surveillance Testing. Retrieved 8/19/20 from https://www.cms.gov/files/document/06-19-2020-frequently-asked-questions-covid-surveillance-testing.pdf
51. Corman, V. M., Lienau, J., & Witzenrath, M. (2019). [Coronaviruses as the cause of respiratory infections]. Internist (Berl), 60(11), 1136-1145. https://doi.org/10.1007/s00108-019-00671-5 (Coronaviren als Ursache respiratorischer Infektionen.) 
52. Cucinotta, D., & Vanelli, M. (2020). WHO Declares COVID-19 a Pandemic. Acta Biomed, 91(1), 157-160. https://doi.org/10.23750/abm.v91i1.9397 
53. Dao Thi, V. L., Herbst, K., Boerner, K., Meurer, M., Kremer, L. P. M., Kirrmaier, D., Freistaedter, A., Papagiannidis, D., Galmozzi, C., Stanifer, M. L., Boulant, S., Klein, S., Chlanda, P., Khalid, D., Barreto Miranda, I., Schnitzler, P., Kräusslich, H.-G., Knop, M., & Anders, S. (2020). A colorimetric RT-LAMP assay and LAMP-sequencing for detecting SARS-CoV-2 RNA in clinical samples. Science Translational Medicine, 12(556), eabc7075. https://doi.org/10.1126/scitranslmed.abc7075 
54. DeBiasi, R. L., Song, X., Delaney, M., Bell, M., Smith, K., Pershad, J., Ansusinha, E., Hahn, A., Hamdy, R., Harik, N., Hanisch, B., Jantausch, B., Koay, A., Steinhorn, R., Newman, K., & Wessel, D. (2020). Severe COVID-19 in Children and Young Adults in the Washington, DC Metropolitan Region. J Pediatr. https://doi.org/10.1016/j.jpeds.2020.05.007 
55. Diao, B., Wen, K., Chen, J., Liu, Y., Yuan, Z., Han, C., Chen, J., Pan, Y., Chen, L., Dan, Y., Wang, J., Chen, Y., Deng, G., Zhou, H., & Wu, Y. (2020). Diagnosis of Acute Respiratory Syndrome Coronavirus 2 Infection by Detection of Nucleocapsid Protein. medRxiv, 2020.2003.2007.20032524. https://doi.org/10.1101/2020.03.07.20032524 
56. Dighe, K., Moitra, P., Alafeef, M., Gunaseelan, N., & Pan, D. (2022). A rapid RNA extraction-free lateral flow assay for molecular point-of-care detection of SARS-CoV-2 augmented by chemical probes. Biosensors and Bioelectronics, 200, 113900. https://doi.org/https://doi.org/10.1016/j.bios.2021.113900 
57. ECDC. (2020, September 18). Testing strategies for SARS-CoV-2. https://www.ecdc.europa.eu/en/covid-19/surveillance/testing-strategies
58. ECDC. (2021, May 3). Guidance for representative and targeted genomic SARS-CoV-2 monitoring. https://www.ecdc.europa.eu/en/publications-data/guidance-representative-and-targeted-genomic-sars-cov-2-monitoring
59. ECDC. (2022, August 15). Diagnostic testing and screening for SARS-CoV-2. European Centre for Disease Prevention and Control. Retrieved 04/18/2022 from https://www.ecdc.europa.eu/en/covid-19/latest-evidence/diagnostic-testing
60. EpitopeDiagnostics. (2020). EDI™ Novel Coronavirus COVID-19 ELISA Kits. http://www.epitopediagnostics.com/covid-19-elisa 
61. Espejo, A. P., Akgun, Y., Al Mana, A. F., Tjendra, Y., Millan, N. C., Gomez-Fernandez, C., & Cray, C. (2020). Review of Current Advances in Serologic Testing for COVID-19. Am J Clin Pathol, 154(3), 293-304. https://doi.org/10.1093/ajcp/aqaa112 
62. FDA. (2020a). ACCELERATED EMERGENCY USE AUTHORIZATION (EUA) SUMMARY SARS-CoV-2 RT-PCR Assay. https://www.fda.gov/media/141192/download 
63. FDA. (2020b, 7/9/2020). CDC Influenza SARS-CoV-2 (Flu SC2) Multiplex Assay. https://www.fda.gov/media/139743/download
64. FDA. (2020c). Coronavirus (COVID-19) Update: FDA Issues First Emergency Authorization for Sample Pooling in Diagnostic Testing. https://www.fda.gov/news-events/press-announcements/coronavirus-covid-19-update-fda-issues-first-emergency-authorization-sample-pooling-diagnostic
65. FDA. (2021a, May 11). Coronavirus (COVID-19) Update: 10/15/21. https://www.fda.gov/news-events/press-announcements/coronavirus-covid-19-update-101521
66. FDA. (2021b, April 22). Illumina COVIDSeq Test. https://www.fda.gov/media/138778/download
67. FDA. (2022a, 02/24/2022). Antibody (Serology) Testing for COVID-19: Information for Patients and Consumers. https://www.fda.gov/medical-devices/coronavirus-covid-19-and-medical-devices/antibody-serology-testing-covid-19-information-patients-and-consumers
68. FDA. (2022b). Coronavirus (COVID-19) Update: FDA Authorizes First COVID-19 Diagnostic Test Using Breath Samples. https://www.fda.gov/news-events/press-announcements/coronavirus-covid-19-update-fda-authorizes-first-covid-19-diagnostic-test-using-breath-samples
69. FDA. (2022c, 04/14/2022). Emergency Use Authorization. Retrieved 04/20/2022 from https://www.fda.gov/emergency-preparedness-and-response/mcm-legal-regulatory-and-policy-framework/emergency-use-authorization
70. FDA. (2022d, January 29). In Vitro Diagnostics EUAs. https://www.fda.gov/medical-devices/coronavirus-disease-2019-covid-19-emergency-use-authorizations-medical-devices/vitro-diagnostics-euas
71. FDA. (2022e). Policy for Coronavirus Disease-2019 Tests During the Public Health Emergency (Revised). FDA. Retrieved 04/20/2022 from https://www.fda.gov/regulatory-information/search-fda-guidance-documents/policy-coronavirus-disease-2019-tests-during-public-health-emergency-revised
72. GenMark_Diagnostics. (2020). ePlex Respiratory Pathogen Panel 2. https://www.fda.gov/media/142902/download 
73. Griffin, D. (2020, December 31). Viral Load as a Predictor of COVID-19 Patient Outcomes. https://www.cuimc.columbia.edu/news/viral-load-predictor-covid-19-patient-outcomes
74. Guo, L., Ren, L., Yang, S., Xiao, M., Chang, D., Yang, F., Dela Cruz, C. S., Wang, Y., Wu, C., Xiao, Y., Zhang, L., Han, L., Dang, S., Xu, Y., Yang, Q.-W., Xu, S.-Y., Zhu, H.-D., Xu, Y.-C., Jin, Q., . . . Wang, J. (2020). Profiling Early Humoral Response to Diagnose Novel Coronavirus Disease (COVID-19). Clinical Infectious Diseases. https://doi.org/10.1093/cid/ciaa310 
