Two different types of detectable antibody responses in SARS-CoV-2 (COVID-19) tell very different stories and may indicate ways to enhance public health efforts against the disease, according to researchers at The University of Texas MD Anderson Cancer Center.
Antibodies to the SARS-CoV-2 spike protein receptor binding domain (S-RBD) are speculated to neutralize virus infection, while the SARS-CoV-2 nucleocapsid protein (N-protein) antibody may often only indicate exposure to the virus, not protections against reinfection.
The results, published today in JCI Insight, highlight findings from a quantitative serological enzyme-linked immunosorbent assay (ELISA) using SARS-CoV-2 S-RBD and N-protein for the detection of circulating antibodies in 138 serial serum samples from confirmed COVID-19 hospitalized patients and 464 healthy and non-COVID-19 serum samples that were collected between June 2017 and June 2020.
Results showed that 3% of healthy and non-COVID-19 samples collected during the pandemic in Houston were positive for the N-protein antibody, but only 1.6% of those had the S-RBD antibody.
Of samples with the S-RBD antibody, 86% had neutralizing capacity – meaning they could prevent reinfection of COVID-19, but only 74% of samples with N-protein had neutralizing capacity. When positive for both, 96.5% exhibited neutralizing capacity.
“These findings suggest that detection of N-protein binding antibodies does not always correlate with presence of S-RBD neutralizing antibodies, and that the presence of the S-RBD antibody is the best indicator of any potential protection against reinfection,” said senior author Raghu Kalluri, M.D., Ph.D., professor and chair of Cancer Biology.
“We caution against the extensive use of N-protein based serology testing for determination of potential COVID-19 immunity, and we believe that accurate and reliable S-RBD serological testing is needed to carefully identify individuals with neutralizing antibodies in order to help advance recovery efforts around the globe.”
At present, some commercially available serological tests confirm only the presence antibodies to the N-protein, with over 200 commercial and hospital laboratory testing facilities currently using these tests. While these tests indicate exposure to the virus, they do not seem to suggest immunity to reinfection.
These findings reiterate the need to educate on what an antibody test result mean for each patient, and that public health efforts should focus on ways to encourage patients to continue vigilant safety precautions even with the presence of N-protein antibodies.
“In addition to serological assessment of the general population, we are hopeful these results will aid in rapid assessment of the efficacy of vaccine candidates as they are translated into the broader population,” said lead author Kathleen McAndrews, Ph.D., postdoctoral fellow in Cancer Biology.
Coronaviral genome and structure
CoV belong to the subfamily Coronavirinaein the family of Coronaviridae of the order Nidovirales. In this subfamily four genera are included Alphacoronavirus, Betacoronavirus, Gammacoronavirus and Deltacoronavirus.
The genome of the virus is a single-stranded positive-sense RNA (+ssRNA) (around 30 kb in size) with a 5′-cap structure and 3′-poly-A tail. The genome and subgenomes of a typical CoV may present six, or even more, open reading frames (ORF). The first ORF (ORF1a/b) encompasses approximately 66% of the whole genome and encodes 16 non-structural proteins (nsp1–16), which are mainly involved in the replication of CoV.
Other ORF encompassing one-third of the genome near the 3′-terminus encode the main structural proteins: spike (S), membrane (M), envelope (E) and nucleocapsid (N) proteins (Chen et al., 2020a).
The different CoV exhibit 54% identity of the whole RNA, with 58% exhibiting identity for the non-structural proteins coding region and 43% identity for the structural protein coding region. Sequence analysis shows that the new CoV incorporates the typical genome structure of CoV and belongs to the cluster of betac-CoV that includes bat-SARS-like (SL)-ZC45, bat-SL ZXC21, SARS-CoV and Middle East respiratory syndrome coronavirus (MERS-CoV). Based on the phylogenetic tree of CoV, 2019-nCoV is more closely related to bat-SL-CoV ZC45 and bat-SL-CoV ZXC21 and more distantly related to SARS-CoV (Chen et al., 2020a).
Four principal structural proteins are essential for assembly of the virion and its associated infective capacity. Homotrimers of S proteins make up the spikes on the viral surface, which are responsible for attachment to receptors on the host cells.
The M protein has three transmembrane domains and shapes the virions, promotes membrane curvature and covers the nucleocapsid. The E protein participates in virus assembly and release, and is involved in viral pathogenesis. The N protein presents two domains, both of which can bind virus RNA genome via different mechanisms.
The N protein binds to non-structural protein 3 (nsp3) protein to help tether the genome to the replication–transcription complex and package the encapsidated genome into virions. The N protein is also an antagonist of interferon and viral encoded repressor of RNA interference, which may be beneficial for viral replication.
Diagnostic tests for the SARS-CoV-2
On 22 May 2019, the database held by the Foundation for Innovative New Diagnostics, which is the WHO Collaborating Centre for Laboratory Strengthening and Diagnostic Technology Evaluation, contained 560 SARS-CoV-2 laboratory tests for the diagnosis of COVID-19.
These comprised 273 molecular assays and 287 immunoassays. Excluding those intended for research use only, 152 of these are molecular assays and 211 immunoassays are CE-marked for in-vitro diagnostic devices. There are principally two types of tests available for COVID-19: viral tests and antibody tests.
The viral tests are direct tests as they are designed to detect the virus and therefore reflect current infection. In contrast, the antibody tests are indirect tests, as they do not detect the virus, but rather ascertain established seroconversion to previous infection, or early seroconversion to ongoing infection.
The recommended test for diagnosis of SARS-CoV-2 infection involves detection of viral RNA using nucleic acid amplification tests (NAAT), such as reverse transcription (RT)-PCR (www.ecdc.europa.eu). In areas with widespread community transmission of SARS-CoV-2 and when laboratory resources are limited, detection by RT-PCR of a single discriminatory target is considered sufficient.
There are still, however, specific technical considerations for laboratory testing, including specimen collection (variable collection methods), which samples to collect (upper or lower respiratory tract biospecimens, or other samples), time of collection in relation to the course of disease, and the availability of different laboratory test methods and kits (not all of which may be standardized or approved by authorities such as the US Food and Drug Administration).
Then there are infrastructure considerations: are the approved laboratory facilities and trained manpower available, can the methodology be rapidly scaled up, and how are test results interpreted and false negatives excluded?
These issues have been faced by the whole scientific community, with a collective response to develop guidance. The currently used protocol was developed and optimized for the detection of the novel CoV at the Charité University Hospital, Geneva, Switzerland in collaboration with several other laboratories in Germany, the Netherlands, China, France, the UK and Belgium (Corman et al., 2020).
Additionally, the existing protocol was further optimized by the Centers for Disease Control (CDC) in the USA through the comprehensive comparison and validation of alternative kits available for nucleic acid extraction and the use of alternative probe and primer sets for efficient SARS-CoV-2 detection in clinical samples (www.cdc.gov/coronavirus).
