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Laboratory tests for the detection of SARS-CoV-2 infection: basic principles and examples

Labortests zum Nachweis einer SARS-CoV-2-Infektion: Grundprinzipien und Beispiele

Review Article Laboratory Diagnostics

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  • corresponding author Khaled R. Alkharsah - Department of Microbiology, College of Medicine, Imam Abdulrahman Bin Faisal University (IAU), Dammam, Saudi Arabia

GMS Ger Med Sci 2021;19:Doc06

doi: 10.3205/000293, urn:nbn:de:0183-0002936

Eingereicht: 29. Oktober 2020
Überarbeitet: 4. Februar 2021
Veröffentlicht: 27. Mai 2021

© 2021 Alkharsah.
Dieser Artikel ist ein Open-Access-Artikel und steht unter den Lizenzbedingungen der Creative Commons Attribution 4.0 License (Namensnennung). Lizenz-Angaben siehe http://creativecommons.org/licenses/by/4.0/.

Abstract

Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) has circulated throughout the world causing the worst pandemic since 1918. All efforts have been marshalled towards testing different treatment approaches, obtaining clinical and epidemiological information, developing suitable diagnostic tests, and developing new vaccines. New ribonucleic acid (RNA)-based and viral vector-based vaccines have been developed and licensed under emergency use in many countries; however, there is a huge demand for vaccines, and it will take some time before a sufficient number of people are vaccinated to stop the circulation of the virus. Therefore, the proper diagnosis and identification of infected individuals are crucial for the isolation and treatment of these patients and tracing of their contacts. Many diagnostic tests and diag-nostic kits have been developed in a relatively short time. This review summarizes the principles of the available laboratory assays that are in use for the detection of SARS-CoV-2 RNA, antigens, or antibodies.

Keywords: COVID-19, SARS-CoV-2, coronavirus, laboratory test, molecular assay, immunoassay

Zusammenfassung

Das Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2), das sich weltweit ausgebreitet hat, ist für eine der größten Pandemien seit 1918 verantwortlich. Seit Beginn der Pandemie laufen weltweit zahlreiche klinische und epidemiologische Studien, um mögliche effektive Behandlungsansätze zu testen und geeignete diagnostische Testverfahren zu entwickeln. Obwohl in einigen Ländern bereits einige RNA- und Vektorimpfstoffe gegen das Virus entwickelt und zugelassen wurden, ist die Nachfrage nach Impfstoffen in vielen Ländern immer noch groß. Aus diesem Grund und bis eine ausreichende Zahl von Menschen weltweit geimpft ist, bleibt eine schnelle Diagnose die entscheidende Maßnahme, um infizierte Personen zu identifizieren und das Infektionsgeschehen zurückzuverfolgen. In relativ kurzer Zeit wurden viele diagnostische Testverfahren und diagnostische Kits entwickelt, die den Nachweis von SARS-CoV-2-RNA, -Antigenen oder -Antikörpern ermöglichen. Dieser Review-Artikel gibt eine Übersicht über verfügbare diagnostische COVID-19-Tests, ihre Prinzipien und Einsatzmöglichkeiten.

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1 Introduction

The pandemic of coronavirus disease 2019 (COVID-19) continues to evolve during 2020 affecting life, health, and economy worldwide, making it the worst pandemic since the influenza pandemic in 1918. In addition to millions of infections and millions of deaths, many millions of people have lost their jobs [1]. Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), the causative agent of COVID-19, emerged in China at the end of 2019 causing a cluster of pneumonia cases and later spread worldwide [2], [3]. SARS-CoV-2 shares genetic homology with the bat coronaviruses from the genus betacoronavirus, family Coronaviridae, and the previously emerged human coronaviruses; severe acute respiratory syndrome Coronavirus (SARS-CoV) and the Middle East respiratory syndrome Coronavirus (MERS-CoV) [2], [3]. The new coronavirus was found to share 79% genetic similarity with the SARS-CoV and only 50% genetic homology with the MERS-CoV, hence its name of SARS-CoV-2 [4].