75. GVN. (2021, https://gvn.org/covid-19/delta-b-1-617-2/). Delta Variant (B.1.617.2). 
76. Helix. (2020, 8/6/20). Helix COVID-19 NGS Test. Retrieved 8/20/20 from https://www.fda.gov/media/140917/download
77. Henderson, L. A., Canna, S. W., Friedman, K. G., Gorelik, M., Lapidus, S. K., Bassiri, H., Behrens, E. M., Ferris, A., Kernan, K. F., Schulert, G. S., Seo, P., MB, F. S., Tremoulet, A. H., Yeung, R. S. M., Mudano, A. S., Turner, A. S., Karp, D. R., & Mehta, J. J. (2020). American College of Rheumatology Clinical Guidance for Multisystem Inflammatory Syndrome in Children Associated With SARS-CoV-2 and Hyperinflammation in Pediatric COVID-19: Version 1. Arthritis Rheumatol. https://doi.org/10.1002/art.41454 
78. Henderson, L. A., Canna, S. W., Friedman, K. G., Gorelik, M., Lapidus, S. K., Bassiri, H., Behrens, E. M., Ferris, A., Kernan, K. F., Schulert, G. S., Seo, P., Son, M. B. F., Tremoulet, A. H., Yeung, R. S. M., Mudano, A. S., Turner, A. S., Karp, D. R., & Mehta, J. J. (2020). American College of Rheumatology Clinical Guidance for Pediatric Patients with Multisystem Inflammatory Syndrome in Children (MIS-C) Associated with SARS-CoV-2 and Hyperinflammation in COVID-19. Version 2. Arthritis Rheumatol. https://doi.org/10.1002/art.41616 
79. Henderson, L. A., Canna, S. W., Friedman, K. G., Gorelik, M., Lapidus, S. K., Bassiri, H., Behrens, E. M., Kernan, K. F., Schulert, G. S., Seo, P., Son, M. B. F., Tremoulet, A. H., VanderPluym, C., Yeung, R. S. M., Mudano, A. S., Turner, A. S., Karp, D. R., & Mehta, J. J. (2022). American College of Rheumatology Clinical Guidance for Multisystem Inflammatory Syndrome in Children Associated With SARS–CoV-2 and Hyperinflammation in Pediatric COVID-19: Version 3. Arthritis & Rheumatology, 74(4), e1-e20. https://doi.org/https://doi.org/10.1002/art.42062 
80. Hirotsu, Y., Maejima, M., Shibusawa, M., Nagakubo, Y., Hosaka, K., Amemiya, K., Sueki, H., Hayakawa, M., Mochizuki, H., Tsutsui, T., Kakizaki, Y., Miyashita, Y., Yagi, S., Kojima, S., & Omata, M. (2020). Comparison of Automated SARS-CoV-2 Antigen Test for COVID-19 Infection with Quantitative RT-PCR using 313 Nasopharyngeal Swabs Including from 7 Serially Followed Patients. Int J Infect Dis. https://doi.org/10.1016/j.ijid.2020.08.029 
81. Hogan, C. A., Sahoo, M. K., & Pinsky, B. A. (2020). Sample Pooling as a Strategy to Detect Community Transmission of SARS-CoV-2. Jama, 323(19), 1967-1969. https://doi.org/10.1001/jama.2020.5445 
82. Hulick, P. (2020, August 21). Next-generation DNA sequencing (NGS): Principles and clinical applications. https://www.uptodate.com/contents/next-generation-dna-sequencing-ngs-principles-and-clinical-applications
83. IDSA. (2020a, December 23). IDSA Algorithm for SARS-CoV-2 Nucleic Acid Testing. IDSA. Retrieved 05/13/2020 from https://www.idsociety.org/globalassets/idsa/practice-guidelines/covid-19/diagnostics/figure-01.png
84. IDSA. (2020b, December 23). Infectious Diseases Society of America Guidelines on the Diagnosis of COVID-19: Molecular Diagnostic Testing. IDSA. Retrieved 05/13/2020 from https://www.idsociety.org/practice-guideline/covid-19-guideline-diagnostics/
85. IDSA. (2020c, August 18). Infectious Diseases Society of America Guidelines on the Diagnosis of COVID-19: Serologic Testing. https://www.idsociety.org/practice-guideline/covid-19-guideline-serology/
86. JHU. (2022, November 11). COVID-19 Dashboard by the Center for Systems Science and Engineering (CSSE) at Johns Hopkins University (JHU). Johns Hopkins University. https://coronavirus.jhu.edu/map.html
87. Jones, V. G., Mills, M., Suarez, D., Hogan, C. A., Yeh, D., Bradley Segal, J., Nguyen, E. L., Barsh, G. R., Maskatia, S., & Mathew, R. (2020). COVID-19 and Kawasaki Disease: Novel Virus and Novel Case. Hosp Pediatr. https://doi.org/10.1542/hpeds.2020-0123 
88. Kawasuji, H., Takegoshi, Y., Kaneda, M., Ueno, A., Miyajima, Y., Kawago, K., Fukui, Y., Yoshida, Y., Kimura, M., Yamada, H., Sakamaki, I., Tani, H., Morinaga, Y., & Yamamoto, Y. (2020). Transmissibility of COVID-19 depends on the viral load around onset in adult and symptomatic patients. PLOS ONE, 15(12), e0243597. https://doi.org/10.1371/journal.pone.0243597 