Similar approaches are being undertaken by other national authorities as they continue to scale up provision for laboratories not using CE-marked assays (www.england.nhs.uk/coronavirus/).
The importance and variability of specimen collection was initially highlighted from comparison of positive rates from pharyngeal, nasal, blood, sputum, faeces, urine and bronchoalveolar lavage fluid specimens and fibrobronchoscope brush biopsy of patients with confirmed COVID-19 (Zou L et al., 2020).
At present the CDC recommend collecting and testing an upper respiratory specimen, with a nasopharyngeal specimen being the preferred choice for swab-based SARS-CoV-2 testing. When collection of a nasopharyngeal swab is not possible, the following are acceptable alternatives: an oropharyngeal specimen, a nasal mid-turbinate specimen (using a flocked tapered swab), an anterior nares (nasal swab) specimen (using a flocked or spun polyester swab) or a nasopharyngeal wash/aspirate or nasal aspirate specimen.
For individuals having invasive procedures, lower respiratory tract specimens are also recommended if available. Although the virus can be detected in other specimens, such as blood and stools, these have been generally less reliable than respiratory specimens.
At present it is recommended that specimens should be collected as soon as possible once a decision has been made to pursue SARS-CoV-2 testing, regardless of the time of symptom onset. The viral load in throat swabs is greatest at the time of viral onset and decreases monotonically thereafter (To et al., 2020; Zou L et al., 2020).
Analysis of these temporal dynamics suggests that viral shedding may begin 2–3 days before the appearance of the first symptoms, facilitating pre-symptomatic or asymptomatic transmission (He et al., 2020). CoV have a number of molecular targets within their +ssRNA genome that can be used for RT-PCR assays.
The WHO has provided primers for the genes that encode the structural proteins of the viral envelope (E) and the nucleocapsid (N), and for the RNA-dependent RNA polymerase (RdRp), which is a key part of the virus’s replication machinery that makes copies of its RNA genome (Corman et al., 2020).
However, there has been no demonstration that any one of these three sequences (E, N or RdRP) might offer an advantage for clinical diagnostic testing, with different targets being preferred by different authorities. For example, the Public Health England assay employs two probes against RdRp; one is a Pan Sarbeco-probe that will detect 2019-nCoV, SARS-CoV and bat-SARS-related CoV, while the second probe is specific to 2019-NCoV. Continued refinement of these NAAT assays is ongoing to facilitate their upscaling, while maintaining laboratory safety, a low cost and a high sensitivity (Won et al., 2020).
Detection of isolated viral antigens
Great efforts have been made in order to develop tests for the rapid detection of SARS-CoV-2 antigens. Antigen detection tests are designed to directly detect viral particles in biological samples such as nasopharyngeal secretions. Several rapid antigen tests have been proposed (Diao et al., 2020); however, the principal concerns are the false-negative rate due to either a low or variable viral load, and the variability in sampling, the latter having the potential to further compound the problem in cases with low viral titres, thereby increasing the false-negative rate (Tang YW et al., 2020).
Diao and colleagues (Diao et al., 2020) have reported preliminary results from the use of a fluorescence immunochromatographic assay for detecting the N protein of SARS-CoV-2 in both nasopharyngeal swab samples and urine from 239 participants, with comparison to NAAT testing where the intersection of the amplification curve and diagnostic threshold line (cycle threshold [Ct] value) was set at either ≤30 or ≤40 (Diao et al., 2020).
With a higher viral load in the sample, the prespecified Ct value may be lower, as fewer replication cycles are required to achieve a detectable signal; however, with a low viral load, a greater number of replication cycles (higher Ct value) will be required for a detectable signal to be attained.
For this assay with a prevalence of 87%, although the positive predictive value (PPV) was 100%, the negative predictive value (NPV) was 32% for a Ct ≤40, increasing to 97% for patients with a higher viral load as demonstrated by a Ct ≤30. This would suggest that, at present, this assay would only be useful in excluding those with high viral loads.
Whether alternative approaches as previously suggested for influenza viruses in children including the use of colloidal gold-labelled immunoglobulin (Ig) G as the detection reagent (Li et al., 2020), to increase the sensitivity of rapid antigen tests for respiratory viruses, are feasible is still under consideration, with monoclonal antibodies specifically against SARS-CoV-2 under development.
Further validation of this technique and similar approaches in larger populations including asymptomatic cases is warranted. Consideration of approaches to try to concentrate antigen and amplify the detection phase are, however, likely to be needed for these methods to have any clinical utility (Loeffelholz et al., 2020).
At the time of writing (25 April 2020), the non-governmental organization Foundation for Innovative New Diagnostics (https://www.finddx.org/) has listed four CE-marked rapid SARS-CoV-2 antigen detection tests, which are primarily lateral flow immunochromatographic assays based on the presence of a colloid gold conjugate pad and a membrane strip pre-coated with antibodies specific to SARS-CoV-2 antigens on a test line.
If SARS-CoV-2 antigens are present in the specimen withdrawn from a nasopharyngeal swab, a visible band appears on the test line as antibody–antigen–antibody gold conjugate complex forms. The evaluation of these diagnostic tests has, however, been limited, and their CE mark means that the manufacturers state that they conform with the relevant EU legislation, but they may still not be available to purchase.
According to European Union Directive 98/79/EC for in-vitro diagnostic devices, in order to affix the CE mark to COVID-19 diagnostic devices to be used by health professionals, the manufacturer has to specify device performance characteristics and self-declare conformity with the safety and performance requirements listed in the Directive. In addition, self-tests intended to be used by patients themselves must also be assessed by a third-party body (a notified body), which for these tests has yet to happen.
Although direct antigen tests are being registered by several health authorities, the sensitivity of these tests is lower than that of RT-PCR, with previous antigen-detecting enzyme-linked immunosorbent assays (ELISA) developed for SARS-CoV having limits of detection of 50 pg/ml (Che et al., 2004; Di et al., 2005).
Furthermore, clarification of their specificity for SARS-CoV-2 is awaited, given the potential for cross-reaction with other human CoV. Despite these limitations, the chief advantages of antigen tests, including their rapidity (10–30 min compared with hours for NAAT testing), ease of interpretation and limited technical skill and infrastructure required compared with NAAT-based testing, continue to make them worth pursuing.
However, experience with influenza antigen testing invites caution as these tests may have low sensitivity and specificity; moreover, as noted, false–negatives rate will be critical (Tang YW et al., 2020). Their greatest utility if they come to fruition may be in symptomatic patients, when the viral load will be at its greatest, to enable accurate triage.
Building an indirect test for SARS-CoV-2: serological testing
In contrast to NAAT-based testing, where as soon as the sequence is known, a diagnostic test can be built, the diagnostic technology and methodology underlying the development of serological tests is quite different, with a substantially longer timeline to obtain a robust product that is suitable for routine deployment.