The early diagnosis of infected individuals and tracing of their contacts, to isolate them and protect susceptible individuals, are integral factors to controlling infection and containing the pandemic. COVID-19 signs and symptoms such as fever, dry cough, fatigue, and shortness of breath are common among other respiratory tract infections such as SARS, MERS, and influenza [5]. Moreover, patients with COVID-19 presented with diverse clinical symptoms based on the severity of their case, such as sore throat, diarrhea, vomiting, and abdominal pain [5]. Therefore, a clinical diagnosis despite being suggestive is not conclusive of COVID-19. Presymptomatic and asymptomatic transmission of SARS-CoV-2 are missed by clinical diagnosis and contribute to the aggravated spread of the virus and increasing the morbidity and mortality of the disease [6]. The surveillance program, therefore, is a priority to rapidly identify cases and allow contact tracing, isolate the infected individuals, and identify the regional spread of the virus.

Laboratory diagnostic tests can identify apparent and asymptomatic infections and facilitate surveillance programs by detecting the viral nucleic acid, the viral antigens, or the antibodies against the viral antigens. The laboratory diagnosis of SARS-CoV-2 infection can not only play a role in identifying new infections but also in understanding the spread of the disease, the level of its containment in a community, and the level of immunity in a population.

Since the emergence of SARS-CoV-2, unprecedented efforts have been made to develop different types of laboratory assays at exceptional speeds. A laboratory developed test (LDT) for the detection of the viral RNA was available very soon after the announcement of the viral genome sequence [7]. Later, multiple molecular and immunoassays were developed and approved for diagnostic detection of SARS-CoV-2 infection. This review describes the principle of laboratory tests currently available for the detection of COVID-19 infection and summarizes their utility and spectrum of use.

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2 General laboratory tests and their interpretation

Several blood parameters have been used to reflect the association with SARS-CoV-2 infection. In general, patients infected with SARS-CoV-2 have normal or more commonly decreased leukocyte counts with lymphocytopenia [8], [9], [10], [11], [12]. However, this was not found to be consistent throughout the course of infection. The leukocyte total and differential count vary between mild and severe infections. With progression to severe COVID-19 symptoms, patients tend to have increased leukocyte counts with lymphocytopenia [13], [14], [15]. Lymphocytopenia seems to be a hallmark of COVID-19 severity. Tan et al. suggested a model whereby the percentage of lymphocytes can be used as a predictor of disease prognosis [16]. A lymphocyte percentage higher than 20% would be associated with recovery while a lymphocyte percentage lower than 5% after 17 days of symptoms indicates that the patient is critically ill and needs ICU admission with a high possibility of death [16]. The mechanism by which lymphocytopenia occurs during COVID-19 has not yet been fully studied. It may be due to the direct infection and destruction of lymphocytes by the virus, resulting from their expression of the angiotensin-converting enzyme 2 (ACE2) receptor [17]. Lymphocytes may also be killed by apoptosis induced by several proinflammatory cytokines produced during the course of the disease such as tumor necrosis factor-alpha [18], [19]. Leukocytosis is also accompanied by increased neutrophil counts in severe COVID-19 cases compared to mild cases [15].

C-reactive protein (CRP) is an acute-phase protein expressed at high concentrations during acute inflammatory infection [20]. CRP has been reported to be high in all patients during the course of COVID-19 [10], [21]. However, the CRP levels were lower in children than in adult patients, which may correlate with the milder symptoms observed in children [22]. CRP levels proportionally correlate with the disease severity and diameter of the lung lesions, being highest in critically ill patients with large lung lesions and lower in milder groups with smaller lung lesions [23]. Serum amyloid A (SAA) is another acute-phase protein produced by the liver. It has also been found to increase in severe COVID-19 cases and predicts a poor prognosis [24], [25]. Increased CRP and SAA expression during acute inflammation is usually driven by inflammatory cytokines such as interleukin 6 (IL-6). IL-6 has repeatedly been reported to be high in critically ill COVID-19 patients. Therefore, several studies have suggested using IL-6 as a prognostic biomarker for the development of severe symptoms [26], [27], [28]. Moreover, IL-6 levels were found to be higher in deteriorating patient cases than in patients discharged after SARS-CoV-2 infection [29]. Therefore, it has been suggested to use an IL-6 inhibitor for the treatment of severe COVID-19 cases [30]. Tocilizumab, an anti-IL-6 receptor antibody, was found to produce a rapid and sustained improvement in patients with COVID-19 pneumonia and hyperinflammatory syndrome [31].