89. Ko, J. H., Joo, E. J., Park, S. J., Baek, J. Y., Kim, W. D., Jee, J., Kim, C. J., Jeong, C., Kim, Y. J., Shon, H. J., Kang, E. S., Choi, Y. K., & Peck, K. R. (2020). Neutralizing Antibody Production in Asymptomatic and Mild COVID-19 Patients, in Comparison with Pneumonic COVID-19 Patients. J Clin Med, 9(7). https://doi.org/10.3390/jcm9072268 
90. Kontou, P. I., Braliou, G. G., Dimou, N. L., Nikolopoulos, G., & Bagos, P. G. (2020). Antibody Tests in Detecting SARS-CoV-2 Infection: A Meta-Analysis. Diagnostics (Basel), 10(5). https://doi.org/10.3390/diagnostics10050319 
91. Kweon, O. J., Lim, Y. K., Kim, H. R., Kim, M. C., Choi, S. H., Chung, J. W., & Lee, M. K. (2020). Antibody kinetics and serologic profiles of SARS-CoV-2 infection using two serologic assays. PLOS ONE, 15(10), e0240395. https://doi.org/10.1371/journal.pone.0240395 
92. LabCorp. (2020a, 3/16/20). ACCELERATED EMERGENCY USE AUTHORIZATION (EUA) SUMMARY. Retrieved 8/21/20 from https://www.fda.gov/media/136151/download
93. LabCorp. (2020b, 04/20/2020). ACCELERATED EMERGENCY USE AUTHORIZATION (EUA) SUMMARY COVID-19 RT-PCR TEST (LABORATORY CORPORATION OF AMERICA). FDA. Retrieved 04/26/2020 from https://www.fda.gov/media/136151/download
94. Lambert-Niclot, S., Cuffel, A., Le Pape, S., Vauloup-Fellous, C., Morand-Joubert, L., Roque-Afonso, A. M., Le Goff, J., & Delaugerre, C. (2020). Evaluation of a Rapid Diagnostic Assay for Detection of SARS-CoV-2 Antigen in Nasopharyngeal Swabs. J Clin Microbiol, 58(8). https://doi.org/10.1128/jcm.00977-20 
95. Li, M., Wei, R., Yang, Y., He, T., Shen, Y., Qi, T., Han, T., Song, Z., Zhu, Z., Ma, X., Lin, Y., Yuan, Y., Zhao, K., Lu, H., & Zhou, X. (2021). Comparing SARS-CoV-2 Testing in Anterior Nasal Vestibular Swabs vs. Oropharyngeal Swabs. Front Cell Infect Microbiol, 11, 653794. https://doi.org/10.3389/fcimb.2021.653794 
96. Li, Y., Yao, L., Li, J., Chen, L., Song, Y., Cai, Z., & Yang, C. (2020). Stability issues of RT-PCR testing of SARS-CoV-2 for hospitalized patients clinically diagnosed with COVID-19. Journal of medical virology, 92(7), 903-908. https://doi.org/10.1002/jmv.25786 
97. Lippi, G., Simundic, A. M., & Plebani, M. (2020). Potential preanalytical and analytical vulnerabilities in the laboratory diagnosis of coronavirus disease 2019 (COVID-19). Clin Chem Lab Med. https://doi.org/10.1515/cclm-2020-0285 
98. Lisboa Bastos, M., Tavaziva, G., Abidi, S. K., Campbell, J. R., Haraoui, L. P., Johnston, J. C., Lan, Z., Law, S., MacLean, E., Trajman, A., Menzies, D., Benedetti, A., & Ahmad Khan, F. (2020). Diagnostic accuracy of serological tests for covid-19: systematic review and meta-analysis. Bmj, 370, m2516. https://doi.org/10.1136/bmj.m2516 
99. Loeffelholz, M. J., & Tang, Y.-W. (2020). Laboratory diagnosis of emerging human coronavirus infections – the state of the art. Emerging Microbes & Infections, 9(1), 747-756. https://doi.org/10.1080/22221751.2020.1745095 
100. Lu, Y., Li, L., Ren, S., Liu, X., Zhang, L., Li, W., & Yu, H. (2020). Comparison of the diagnostic efficacy between two PCR test kits for SARS-CoV-2 nucleic acid detection. Journal of Clinical Laboratory Analysis, 34(10), e23554. https://doi.org/10.1002/jcla.23554 
101. Ludwig, S., & Zarbock, A. (2020). Coronaviruses and SARS-CoV-2: A Brief Overview. Anesth Analg. https://doi.org/10.1213/ane.0000000000004845 
102. LumiraDx. (2020). SARS-CoV-2 Ag Test. https://www.fda.gov/media/141304/download
103. Mak, G. C., Cheng, P. K., Lau, S. S., Wong, K. K., Lau, C. S., Lam, E. T., Chan, R. C., & Tsang, D. N. (2020). Evaluation of rapid antigen test for detection of SARS-CoV-2 virus. J Clin Virol, 129, 104500. https://doi.org/10.1016/j.jcv.2020.104500 
104. Mandavilli, A. (2021, July 30). C.D.C. Internal Report Calls Delta Variant as Contagious as Chickenpox. https://www.nytimes.com/2021/07/30/health/covid-cdc-delta-masks.html
105. Mathieu, E., Ritchie, H., Rodés-Guirao, L., Appel, C., Giattino, C., Hasell, J., Macdonald, B., Dattani, S., Beltekian, D., Ortiz-Ospina, E., & Roser, M. (2022, November 11). Coronavirus (COVID-19) Vaccinations. https://ourworldindata.org/covid-vaccinations?country=USA
106. Mawhorter, M. E., Nguyen, P., Goldsmith, M., Owens, R. G., Baer, B., & Raman, J. D. (2022). Diagnostic yield and costs associated with a routine pre-operative COVID-19 testing algorithm for asymptomatic patients prior to elective surgery. Am J Clin Exp Urol, 10(5), 341-344. 