The principal difference is that antibody tests require identification of distinct proteins that form the viral coat, with elucidation of which proteins are most divergent from previous CoV proteins, then identification of specific antibodies to these proteins that are part of the acquired immune response to viral exposure, and finally testing to ensure that there is limited cross-reactivity with antibodies developed to other historical CoV.
With the previous two CoV, a variety of assays encompassing different methodologies were developed, including ELISA, chemiluminescence, western blotting, protein microarray and immunofluorescence platforms. However, only ELISA and chemiluminescence were deemed suitable for clinical application because of costs, time-to-results, relative simplicity and ability to scale to very large throughput. It is these platforms that are once again being examined for detection of antibodies to SARS-CoV-2.
Appraisal of test performance
Appropriate thresholds for sensitivity and specificity of an antibody test depend on its purpose and must be considered prior to implementation. For diagnosis in symptomatic patients, high sensitivity is required (generally ≥90%). In this context, a slight reduction in specificity may be acceptable as some false-positive results may be tolerated, provided other potential diagnoses are considered and there is acceptance that overdiagnosis may result in unnecessary interventions that, for SARS-CoV-2, may include quarantining.
However, if antibody tests were deployed as an individual-level approach to inform release from social isolation and return to normal activities, high specificity would be essential, as false-positive results would return non-immune individuals to risk of exposure. It is with these purposes in mind that the UK Medicines and Healthcare products Regulatory Agency set a minimum 98% specificity threshold for flow immunoassays (LFIA).
This is particularly challenging, particularly given the scale of validation study required for a suitable candidate LFIA, because to demonstrate a high specificity if the true underlying value were 98%, 1000 negative controls would be required to estimate the specificity of an assay to ±1% with approximately 90% power.
As part of the evaluation of test performance, the influence of population prevalence also needs to be considered, acknowledging that this is at present rapidly changing (Brenner and Gefeller, 1997). This can be considered as the proportion of all positive tests that are wrong, as well as the number of incorrect positive tests per 1000 people tested.
For example, for a point of care (POC) test with 70% sensitivity and 98% specificity, the proportion of positive tests that are wrong is 35% at 5% population seroprevalence (19 false positives/1000 tested), 13% at 20% seroprevalence (16 false positives/1000 tested) and 3% at 50% seroprevalence (10 false positives/1000 tested).
According to available data, the prevalence of seropositivity is still low. The prevalence of antibodies to SARS-CoV-2 among a high risk category such as healthcare personnel is 5.9% in Utah, USA (Masden et al., 2020), 5.4% in Lyon, France (Solodky et al., 2020), 17.3% in Trieste (Comar et al., 2020), 5.25% in Padua (Tosato et al. 2020) and 1.5% in Bari, Italy (Paradiso et al., 2020a), 1.6% in Germany (Korth et al., 2020) and 2.6% in Barcelona, Spain (Tuaillon et al., 2020). In the general population it has been reported as being 0.13% in Rio Grand do Sul, Brazil (Silveira et al., 2020), 1.5% in Santa Clara, California (Benavid et al. 2020), 1.79 % in Idaho (Bryan et al., 2020) and 7.1% in Atlanta, USA (Zou J et al., 2020), 1.2% in Edinburgh, Scotland (Thompson et al., 2020), 3% in Paris, France (Grzelak et al., 2020), 1.7% in Denmark (Erikstrup et al., 2020), 3.3% in Kobe, Japan (Doi et al., 2020), 9.6% in Wuhan, China (Wu et al., 2020) and 21% in Guilan, Iran (Shakiba et al., 2020). Large-scale seroprevalence studies are ongoing, but understanding the background rate is essential for accurate interpretation of diagnostic tests.
The potential risk of a test providing false reassurance and release from being sheltered for non-immune individuals can therefore be widely based on the underlying seroprevalence, and this still assumes antibody positivity as a correlate of protective immunity, which may be incorrect.
Dynamics of seroconversion
Understanding viral and host interactions during the acute and convalescent phases is critical to be able to understand both the timing of initial seroconversion after exposure to SARS-CoV-2 and the subsequent duration of antibodies. However, at present the studies regarding seroconversion are being developed in parallel to the assays, limiting some conclusions.
The data do suggest that seroconversion after exposure to SARS-CoV-2 is very similar to that after other acute viral infections, with IgG concentration beginning to rise as IgM concentrations reach a plateau (Figure 1 ). However, observations have shown that IgM and IgA growth is relatively slow related to other respiratory viruses, which has been suggested to contribute to the heterogeneous pathogenicity of SARS-CoV-2 in COVID-19 patients (Zhao J et al., 2020).
The most comprehensive study to date of seroconversion assessed 173 patients affected by COVID-19 using an assay developed to detect antibodies against the receptor binding domain of the S protein of SARS-CoV-2 (Zhao J et al., 2020). The median seroconversion times of total antibody, IgM and IgG were 11, 12 and 14 days (Zhao J et al., 2020).
The respective seroconversion rates for total antibody, IgM and IgG were 93.1%, 82.7% and 64.7% (Zhao J et al., 2020), with the cumulative seroconversion curve suggesting that the rate for total antibody and IgM reached 100% 30 days after the onset. These studies have also highlighted the temporal nature of testing as, despite all patients subsequently being confirmed as COVID-19 positive, in the early phase of illness (within 7 days since the onset) the NAAT test only exhibited 66.7% sensitivity, with the antibody assays having an even lower positive rate of 38.3% (Zhao J et al., 2020).
However, the sensitivity of antibody overtook that of RNA testing from day 8 after symptom onset and reached over 90% across day 12 after onset. Among samples from patients in the later phase (day 15–39 after onset), the sensitivities for total antibody, IgM and IgG were 100.0%, 94.3% and 79.8%, respectively.
In contrast, RNA was detectable in only 45.5% of samples from days 15–39. In a separate small series of nine cases, seroconversion occurred after 7 days in 50% of patients (after 14 days in all) but was not followed by a rapid decline in viral load (Wolfel et al., 2020). An analysis of 285 patients further supports IgG seroconversion within 19 days after symptom onset (Long et al., 2020).
Collectively, these data suggest that there is a role for both tests depending on where the patient is on their infection journey, with the combined use of NAAT and antibody tests markedly improving the sensitivity of a pathogenic diagnosis for COVID-19 patients in different phases of the illness.
With respect to antibody titres and disease severity, critically ill hospitalized patients have been reported to exhibit significantly higher antibody titre values than non-critical patients in some (Long et al., 2020; Zhao J et al., 2020) but not all studies. In the previous epidemics of SARS-CoV and MERS-CoV, antibody titres were positively associated with disease severity (Choe et al., 2017; Okba et al., 2019).