107. Mboumba Bouassa, R.-S., Tonen-Wolyec, S., Veyer, D., Péré, H., & Bélec, L. (2022). Analytical performances of the AMPLIQUICK® Respiratory Triplex assay for simultaneous detection and differentiation of SARS-CoV-2, influenza A/B and respiratory syncytial viruses in respiratory specimens. PLOS ONE, 17(1), e0262258. Retrieved 2022, from 
108. McIntosh, K. (2021, April 2). COVID-19: Clinical features. Wolter Kluwer. Retrieved 08/19/2020 from https://www.uptodate.com/contents/covid-19-clinical-features
109. Miller, J. M., Binnicker, M. J., Campbell, S., Carroll, K. C., Chapin, K. C., Gilligan, P. H., Gonzalez, M. D., Jerris, R. C., Kehl, S. C., Patel, R., Pritt, B. S., Richter, S. S., Robinson-Dunn, B., Schwartzman, J. D., Snyder, J. W., Telford, S., 3rd, Theel, E. S., Thomson, R. B., Jr., Weinstein, M. P., & Yao, J. D. (2018). A Guide to Utilization of the Microbiology Laboratory for Diagnosis of Infectious Diseases: 2018 Update by the Infectious Diseases Society of America and the American Society for Microbiology. Clin Infect Dis, 67(6), e1-e94. https://doi.org/10.1093/cid/ciy381 
110. Morell, A., Skvaril, F., Noseda, G., & Barandun, S. (1973). Metabolic properties of human IgA subclasses. Clinical and experimental immunology, 13(4), 521-528. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1553728/ 
111. Morris, S. B., Schwartz, N. G., Patel, P., Abbo, L., Beauchamps, L., Balan, S., Lee, E. H., Paneth-Pollak, R., Geevarughese, A., Lash, M. K., Dorsinville, M. S., Ballen, V., Eiras, D. P., Newton-Cheh, C., Smith, E., Robinson, S., Stogsdill, P., Lim, S., Fox, S. E., . . . Godfred-Cato, S. (2020). Case Series of Multisystem Inflammatory Syndrome in Adults Associated with SARS-CoV-2 Infection - United Kingdom and United States, March-August 2020. MMWR Morb Mortal Wkly Rep, 69(40), 1450-1456. https://doi.org/10.15585/mmwr.mm6940e1 
112. Nackerdien, Z. (2020, December 6). Viral Load Peaks in First Week of COVID-19 Symptom Onset. Retrieved January 31 from https://www.medpagetoday.org/infectiousdisease/covid19/90035
113. Nagura-Ikeda, M., Imai, K., Tabata, S., Miyoshi, K., Murahara, N., Mizuno, T., Horiuchi, M., Kato, K., Imoto, Y., Iwata, M., Mimura, S., Ito, T., Tamura, K., & Kato, Y. (2020). Clinical evaluation of self-collected saliva by RT-qPCR, direct RT-qPCR, RT-LAMP, and a rapid antigen test to diagnose COVID-19. J Clin Microbiol. https://doi.org/10.1128/jcm.01438-20 
114. NIH. (2020, December 28). Phase 3 trial of Novavax investigational COVID-19 vaccine opens. Retrieved January 30 from https://www.nih.gov/news-events/news-releases/phase-3-trial-novavax-investigational-covid-19-vaccine-opens
115. NIH. (2022a, September 26). Clinical Spectrum of SARS-CoV-2 Infection. National Institutes of Health. https://www.covid19treatmentguidelines.nih.gov/overview/clinical-spectrum/
116. NIH. (2022b, August 8). Testing for SARS-CoV-2 Infection. National Institutes of Health. https://www.covid19treatmentguidelines.nih.gov/overview/sars-cov-2-testing/
117. Okba, N. M. A., Müller, M. A., Li, W., Wang, C., GeurtsvanKessel, C. H., Corman, V. M., Lamers, M. M., Sikkema, R. S., de Bruin, E., Chandler, F. D., Yazdanpanah, Y., Le Hingrat, Q., Descamps, D., Houhou-Fidouh, N., Reusken, C., Bosch, B. J., Drosten, C., Koopmans, M. P. G., & Haagmans, B. L. (2020). Severe Acute Respiratory Syndrome Coronavirus 2-Specific Antibody Responses in Coronavirus Disease 2019 Patients. Emerg Infect Dis, 26(7). https://doi.org/10.3201/eid2607.200841 
118. OSHA. (2020). Guidance on Preparing Workplaces for COVID-19. Occupational Safety and Health Administration. Retrieved 04/29/2020 from https://www.osha.gov/Publications/OSHA3990.pdf
119. OSHA. (2021, June 10). Protecting Workers: Guidance on Mitigating and Preventing the Spread of COVID-19 in the Workplace. https://www.osha.gov/coronavirus/safework#workers-who-have-had-covid-19
120. Oude Munnink, B. B., Nieuwenhuijse, D. F., Stein, M., O’Toole, Á., Haverkate, M., Mollers, M., Kamga, S. K., Schapendonk, C., Pronk, M., Lexmond, P., van der Linden, A., Bestebroer, T., Chestakova, I., Overmars, R. J., van Nieuwkoop, S., Molenkamp, R., van der Eijk, A. A., GeurtsvanKessel, C., Vennema, H., . . . The Dutch-Covid-19 response, t. (2020). Rapid SARS-CoV-2 whole-genome sequencing and analysis for informed public health decision-making in the Netherlands. Nature Medicine, 26(9), 1405-1410. https://doi.org/10.1038/s41591-020-0997-y 
121. Padoan, A., Cosma, C., Sciacovelli, L., Faggian, D., & Plebani, M. (2020). Analytical performances of a chemiluminescence immunoassay for SARS-CoV-2 IgM/IgG and antibody kinetics. Clin Chem Lab Med. https://doi.org/10.1515/cclm-2020-0443 
122. Palavecino, E. (2015). One Sample, Multiple Results The Use of Multiplex PCR for Diagnosis of Infectious Syndromes. Retrieved 11/1 from https://www.aacc.org/publications/cln/articles/2015/april/one-sample-multiple-results