In a limited case series (n = 57 confirmed SARS-CoV-2 cases), six patients with detectable viral RNA in the blood were at increased risk of severe disease progression compared with those with low titres, but unfortunately the authors did not measure antibody titres (Chen W et al., 2020b). Clarification is awaited of whether even in previously healthy individuals a high viral titre and/or high antibody titre can predict disease severity and likely progression.
Diagnostic performance of the immunoassays
Our extensive search identified 24 peer-reviewed articles (Xiao DAT et al., 2020; Zhao J et al., 2020; Du Z et al., 2020; Guo L et al., 2020; Jin Y et al., 2020; Pan Y et al., 2020; Padoan A et al., 2020; Zhong L et al., 2020; Infantino M et al., 2020; Xiang F et al., 2020; Long QX et al., 2020; Perera R et al., 2020; Qu J et al., 2020; Zhao R et al., 2020; Cai X et al., 2020; Hou H et al., 2020; Lippi G et al., 2020; Sun B et al., 2020; To K et al., 2020; Xie J et al., 2020; Bryan A et al., 2020; Jaaskeilanen A et al., 2020; Montesinos I et al., 2020; Tang MS et al., 2020) and 25 pre-print studies (Wang X et al., 2020; Lassauniere R et al., 2020; Yangchun F, 2020; Liu R et al., 2020; Lin D et al., 2020; Lou B et al., 2020; Liu L et al., 2020; Jia X et al., 2020; Zhang J et al., 2020; Xiang J et al., 2020; Hu Q et al., 2020; Ma H et al., 2020; Qian C et al., 2020; National COVID Testing Scientific Advisory Panel, 2020; Burbelo PD et al., 2020; Adams ER et al., 2020; Meyer B et al., 2020; Norman M et al., 2020; Tuaillon E et al., 2020; Wajnberg A et al., 2020; Wan Y et al., 2020; Xiao T et al., 2020; Zhou Q et al., 2020; Ozturk T et al., 2020; Rosado J et al., 2020) reporting on the sensitivity and specificity of immunoassays for COVID-19, with a sample size ranging from 16 to 6001 subjects (Table 1). Most studies were conducted in China, with only a few coming from western countries.
Overall sensitivity ranged from 0% to 100% and specificity from 78% to 100%, with performance highly time-sensitive, reflecting the dynamics of seroconversion. In general, most assays performed better shortly after initial symptom resolution, accepting the very limited time frames evaluated for all studies to date.
In an evaluation of nine commercially available SARS-CoV-2 immunoassays, the sensitivities varied with the duration of disease: early phase, 7–13 days after the onset of disease symptoms (sensitivities 40–86%); middle phase, 14–20 days after the onset of disease symptoms (sensitivities 67–100%); and late phase, ≥21 days after the onset of disease symptoms (sensitivities 78–89%) (Lassauniere et al., 2020).
The range of assays being released is extensive, with apparently very limited validation. Gonzalez and colleagues reviewed four web databases for SARS-CoV-2 immunoassay, and by 4 April 2020, 226 immunoassays from 20 different countries had already been listed. The technical data sheet was available online for only 22% of the tests, and despite 23 claiming regulatory certification only four had PubMed-listed papers (González JM et al., 2020).
Despite wide claims for sensitivity and specificity, practically at present it is almost impossible to conclude which antibody test would be the one to use. A pragmatic choice would be to use an automated immunoassay that is scalable, from a well-known established manufacturer, with a complete and clear technical data sheet, which has received regulatory certification issued by the health authority and been independently validated.
In accordance with this, the most recent novel assays use fully automated chemiluminescence immunoassays implemented on high-throughput laboratory instrumentation. These systems include the MAGLUMI 2000 Plus 2019-nCov IgM and IgG assays (Snibe, China), which has been independently validated in accordance with the Clinical and Laboratory Standards Institute EP15-A3 guideline (Padoan et al. 2020) and the CE-marked Euroimmun, Italy Anti-SARS-CoV-2 IgA and IgG assays, with others, including from Beckman Coulter, Italy for their Access platform and Roche Diagnostics, Italy for their Elecsys platform, under development.
However, in independent validation the EUROIMMUN assay exhibited some cross-reactivity in both ELISA with serum samples from the two seasonal CoV patients (HCoV-OC43) who had previously cross-reacted with the MERS-CoV S1 IgG ELISA (Okba et al., 2019). On comparison of their respective performances for 131 known cases, there was concordance for the IgG assays of only 88% (kappa statistic 0.47; 95% confidence interval [CI] 0.26–0.68).
Despite involving different immunoglobulin classes, an analogous analysis between MAGLUMI 2019-nCoV IgM positive/negative and EUROIMMUN Anti-SARS-CoV-2 IgA positive/negative results yielded an overall concordance of 90% (kappa statistic 0.39; 95% CI 0.14–0.65). The IgG assays also exhibited different concordance during the early phases of symptom onset, with concordance improving 10–21 days after symptom onset.
Further studies with longer timelines and known cases with a range of symptoms will help confirm the alignment of these assays. Inevitably, it is anticipated that there will be an enormous number of studies comparing the available assays, with the advantages and disadvantages of the respective assays being discussed at length.
Rapid serological tests
POC immunoassays have also been developed for the rapid detection of SARS-CoV-2 antibodies (IgG and IgM). The primary advantage of these tests, as with a home pregnancy test, is being able to obtain a diagnosis without sending samples to centralized laboratories.
This enables communities without the necessary laboratory infrastructure to detect SARS-CoV-2-exposed subjects using only finger prick testing rather than formal blood draws, thereby reducing training requirements and allowing clinicians to have a validated test at the bedside. As these devices are cheap to manufacture, store and distribute, and provided that a positive antibody test were confirmed to be an accurate surrogate for immunity to infection, they would also be able inform decision making.
This would particularly be the case as secure confirmation of antibody status would reduce anxiety, provide confidence to allow individuals to relax social distancing measures, and guide policy makers in the staged release of population lockdown, potentially in tandem with digital approaches to contact tracing.
The rapid POC immunoassays are generally LFIA (Li et al., 2020). In lateral flow assays, a membrane strip is coated with two lines: gold nanoparticle–antibody conjugates are located on one line and bind antibodies on the other. The blood sample from the patient is put on the membrane, and the proteins are drawn through the membrane strip by capillary action. As it passes the first line, the antigen binds to the gold nanoparticle–antibody conjugate, and the complex flows across the membrane. Generally, rapid assays have a low diagnostic performance compared with ELISA assays, which is explained not only by the well-known technical differences between the two methodologies, but also by possible low antibody concentrations that may further contribute to the false-negative results observed with the rapid tests.
At present, 12 peer-reviewed articles (Li Z et al., 2020; Cassaniti I et al., 2020; Lee YL, et al., 2020; Shen B et al., 2020; Dohla M et al., 2020; Hoffman T et al., 2020; Imai K et al., 2020; Pan Y et al., 2020b; Spicuzza et al., 2020; Yong G et al., 2020; Demey B et al., 2020; Montesinos I et al., 2020) and 9 pre-print studies (Garcia FP et al., 2020; Lassauniere R et al., 2020; Liu Y et al., 2020; Yong G et al., 2020; Lou B et al., 2020; Bendavid E et al., 2020; Paradiso AV et al., 2020a; National COVID testing Scientific Advisory Board, 2020; Tuaillon E et al., 2020) have reported on the diagnostic performance of rapid assays (summarized in Table 1).
In the published studies, sensitivity and specificity ranged from 9% to 88.6% and from 88.9% to 91.7%, respectively (Table 1), while in the pre-print articles sensitivity and specificity ranged from 30% to 98.8% and from 89% to 100%, respectively. Of note the sensitivity of these tests performed in countries other than China were substantially lower than those reported for studies conducted in China.
Extensive evaluation of manufacturers’ claims related to the performance of these tests and optimal timing will be required before they are suitable for widespread routine clinical use. For example, the performance of the VivaDiag, VivaCheck Biotech, China COVID-19 IgM/IgG Rapid Test was evaluated in 30 cases 7 days after confirmed NAAT testing, and despite this five (16.7%) cases were negative for both IgG and IgM (Cassaniti et al., 2020).
Furthermore, when evaluating 50 patients with acute illness presenting in the emergency room, of whom 38 were positive on RT-PCR, the sensitivity of the VivaDiag COVID-19 IgM/IgG Rapid Test was only 18.4%, its specificity was 91.7%, while the NPV was 26.2% and the PPV was 87.5% (Cassaniti et al., 2020).
The same VivaDiag test was evaluated in 525 healthcare workers in Italy, with only six testing positive; none was positive by NAAT testing or symptomatic, and only three had a confirmed positive result on the MAGLUMI chemiluminescence IgG assay (Paradiso et al., 2020b). Evaluation of six POC tests in a mix of 111 patients with COVID-19, other CoV or other viruses and negative controls revealed sensitivities ranging from 83% to 93% and NPV of 74–92% (Lassauniere et al., 2020).
In keeping with other studies, the diagnostic performance of these tests reflected the duration of the illness, with the worst performance observed in the first 2 weeks after symptom onset (Lassauniere et al., 2020). Finally, formal evaluation of nine commercially available LFIA in a case-control mix of 182 samples revealed sensitives of 55–70% (National COVID Testing Scientific Advisory Panel, 2020).
For all studies to date, sample size has been limited, and further testing across a large diverse population from a range of geographical locations and ethnic groups is required, with inclusion of children and individuals with autoimmune disease and immunosuppression. With extensive evaluation, it is likely that technical performance may deteriorate. At present, evaluation of the current LFIA devices suggest that although they may provide some information for population-level surveys, their performance is inadequate for most individual patient applications.
Clinical interpretation of the COVID-19 tests
The interpretation of a test for SARS-CoV-2 will depend on a combination of the accuracy of the test and the estimated risk of COVID-19 prior to performing the test (Watson et al., 2020). A positive direct antigen test and specifically NAAT is strongly suggestive of current infection due to its high specificity but moderate sensitivity, and the patient can be reassured that the clinician is confident that they have COVID-19 and should be managed in accordance with local policies regarding positive cases.
In contrast, negative tests need to be interpreted with caution, and a single negative SARS-CoV-2 test in a patient with strongly suggestive symptoms should not be relied upon to exclude COVID-19. In this situation, it would still be safer for the patient to be treated as a positive case, and local policies regarding re-testing and isolation to be followed.
For the serological tests, the clinical implication of seroconversion with respect to future immunity continues to be elucidated, but similar principles for evaluating the test result in the clinical context and history of previous infection or exposure is critical, particularly as a false-positive result could lead to false reassurance and inappropriate behaviour that might enhance community disease transmission.
Summary of the original articles reporting on SARS-CoV-2 antibody testing
|Author, Year||Design of the study||N||Population||Country of the test population||Antibody used||Methodology||Main findings and/or conclusions||Sensitivity (%)||Specificity (%)||PPV||NPV|
|Li Z et al., 2020||Retrospective||525||397 RNA-positive patients, 128 controls||China||Commercial assay||Jiangsu Medomics Medical Technologies, LFIA||The test time was from day 8 to day 33 after symptoms appeared. The IgM–IgG combined assay has better utility and sensitivity than a single IgM or IgG test. Results demonstrate that the IgG–IgM combined antibody test kit can be used as a POC test||88.66||90.63||NA||NA|
|Xiao D et al., 2020||Prospective||34||SARS-CoV-2 confirmed patients||China||Commercial assay||Shenzhen Yahuilong Biotechnology, chemiluminescence assay||After 2 weeks from the onset of symptom, all but two subjects had positive results from the test. From the 5th to 7th weeks IgM became negative, while all had high levels of IgG||94.1||NA||NA||NA|
|Zhao J et al., 2020||Prospective||535 samples from 173 subjects||173 RNA-positive patients||China||Commercial assay||Beijing Wantai Biological Pharmacy Enterprise, ELISA||The seroconversion rates for Ab, IgM and IgG were 93.1%, 82.7% and 64.7%. The cumulative seroconversion curve showed that the rates for antibody and IgM reached 100% around 1 month of illness||100 (>15 days)||NA||NA||NA|
|Du Z et al., 2020||Retrospective||60||Convalescent patients (6–7 weeks from onset)||China||Commercial assay||ELISA||All patients tested positive for IgG against the virus, while 13 patients tested negative for IgM||78 IgM|
|Cassaniti I et al., 2020||Prospective||110||30 RNA-positive patients, 50 patients with respiratory symptoms, 30 controls||China||Commercial assay||Rapid VivaDiag IgM /IgG immunoassay||The rapid test is not recommended for triage of patients with suspected COVID-19 in the emergency room||18.4||91.7||87.5||26.2|