123. Patel, P., DeCuir, J., Abrams, J., Campbell, A. P., Godfred-Cato, S., & Belay, E. D. (2021). Clinical Characteristics of Multisystem Inflammatory Syndrome in Adults: A Systematic Review. JAMA Network Open, 4(9), e2126456-e2126456. https://doi.org/10.1001/jamanetworkopen.2021.26456 
124. Peacock, W. F., Soto-Ruiz, K. M., House, S. L., Cannon, C. M., Headden, G., Tiffany, B., Motov, S., Merchant-Borna, K., Chang, A. M., Pearson, C., Patterson, B. W., Jones, A. E., Miller, J., Varon, J., Bastani, A., Clark, C., Rafique, Z., Kea, B., Eppensteiner, J., . . . Young, S. (2022). Utility of COVID-19 antigen testing in the emergency department. Journal of the American College of Emergency Physicians Open, 3(1), e12605. https://doi.org/https://doi.org/10.1002/emp2.12605 
125. Pfefferle, S., Reucher, S., Nörz, D., & Lütgehetmann, M. (2020). Evaluation of a quantitative RT-PCR assay for the detection of the emerging coronavirus SARS-CoV-2 using a high throughput system. Euro Surveill, 25(9). https://doi.org/10.2807/1560-7917.Es.2020.25.9.2000152 
126. Poljak, M., Korva, M., Gašper, N. K., Komloš, K. F., Sagadin, M., Uršič, T., Županc, T. A., Petrovec, M., & McAdam, A. J. (2020). Clinical Evaluation of the cobas SARS-CoV-2 Test and a Diagnostic Platform Switch during 48 Hours in the Midst of the COVID-19 Pandemic. Journal of Clinical Microbiology, 58(6), e00599-00520. https://doi.org/doi:10.1128/JCM.00599-20 
127. Poplar. (2020). EMERGENCY USE AUTHORIZATION (EUA) SUMMARY OF THE POPLAR SARS-COV-2 TMA POOLING ASSAY. https://www.fda.gov/media/140792/download
128. Qiagen_GmbH. (2020, 03/2020). QIAstat-Dx® Respiratory SARS-CoV2 Panel Instructions for Use (Handbook). FDA. Retrieved 04/27/2020 from https://www.fda.gov/media/136571/download
129. Quidel_Corporation. (2020, 05/2020). Sofia 2 SARS Antigen FIA. FDA. Retrieved 05/12/2020 from https://www.fda.gov/media/137885/download
130. Roberts, M. (2021, January 29). Covid-19: Novavax shows 89% efficacy in UK trials. Retrieved January 30 from https://www.bbc.com/news/uk-55850352
131. Ryding, S. (2020, June 24). What is Viral Load? Retrieved January 31 from https://www.news-medical.net/health/What-is-Viral-Load.aspx
132. SansureBiotech. (2020, 05-04-2020 ). Novel Coronavirus (2019-nCoV) Nucleic Acid Diagnostic Kit (PCR-Fluorescence Probing). https://www.fda.gov/media/137651/download
133. Scohy, A., Anantharajah, A., Bodéus, M., Kabamba-Mukadi, B., Verroken, A., & Rodriguez-Villalobos, H. (2020). Low performance of rapid antigen detection test as frontline testing for COVID-19 diagnosis. J Clin Virol, 129, 104455. https://doi.org/10.1016/j.jcv.2020.104455 
134. Seo, G., Lee, G., Kim, M. J., Baek, S. H., Choi, M., Ku, K. B., Lee, C. S., Jun, S., Park, D., Kim, H. G., Kim, S. J., Lee, J. O., Kim, B. T., Park, E. C., & Kim, S. I. (2020). Rapid Detection of COVID-19 Causative Virus (SARS-CoV-2) in Human Nasopharyngeal Swab Specimens Using Field-Effect Transistor-Based Biosensor. ACS Nano, 14(4), 5135-5142. https://doi.org/10.1021/acsnano.0c02823 
135. Sri Santosh, T., Parmar, R., Anand, H., Srikanth, K., & Saritha, M. (2020). A Review of Salivary Diagnostics and Its Potential Implication in Detection of Covid-19. Cureus, 12(4), e7708. https://doi.org/10.7759/cureus.7708 
136. Taylor, J., Carter, R. J., Lehnertz, N., Kazazian, L., Sullivan, M., Wang, X., Garfin, J., Diekman, S., Plumb, M., Bennet, M. E., Hale, T., Vallabhaneni, S., Namugenyi, S., Carpenter, D., Turner-Harper, D., Booth, M., Coursey, E. J., Martin, K., McMahon, M., . . . Lynfield, R. (2020). Serial Testing for SARS-CoV-2 and Virus Whole Genome Sequencing Inform Infection Risk at Two Skilled Nursing Facilities with COVID-19 Outbreaks - Minnesota, April-June 2020. MMWR Morb Mortal Wkly Rep, 69(37), 1288-1295. https://doi.org/10.15585/mmwr.mm6937a3 
137. The_Native_Antigen_Company. (2020, 03/24/2020). Why We Need Antigen and Antibody Tests for COVID-19. The Native Antigen Company. Retrieved 04/21/2020 from https://thenativeantigencompany.com/why-we-need-antigen-and-antibody-tests-for-covid-19/
138. To, K. K. W., Yip, C. C. Y., Lai, C. Y. W., Wong, C. K. H., Ho, D. T. Y., Pang, P. K. P., Ng, A. C. K., Leung, K. H., Poon, R. W. S., Chan, K. H., Cheng, V. C. C., Hung, I. F. N., & Yuen, K. Y. (2019). Saliva as a diagnostic specimen for testing respiratory virus by a point-of-care molecular assay: a diagnostic validity study. Clin Microbiol Infect, 25(3), 372-378. https://doi.org/10.1016/j.cmi.2018.06.009 
139. Tregoning, J. S., Brown, E. S., Cheeseman, H. M., Flight, K. E., Higham, S. L., Lemm, N. M., Pierce, B. F., Stirling, D. C., Wang, Z., & Pollock, K. M. (2020). Vaccines for COVID-19. Clinical and experimental immunology, 202(2), 162-192. https://doi.org/10.1111/cei.13517 
140. UCSD. (2020). UCSD RC SARS-CoV-2 Assay https://www.fda.gov/media/140712/download
141. US. (2020, 03/27/2020). H.R.748 - CARES Act. Retrieved 05/19/2020 from https://www.congress.gov/116/bills/hr748/BILLS-116hr748enr.pdf