|Guo L et al., 2020||Prospective||208 samples from 140 subjects||82 confirmed and 58 probable cases||China||In-house assay||ELISA for IgA, IgM and IgG||IgA, IgM and IgG were detected in 92.7%, 85.4% and 77.9% of samples from a median time of 5 days from the onset of symptoms||75.6 (IgM in confirmed cases)|
93.1 (IgM in probable cases)
|Jin Y et al., 2020||Retrospective||76||43 RNA-positive patients, 33 probable cases||China||Commercial assay||Shenzhen YHLO Biotech, chemiluminescence assay||Viral serological testing is an effective means of diagnosing SARS-CoV-2 infection. The positive rate and titre variance of IgG are higher than those of IgM||48.1 IgM|
|Pan Y et al., 2020||Retrospective||105||105 patients||China||In-house assay||Immunochromatography||The positive rates of Ig in the early stage are relatively low and gradually increase during disease progression. The IgM-positive rate rose from 11.1% in early-stage to 74.2% in late-stage disease. The IgG-positive rate in confirmed patients was 3.6% in early-stage and 96.8% in late-stage disease||68.6||NA||NA||NA|
|Padoan A et al., 2020||Retrospective||87 samples from 37 subjects||37 patients||Italy||Commercial assay||Snibe, MAGLUM 2000 Plus 2019-nCov IgM and IgG assays||After the 11th day from the onset of symptoms, all patients were found to be positive for IgG (100%), while the higher positivity of IgM (88%) was achieved only after the 13th day. The imprecision and repeatability of the test were acceptable||88 IgM|
|Zhong L et al., 2020||Cross-sectional||347||47 RNA-positive patients, 300 controls||China||Commercial assay||ELISA and chemiluminescence detection assay||The ELISA and chemiluminescence methods were consistent in detecting IgG and IgM antibodies by the recombinant N and S proteins of SARS-CoV-2||97.9 IgM|
|Infantino M et al., 2020||Cross-sectional||125||61 RNA-positive patients and 64 controls||China||Commercial assay||iFlash CLIA, Chemiluminescence assay||The ROC AUC were 0.918 and 0.980 for IgM and IgG anti-SARS CoV-2 antibodies||73.3 (IgM)|
|Xiang F et al., 2020||Retrospective||216 samples from 109 subjects||85 confirmed and 24 suspected cases||China||Commercial assays||Zhu Hai LivZon Diagnostics, ELISA||The seropositive rate of IgM increased gradually and notably. IgG was increased sharply on the 12th day after onset. Diagnostic performance calculated from samples obtained after 13 days from the onset||77.3 IgM|
|Lee YL, et al., 2020||Retrospective||33 samples from 14 subjects, 28 samples from 28 controls||14 RNA-positive patients and 28 controls||China||Commercial assay||Alltest, Rapid Test||Antibody response varied with different clinical manifestations and disease severity. Patients with symptoms and development of anti-SARSCoV-2 IgM antibodies had a shorter duration of a positive rRT-PCR result and no worsening clinical conditions compared with those without the presence of anti-SARS-CoV-2 IgM antibodies||78.6||100||NA||NA|
|Long QX et al., 2020||Cross-sectional||285 patients||285 RNA-positive patients||China||Commercial assay||Chemiluminescence Bioscience assay||The positive rate of IgG reached 100% around 17–19 days after symptom onset, while the IgM seroconversion rate reached its peak of 94.1% around 20–22 days after symptom onset||94 (IgM)|
|Perera R et al., 2020||Retrospective||51 samples from 24 patients||24 RNA-positive patients||China||In-house assay||ELISA||IgG and IgM were reliably positive after 29 days from illness onset with no detectable cross-reactivity in age-stratified controls||74||100||NA||NA|
|Qu J et al., 2020||Retrospective||347 samples from 41 patients and 38 samples from controls||41 RNA-positive patients and 38 controls||China||Commercial assay||YHLO Biotech, chemiluminescence assay||The majority of the patients developed robust antibody responses between 17 and 23 days after illness onset||87.8 (IgM)|
|Shen B et al., 2020||Prospective||150 patients||150 suspected cases, of whom 97 were RNA positive||China||Commercial assay||Shanghai Outdo Biotech, rapid immunochromatography test||The colloidal gold immunochromatography assay for SARS-Cov-2-specific IgM/IgG antibody shows the potential for a useful rapid diagnosis test for COVID-19||71||96||97||64|
|Zhao R et al., 2020||Retrospective||481||69 affected subjects and 412 controls||China||In-house assay||ELISA||The overall accuracy of the ELISA test was 97.3%||97.5||97.5||NA||NA|
|Cai X et al., 2020||Retrospective||276 samples from 276 subjects, 200 samples from 200 controls||276 RNA-positive patients, and 200 healthy controls||China||In-house assay||Chemiluminescence assay||Combining immunoassay with real-time RT-PCR might enhance the diagnostic accuracy of COVID-19||57.2 (IgM)|
|Dohla M et al., 2020||Prospective||Samples from 49 symptomatic patients||22 RNA-positive and 27 RNA-negative patients||Germany||Commercial assay||Rapid test||The rapid test was substantially inferior to the RT-qPCR testing and should therefore be used for neither individual risk assessment nor decisions on public health measures||36.4||88.9||72.7||63.1|
|Hoffman T et al., 2020||Cross-sectional||Samples from 153 subjects||29 RNA-positive patients and 124 controls||China||Commercial assay||Zhejiang Orient Gene Biotech, rapid COVID test||The test is suitable for assessing previous virus exposure, although negative results may be unreliable during the first weeks after infection||69 (IgM)|
|Hou H et al., 2020||Retrospective||338 subjects||338 RNA-positive patients||China||Commercial Assay||YHLO, ELISA||Quantitative detection of IgM and IgG antibodies against SARS-CoV-2 quantitatively has potential significance for evaluating the severity and prognosis of COVID-19||82.7 (IgM)|
|Imai K et al., 2020||Retrospective||139 samples from 112 patients and 48 controls||112 RNA-positive patients and 48 controls||Artron, Canada||Commercial assay||Artton, One Step IgM/IgG Rapid Test||Immunoassay had low sensitivity during the early phase of infection, and thus immunoassay alone is not recommended for initial diagnostic testing for COVID-19||40||NA||NA||NA|
|Lippi G et al., 2020||Prospective||48 patients||48 RNA-positive patients||Snibe Cehmiluminescence Maglumi, Italy Euroimmun, Italy||Commercial assays||Snibe, Chemiluminescence MAGLUMI; EUROIMMUN, ELISA||The results of MAGLUMI are well aligned with those of the EUROIMMUN test||10 (<5days)|
100 (>10 days)
|Pan Y et al., 2020b||Retrospective||86 samples from 67 cases||67 RNA-positive patients||China||Commercial assay||Zhuhai Livzon Diagnostic, rapid lateral flow assays||Serology may be considered a supplementary approach in clinical diagnosis||11 (<7 days)|
92 (7–14 days)
96 (>14 days)
|Spicuzza et al., 2020||Cross-sectional||41 subjects||27 RNA-positive patients, 7 symptomatic RNA-negative patients and 7 controls||China||Commercial assay||Beijing Diagreat Biotechnologies, rapid lateral flow assay||Antibody test is quite reliable and useful as it has the advantage of being a POC test that gives a response within minutes||83||93||NA||NA|
|Sun B et al., 2020||Cross-sectional||130 samples from 38 patients, 16 samples from 16 controls||38 RNA-positive patients and 16 controls||China||In-house assay||ELISA||IgM and IgG increased gradually after symptom onset and can be used for detection of SARS-CoV-2 infection. Analysis of the dynamics of IgG may help to predict prognosis||75 (after 1 week)|
94.7 (after 2 weeks)
100 (after 3 weeks)
|To K et al., 2020||Cross-sectional||16 patients||16 RNA-positive patients||China||In-house assay||ELISA||Serological assay can complement RT-qPCR for diagnosis||88 (IgM)|
|Xie J et al., 2020||Prospective||56 patients||56 symptomatic patients||China||Commercial assay||YHLO Biological Technology, chemiluminescence assay||A combination of nucleic acid and Ig testing is a more accurate approach for diagnosing COVID-19||93.7 (IgM)|
|Yonh G et al., 2020||Retrospective||76 samples from 38 patients||38 symptomatic patients||China||Commercial assay||Rapid assay GICA kit||Antibody detection could be used as an effective indicator of the virus in the absence of viral RNA||50 (IgM)|
|Bryan A et al., 2020||Cross-sectional||6001 subjects||1020 controls and 125 patients. 4856 subjects from the general population||USA||Commercial assay||Abbott, chemiluminescence SARS-CoV-2 IgG test||This study demonstrates excellent analytical performance of the Abbott SARS-CoV2 test as well as the limited circulation of the virus in the western USA||53.1 (day 7)|
82.4 (day 10)
96.9 (day 14)
100 (day 17)
|Demey B et al., 2020||Prospective||21 subjects||21 RNA-positive patients||France||Commercial assays||Four rapid lateral flow assays||Immunochromatographic tests for detection of the virus may have a role in the diagnosis of COVID-19||9-24 (day 5)|
67–82 (day 10)
100 (day 15)
|Jaaskeilanen A et al., 2020||Retrospective||77 subjects||40 RNA-positive patients and 37 controls||Germany||Commercial assay||EUROIMMUN, ELISA||The median time after onset of symptoms was 12 days (13 patients, range 5–20 days) for detection of IgG and 11 days (24 patients, range 5–20 days) for detection of IgA||NA||91.9 (IgG)|
|Montesinos J et al., 2020||Retrospective||400 subjects||272 controls and 128 RNA-positive patients||Germany||Commercial assays||MAGLUMI, chemiluminescence; EUROIMMUN, ELISA; rapid assay||The sensitivity of the tests increased with time from onset of symptoms||64.3 (MAGLUMI)|
70 (rapid assay)
|Tang MS et al., 2020||Retrospective||201 subjects||48 patients and 153 controls||Abbott USAEuroimmun USA||Commercial assays||Abbott, chemiluminescence assay; EUROIMMUN, ELISA||Both assays have poor sensitivity during the first days of the disease. Abbott tests generally performed better than the EUROIMMUN test||Abbott:|
0 (<3 days)
30 (3–7 days)
47.8 (8–13 days)
93.8 (>14 days)
0 (<3 days)
25 (3–7 days)
56.5 (8–13 days)
85.4 (>14 days)
|Wang X et al., 2020||Prospective study with longitudinal follow-up||117 samples in 70 subjects||Inpatients and convalescent patients||China||In-house assay||Modified cytopathogenic assay||The seropositivity rate reached up to 100.0% within 20 days after onset. Patients with a worse clinical classification had a higher antibody titre||100||NA||NA||NA|
|Garcia PF et al., 2020||Prospective||163||55 RNA-positive patients, 63 RNA-negative patients, 45 controls||China||Commercial Assay||AllTest, COV 19 IgG IgM immunoassay||The sensitivity of the test was 73.9% after 2 weeks from the onset of symptoms||73.9||100||NA||NA|
|Lassauniere R et al., 2020||Cross-sectional||111||30 SARS-CoV-2 patients, 10 healthy controls, 71 patients with respiratory diseases other than SARS-CoV-2||Denmark||Commercial assays||3 ELISA tests and 6 POC lateral flow tests||The diagnostic performance of the commercial assays analysed may vary||65–90 (ELISA)|
|Yangchun F, 2020||Cross-sectional||294||186 RNA-positive patients, 98 RNA-negative patients||China||Commercial assay||ELISA||Antibody testing has a very good diagnostic performance in identifying positive subjects||96.1 (IgG)||92.4 (IgG)||96.09 (IgG)||90.1 (IgG)|
|Liu R et al., 2020||Retrospective||133||Samples from patients||China||Commercial Assay||YHLO, IG detection kit||In symptomatic patients, IgM was superior to RT-PCR in detecting affected subjects. The positive rate for IgM was 79.55% in moderate cases, 82.69% 156 in severe cases and 72.97% in critical cases. The IgG antibody test positive rate was 93.18% in moderate cases, 100.00% in severe cases and 97.30% in critical cases||78.95 (IgM)|
|Liu Y et al., 2020||Retrospective||179||Patients, RNA positive (n = 90) and RNA negative (n = 89)||China||Commercial assay||Rapid immunoassay||The accuracy of the antibody testing increased over time (from 40% in the first week from onset of symptoms to 93.9% 2 weeks later)||85.6||91||95.1||82.7|
|Yong G et al., 2020||Retrospective||38||Patients||China||Commercial assay||Rapid assay GICA IgG IgM detection kit||The accuracy of the test 8 days after the onset of symptoms||50||92.1||NA||NA|
|Lin D et al., 2020||Retrospective||149||79 RNA-positive patients||China||Commercial assay||Darui Biotech, ELISA||The sensitivity of the test increased with time from onset of the disease||82.2||97.5||NA||NA|
|Lou B et al., 2020||Cross-sectional||380||80 RNA-positive patients. 300 healthy controls||China||Commercial assay||ELISA and lateral flow assay||The overall seroconversion rate was 98.8% at a median of 9 days from the onset of disease||98.8||94.3||NA||NA|
|Liu L et al., 2020||Cross-sectional||238||238 patients, 153 of them RNA-positive. 120 controls||China||Commercial assay||Lizhu, ELISA||Antibody detection should be used as a major viral diagnostic test for patients with symptoms for more than 10 days. The combination of ELISA and RT-PCR assays will greatly improve detection efficacy, even in the early stage of infection||81.5||NA||NA||NA|
|Bendavid E et al., 2020||Cross-sectional||3300||3300 subjects from the general population||China||Commercial||Premier Biotech, LFIA||The population prevalence of COVID-19 in Santa Clara, CA, ranged from 2.49% to 4.16%, 50- to 85-fold more than reported cases||80.3||99.5||NA||NA|