142. Van Beusekom, M. (2020, December 8). Phase 3 trials show AstraZeneca COVID vaccine has up to 90% efficacy. University of Minnesota. Retrieved January 30 from https://www.cidrap.umn.edu/news-perspective/2020/12/phase-3-trials-show-astrazeneca-covid-vaccine-has-90-efficacy
143. Verdoni, L., Mazza, A., Gervasoni, A., Martelli, L., Ruggeri, M., Ciuffreda, M., Bonanomi, E., & D'Antiga, L. (2020). An outbreak of severe Kawasaki-like disease at the Italian epicentre of the SARS-CoV-2 epidemic: an observational cohort study. Lancet. https://doi.org/10.1016/s0140-6736(20)31103-x 
144. Villaverde, S., Domínguez-Rodríguez, S., Sabrido, G., Pérez-Jorge, C., Plata, M., Romero, M. P., Grasa, C. D., Jiménez, A. B., Heras, E., Broncano, A., Núñez, M. D. M., Illán, M., Merino, P., Soto, B., Molina-Arana, D., Bermejo, A., Mendoza, P., Gijón, M., Pérez-Moneo, B., . . . Epidemiological Study of, C.-i. C. o. t. S. S. o. P. W. G. (2021). Diagnostic Accuracy of the Panbio Severe Acute Respiratory Syndrome Coronavirus 2 Antigen Rapid Test Compared with Reverse-Transcriptase Polymerase Chain Reaction Testing of Nasopharyngeal Samples in the Pediatric Population. The Journal of pediatrics, 232, 287-289.e284. https://doi.org/10.1016/j.jpeds.2021.01.027 
145. Wang, F., Huang, S., Gao, R., Zhou, Y., Lai, C., Li, Z., Xian, W., Qian, X., Li, Z., Huang, Y., Tang, Q., Liu, P., Chen, R., Liu, R., Li, X., Tong, X., Zhou, X., Bai, Y., Duan, G., . . . Liu, L. (2020). Initial whole-genome sequencing and analysis of the host genetic contribution to COVID-19 severity and susceptibility. Cell Discovery, 6(1), 83. https://doi.org/10.1038/s41421-020-00231-4 
146. Wang, R., Qian, C., Pang, Y., Li, M., Yang, Y., Ma, H., Zhao, M., Qian, F., Yu, H., Liu, Z., Ni, T., Zheng, Y., & Wang, Y. (2020). opvCRISPR: One-pot visual RT-LAMP-CRISPR platform for SARS-cov-2 detection. Biosensors and Bioelectronics, 172, 112766. https://doi.org/https://doi.org/10.1016/j.bios.2020.112766 
147. WHO. (2020a, 09/11/20). Diagnostic testing for SARS-CoV-2. Retrieved 11/08/20 from https://www.who.int/publications/i/item/diagnostic-testing-for-sars-cov-2
148. WHO. (2020b, 04/24/2020). "Immunity passports" in the context of COVID-19. World Health Organization. Retrieved 04/25/2020 from https://www.who.int/news-room/commentaries/detail/immunity-passports-in-the-context-of-covid-19
149. WHO. (2020c, 05/15/2020). Multisystem inflammatory syndrome in children and adolescents with COVID-19. World Health Organization. Retrieved 05/18/2020 from https://www.who.int/publications-detail/multisystem-inflammatory-syndrome-in-children-and-adolescents-with-covid-19
150. WHO. (2020d, 04/17/2020). Q&A on coronaviruses (COVID-19): What are the symptoms of COVID-19? Retrieved 05/20/2020 from https://www.who.int/emergencies/diseases/novel-coronavirus-2019/question-and-answers-hub/q-a-detail/q-a-coronaviruses
151. WHO. (2021a, October 6). Antigen-detection in the diagnosis of SARS-CoV-2 infection. World Health Organization. Retrieved 11/08/2020 from https://www.who.int/publications/i/item/antigen-detection-in-the-diagnosis-of-sars-cov-2infection-using-rapid-immunoassays
152. WHO. (2021b, November 2021). COVID-19 Clinical management: living guidance. World Health Organization. Retrieved April 19 from https://apps.who.int/iris/bitstream/handle/10665/349321/WHO-2019-nCoV-clinical-2021.2-eng.pdf
153. WHO. (2021c). COVID-19 natural immunity. file:///C:/Users/AHCS8330/Downloads/WHO-2019-nCoV-Sci-Brief-Natural-immunity-2021.1-eng.pdf 
154. WHO. (2022a, November 11). Coronavirus disease (COVID-19) Pandemic. World Health Organization. https://www.who.int/emergencies/diseases/novel-coronavirus-2019
155. WHO. (2022b, 2022). Middle East respiratory syndrome coronavirus (MERS-CoV). World Health Organization. Retrieved 08/19/2020 from https://www.who.int/emergencies/mers-cov/en/
156. WHO. (2022c). SARS (Severe Acute Respiratory Syndrome). World Health Organization. Retrieved 04/19/2022 from https://www.who.int/ith/diseases/sars/en/
157. Woof, J. M., & Kerr, M. A. (2006). The function of immunoglobulin A in immunity. The Journal of Pathology, 208(2), 270-282. https://doi.org/10.1002/path.1877 
158. Wright, D., & CMS. (2021, March 16). Nursing Home Reopening Recommendations for State and Local Officials. CMS. Retrieved 05/20/2020 from https://www.cms.gov/files/document/nursing-home-reopening-recommendations-state-and-local-officials.pdf
159. Wu, F., Liu, M., Wang, A., Lu, L., Wang, Q., Gu, C., Chen, J., Wu, Y., Xia, S., Ling, Y., Zhang, Y., Xun, J., Zhang, R., Xie, Y., Jiang, S., Zhu, T., Lu, H., Wen, Y., & Huang, J. (2020). Evaluating the Association of Clinical Characteristics With Neutralizing Antibody Levels in Patients Who Have Recovered From Mild COVID-19 in Shanghai, China. JAMA Intern Med. https://doi.org/10.1001/jamainternmed.2020.4616 