|Paradiso AV et al., 2020a||Prospective||191||191 symptomatic patients||China||Commercial||Rapid VivaDiag IgM /IgG immunoassay||The performance of the test at the onset of symptoms was low. Sensitivity was 66.7% 15 days later||30||89||NA||NA|
|Jia X et al., 2020||Retrospective||59||59 suspected patients, 24 of whom were RNA positive||China||Commercial assay||Diagreat, immunofluorescence assay||IgM and IgG may provide a quick, simple and accurate detection method for suspected COVID-19 patients||87.5||NA||NA||NA|
|Zhang J et al., 2020||Retrospective||736||228 suspected cases, 3 positive. 508 controls||China||Commercial assay||Shenzhen Yahuilong Biotechnology, chemiluminescence assay||Detection of specific antibodies in patients with fever can be a good complement to nucleic acid diagnosis for early diagnosis of suspected cases||100||97||75||100|
|Xiang J et al., 2020||Retrospective||189||154 patients, 35 controls||China||Commercial assays||Zhu Hai Liv Zon Diagnostics, ELISA and GICA assays||There is no difference between the sensitivity of ELISA and GICA assay; both are simple and fast, and the results can be used for clinical reference||87.3 (ELISA)|
|Hu Q et al., 2020||Prospective||993 samples from 221 subjects||221 hospitalized patients||China||Commercial assay||BioScience, chemiluminescence assay||IgG and IgM antibodies examined every 3 days revealed increasing antibody levels that peaked on day 19–21. SARS-CoV-2 IgG and IgM antibody testing should be combined with RT-PCR as an early diagnosis method||73.6 IgM|
(day 13–18 after the onset)
|Ma H et al., 2020||Cross-sectional||216 samples from 87 subjects||87 RNA-positive patients||China||In-house assay||Chemiluminescence||Measuring SARS-CoV-2 specific antibodies IgA, IgM and IgG in serum provides better serological testing with improved sensitivity and specificity||98.6 IgA|
|98.1 IgA 92.3 IgM 99.8 IgG||NA||NA|
|Qian C et al., 2020||Prospective, multicentric||2061 subjects from 10 hospitals||972 non-COVID patients, 586 controls, 503 RNA-positive patients||China||Commercial assay||Shenzhen YHLO Biotech, chemiluminescence assay||The assay showed a coefficient of variation of less than 5%. SARS-CoV-2 IgM and IgG showed clinical specificity of over 97% and 86.54% for suspected cases||85.8 IgM|
|National COVID testing Scientific Advisory Board, 2020||Cross-sectional||182||40 RNA-positive patients, 142 controls||UK||Commercial assays||Elisa and 9 commercial LFIA||The performance of current LFIA devices is inadequate for most individual patient applications. ELISA can be calibrated to be specific for detecting and quantifying SARSCoV-2 IgM and IgG and is highly sensitive for IgG from 10 days following symptom onset||85 (ELISA)|
55-70 (LFIA versus RT-PCR)
65–85 (LFIA versus ELISA)
|Burbelo PD et al., 2020||Cross-sectional||100||68 patients, 32 controls||USA||In-house assay||Luciferase 44 immunoprecipitation assay systems to the nucleocapsid (NP) and spike proteins (SP)||Antibody to the nucleocapsid protein of SARS-CoV-2 is more sensitive than 56 spike protein antibody for detecting early infection||100 (anti-NP)|
|Adams ER et al., 2020||Retrospective||834 samples||270 positive samples, 564 negative samples||UK||Commercial assay||Mologic, ELISA||The ELISA tested had good diagnostic performance||88||97||NA||NA|
|Meyer B et al., 2020||Retrospective||357 subjects||176 controls, 181 RNA-positive patients||Germany||Commercial assay||EUROIMMUN, ELISA||The assay displays an optima diagnostic accuracy using IgG, with no obvious gain from IgA serology||82||100||100||46|
|Norman M et al., 2020||Retrospective||81 subjects||81 subjects||USA||In-house assay||Single Molecular Array (SIMOA)||The SIMOA serological platform provides a powerful analytical tool||86||100||NA||NA|
|Tuaillon E et al., 2020||Prospective||58||38 RNA-positive patients and 20 controls||Euroimmun, GermanyIdVet, France||Commercial assay||Elisa by EUROIMMUN and IdVet and 5 rapid lateral flow tests||The second week of COVID-19 seems to be the best period for assessing the sensitivity of commercial serological assays||86.7 (ELISA)|
80–93.3 (rapid tests)
65–100 (rapid tests)
|Wajnberg A et al., 2020||Prospective||1343 subjects||1343 symptomatic subjects, of whom 624 were RNA-positive||Roche, USA||Commercial assay||Roche, chemiluminescence assay||The vast majority of confirmed COVID-19 patients seroconvert, potentially providing immunity to reinfection||82||NA||NA||NA|
|Wan Y et al., 2020||Retrospective||180||50 RNA-positive patients and 130 controls||China||Commercial assay||Four chemiluminescence assay systems||Systems for CoVID-2019 IgM/IgG antibody test may perform differently||26–92||78–99||NA||NA|
|Xiao T et al., 2020||Retrospective||56 subjects||56 RNA-positive patients (33 symptomatic and 23 asymptomatic)||China||Commercial assay||Chemiluminescence microparticle immunoassay||Asymptomatic carriers were found to have a lower initial viral load, undetectable IgM and moderate levels of IgG||90.9|
|Zhou Q et al., 2020||Retrospective||419 subjects||19 RNA-positive patients and 400 controls||China||Commercial assay||Chemiluminescence||Viral serological testing is an effective means of detecting SARS-CoV-2 infection||91.6||NA||NA||NA|
|Ozturk T et al., 2020||Cross-sectional||148 subjects||32 RNA-positive patients, 116 controls||USA||Commercial assay||GenScript, ELISA||There is a complex relationship between antibody levels, disease severity and time since symptom onset, so caution is needed in using serological assay to inform public policies||88.9||92.3||NA||NA|
|Rosado J et al., 2020||Retrospective||594||259 RNA-positive patients, 335 controls||France||In-house assay||Multiplex serological assay using a serological signature of IgG to four antigens||Serological signatures based on antibody responses to multiple antigens can provide more accurate and robust serological classification of individuals with previous SARS-CoV-2 infection||96.1||99.1||NA||NA|
AUC, area under the curve; ELISA, enzyme-linked immunosorbent assay; GICA, gold immunochromatographic assay; Ig; LFIA, lateral flow immunoassay; NA, not available; NPV, negative predictive value; PPV, positive predictive value; qPCR, quantitative PRC; POC, point of care; ROC, receiver operating characteristic; RT-PCR, reverse transcription-PCR; SARS-CoV-2, severe acute respiratory syndrome coronavirus 2.
REFERENCE LINK : https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7293848/
More information: Kathleen M. McAndrews et al, Heterogeneous antibodies against SARS-CoV-2 spike receptor binding domain and nucleocapsid with implications on COVID-19 immunity, JCI Insight (2020). DOI: 10.1172/jci.insight.142386