160. Wulff, N. H., Tzatzaris, M., & Young, P. J. (2012). Monte Carlo simulation of the Spearman-Kaerber TCID50. J Clin Bioinforma, 2(1), 5. https://doi.org/10.1186/2043-9113-2-5 
161. Xiao, D. A. T., Gao, D. C., & Zhang, D. S. (2020). Profile of Specific Antibodies to SARS-CoV-2: The First Report. J Infect. https://doi.org/10.1016/j.jinf.2020.03.012 
162. Yang, X., Yu, Y., Xu, J., Shu, H., Xia, J., Liu, H., Wu, Y., Zhang, L., Yu, Z., Fang, M., Yu, T., Wang, Y., Pan, S., Zou, X., Yuan, S., & Shang, Y. (2020). Clinical course and outcomes of critically ill patients with SARS-CoV-2 pneumonia in Wuhan, China: a single-centered, retrospective, observational study. Lancet Respir Med, 8(5), 475-481. https://doi.org/10.1016/s2213-2600(20)30079-5 
163. Yau, F., Ferreira, R., Kamali, R., Bird, P. W., Halliwell, R., Patel, H., Nicoara, D. C., Woltmann, G., & Tang, J. W. (2021). Clinical utility of a rapid 'on-demand' laboratory-based SARS-CoV-2 diagnostic testing service in an acute hospital setting admitting COVID-19 patients. Clin Infect Pract, 12, 100086. https://doi.org/10.1016/j.clinpr.2021.100086 
164. Yelin, I., Aharony, N., Shaer Tamar, E., Argoetti, A., Messer, E., Berenbaum, D., Shafran, E., Kuzli, A., Gandali, N., Shkedi, O., Hashimshony, T., Mandel-Gutfreund, Y., Halberthal, M., Geffen, Y., Szwarcwort-Cohen, M., & Kishony, R. (2020). Evaluation of COVID-19 RT-qPCR test in multi-sample pools. Clin Infect Dis. https://doi.org/10.1093/cid/ciaa531 
165. Zhang, Y. V., Wiencek, J., Meng, Q. H., Theel, E. S., Babic, N., Sepiashvili, L., Pecora, N. D., Slev, P., Cameron, A., Konforte, D., & the, A. C. S. T. T. F. (2021). AACC Practical Recommendations for Implementing and Interpreting SARS-CoV-2 EUA and LDT Serologic Testing in Clinical Laboratories. Clinical Chemistry. https://doi.org/10.1093/clinchem/hvab051 
166. Zhao, J., Yuan, Q., Wang, H., Liu, W., Liao, X., Su, Y., Wang, X., Yuan, J., Li, T., Li, J., Qian, S., Hong, C., Wang, F., Liu, Y., Wang, Z., He, Q., Li, Z., He, B., Zhang, T., . . . Zhang, Z. (2020). Antibody responses to SARS-CoV-2 in patients of novel coronavirus disease 2019. Clinical Infectious Diseases. https://doi.org/10.1093/cid/ciaa344 
167. Zimmer, C., Corum, J., Wee, S., & Kristoffersen, M. (2022, August 31). Coronavirus Vaccine Tracker. https://www.nytimes.com/interactive/2020/science/coronavirus-vaccine-tracker.html

Coding Section

CPT	Code Description
86318	Immunoassay for infectious agent antibody(ies), qualitative or semiquantitative, single step method (e.g., reagent strip)
86328	Immunoassay for infectious agent antibody(ies), qualitative or semiquantitative, single step method (e.g., reagent strip); severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19])
86408	Neutralizing antibody, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID19]); screen
86409	Neutralizing antibody, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID19]); titer
86413	Severe acute respiratory syndrome coronavirus 2 (SARSCoV-2) (Coronavirus disease [COVID-19]) antibody, quantitative
86769	Antibody; severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19])
86790	Antibody: virus, not elsewhere specified
87426	Infectious agent antigen detection by immunoassay technique, (e.g., enzyme immunoassay [EIA], enzyme-linked immunosorbent assay [ELISA], immunochemiluminometric assay [IMCA]) qualitative or semiquantitative, multiple-step method; severe acute respiratory syndrome coronavirus (e.g., SARS-CoV, SARS-CoV-2 [COVID-19])
87428	Infectious agent antigen detection by immunoassay technique, (e.g., enzyme immunoassay [EIA], enzyme-linked immunosorbent assay [ELISA], fluorescence immunoassay [FIA], immunochemiluminometric assay [IMCA]) qualitative or semiquantitative; severe acute respiratory syndrome coronavirus (e.g., SARS-CoV, SARS-CoV-2 [COVID-19]) and influenza virus types A and B
87631	Infectious agent detection by nucleic acid (DNA or RNA); respiratory virus (e.g., adenovirus, influenza virus, coronavirus, metapneumovirus, parainfluenza virus, respiratory syncytial virus, rhinovirus), includes multiplex reverse transcription, when performed, and multiplex amplified probe technique, multiple types or subtypes, 3 – 5 targets
87632	Infectious agent detection by nucleic acid (DNA or RNA); respiratory virus (e.g., adenovirus, influenza virus, coronavirus, metapneumovirus, parainfluenza virus, respiratory syncytial virus, rhinovirus), includes multiplex reverse transcription, when performed, and multiplex amplified probe technique, multiple types or subtypes, 6 – 11 targets
87633	Infectious agent detection by nucleic acid (DNA or RNA); respiratory virus (e.g., adenovirus, influenza virus, coronavirus, metapneumovirus, parainfluenza virus, respiratory syncytial virus, rhinovirus), includes multiplex reverse transcription, when performed, and multiplex amplified probe technique, multiple types or subtypes, 12-25 targets
87635	Infectious agent detection by nucleic acid (DNA or RNA); severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19]), amplified probe technique
87797	Infectious agent detection by nucleic acid (DNA or RNA), not otherwise specified; direct probe technique, each organism
87798	Infectious agent detection by nucleic acid (DNA or RNA), not otherwise specified; amplified probe technique, each organism
87799	Infectious agent detection by nucleic acid (DNA or RNA), not otherwise specified; quantification, each organism
87811	Infectious agent antigen detection by immunoassay with direct optical (i.e., visual) observation; severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19])
0115U	Respiratory infectious agent detection by nucleic acid (DNA and RNA), 18 viral types and subtypes and 2 bacterial targets, amplified probe technique, including multiplex reverse transcription for RNA targets, each analyte reported as detected or not detected Proprietary test: ePlex Respiratory Pathogen (RP) Panel Lab/Manufacturer: GenMark Diagnostics, Inc
0202U	Infectious disease (bacterial or viral respiratory tract infection), pathogen specific nucleic acid (DNA or RNA), 22 targets including severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), qualitative RT-PCR, nasopharyngeal swab, each pathogen reported as detected or not detected Proprietary test: BioFire® Respiratory Panel 2.1 (RP2.1) Lab/Manufacturer: BioFire®Diagnostics, LLC
0223U	Infectious disease (bacterial or viral respiratory tract infection), pathogen-specific nucleic acid (DNA or RNA), 22 targets including severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), qualitative RT-PCR, nasopharyngeal swab, each pathogen reported as detected or not detected Proprietary test: QIAstat-Dx Respiratory SARS CoV-2 Panel Lab/Manufacturer: QIAGEN Sciences
0224U	Antibody, severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19]), includes titer(s), when performed Proprietary test: COVID-19 Antibody Test Lab/Manufacturer: Mt Sinai, Mount Sinai Laboratory
0225U	Infectious disease (bacterial or viral respiratory tract infection) pathogen-specific DNA and RNA, 21 targets, including severe acute respiratory syndrome coronavirus 2 (SARSCoV-2), amplified probe technique, including multiplex reverse transcription for RNA targets, each analyte reported as detected or not detected Proprietary test: ePlex® Respiratory Pathogen Panel 2 Lab/Manufacturer: GenMark Dx/GenMark Diagnostics, Inc
0226U	Surrogate viral neutralization test (sVNT), severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19]), ELISA, plasma, serum Proprietary test: Tru-Immune™ Lab/Manufacturer: Ethos Laboratories/GenScript® USA Inc
C9803	Hospital outpatient clinic visit specimen collection for severe acute respiratory syndrome coronavirus 2 (sars-cov-2) (coronavirus disease [covid-19]), any specimen source
G2023	Specimen collection for severe acute respiratory syndrome coronavirus 2 (sars-cov-2) (coronavirus disease [covid-19]), any specimen source
G2024	Specimen collection for severe acute respiratory syndrome coronavirus 2 (sars-cov-2) (coronavirus disease [covid-19]) from an individual in a SNF or by a laboratory on behalf of a HHA, any specimen source
U0001	CDC Novel Coronavirus (2019-nCoV) Real-Time RT-PCR Diagnostic Panel
U0002	Non-CDC laboratory test for 2019-nCoV (COVID-19), any method
U0003	Infectious agent detection by nucleic acid (DNA or RNA); severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19]), amplified probe technique, making use of high throughput technologies as described by CMS-2020-01-R
U0004	2019-nCoV Coronavirus, SARS-CoV-2/2019-nCoV (COVID-19), any technique, multiple types or subtypes (includes all targets), non-CDC, making use of high throughput technologies as described by CMS-2020-01-R
U0005	Infectious agent detection by nucleic acid (DNA or RNA); severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) (Coronavirus disease [COVID-19]), amplified probe technique, CDC or non-CDC, making use of high throughput technologies, completed within two calendar days from date and time of specimen collection. (List separately in addition to either HCPCS code U0003 or U0004)

Procedure and diagnosis codes on Medical Policy documents are included only as a general reference tool for each Policy. They may not be all-inclusive.

This medical policy was developed through consideration of peer-reviewed medical literature generally recognized by the relevant medical community, U.S. FDA approval status, nationally accepted standards of medical practice and accepted standards of medical practice in this community, Blue Cross Blue Shield Association technology assessment program (TEC) and other non-affiliated technology evaluation centers, reference to federal regulations, other plan medical policies, and accredited national guidelines.

"Current Procedural Terminology © American Medical Association. All Rights Reserved" 

History From 2023 Forward     

03/07/2023

New Policy