D-dimer is a product of fibrin breakdown during the degradation of vascular thrombi and is therefore a good indicator of hemostatic abnormalities and intravascular thrombosis [32]. Normal levels of D-dimer can rule out a suspected pulmonary embolism or deep vein thrombosis in patients with low risk [33]. High levels of D-dimer are indicative of venous thromboembolism but also of other conditions such as pregnancy, impaired renal function, and atrial fibrillation [33]. In severe COVID-19 cases, the risk of developing pulmonary embolism increases with disease progression after the first indication of high D-dimer levels [34]. D-dimer levels were also found to stay high or increase in the non-survivor group of COVID-19 patients compared to survivors [35]. D-dimer levels higher than a cutoff of 1,570 ng/mL were suggestive of asymptomatic deep vein thrombosis in patients with COVID-19 pneumonia [36]. Moreover, COVID-19 patients with D-dimer levels higher than 2,000 ng/mL on hospital admission had a higher incidence of death compared to patients with lower D-dimer levels [37].

Other blood biomarkers have also been found to be elevated in COVID-19 patients; however, these other blood biomarkers are not specific indicators of infection but are more useful in the prognosis of disease progression.

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3 Molecular testing

Suitable samples and proper sampling are crucial for the molecular detection of respiratory viruses. Different types of respiratory tract samples are used to diagnose SARS-CoV-2 infection. A higher rate of positive results is obtained from lower respiratory tract samples; therefore, sputum samples are the better samples when available [38]. If a sputum sample is not available, other upper respiratory tract samples such as nasopharyngeal swabs, oropharyngeal swabs, throat swabs, nasal swabs, or saliva can be used for the detection of SARS-CoV-2 RNA. Nasopharyngeal swabs seem to have a better yield than oropharyngeal and throat swabs detecting more than 80% of cases [38], [39]. Saliva samples are considered a non-invasive alternative for nasopharyngeal swabs, as the two samples show more than 97% agreement [40]. Thermal inactivation of samples prior to testing as a form of personal protection is reported to reduce the RNA copy number by approximately 50% and TRIzol inactivation by about 40%; therefore, it is not recommended [41], [42]. The commonly used nucleic acid extraction protocols and the available nucleic acid extraction kits include a lysis step which is expected to inactivate viruses. However, the efficiency of inactivation varies depending on the lysis buffers employed and the type of sample; therefore, the samples should be considered as potentially infectious [43].

3.1 Real-time reverse transcriptase polymerase chain reaction (rRT-PCR)

The real-time reverse transcriptase polymerase chain reaction (rRT-PCR) is, thus far, the gold standard test for the detection of SARS-CoV-2 RNA in clinical samples and plays an important role in the identification of cases [44]. Several protocols have been designed to target different fragments of the viral genome including genes encoding structural proteins, such as spike (S), transmembrane (M), envelope (E), and nucleocapsid (N), or species-specific genes encoding functional proteins such as the RNA-dependent RNA polymerase (RdRp), open reading frame 1a (ORF1a), and ORF1b (Figure 1 [Fig. 1]). The World Health Organization (WHO) recommends the laboratory confirmation of the COVID-19 case by nucleic acid amplification test (NAAT)-based assays in two different spatial categories. In areas with no known circulation of SARS-CoV-2, positive assay results from two different genomic targets specific for SARS-CoV-2 are required to confirm the COVID-19 case, or one positive result for a genomic target from the Betacoronavirus genus and sequencing of another genomic fragment for SARS-CoV-2 [45]. For areas with a wide spread of SARS-CoV-2, a positive result from one SARS-CoV-2 discriminatory genomic target is considered sufficient [45]. The American Food and Drug Administration (FDA) has approved the use of over 65 in vitro diagnostic kits for the detection of SARS-CoV-2 under Emergency Use Authorizations (EUA) (FDA-approved kits). These kits are authorized for diagnostic use during the existing circumstances and require further validation. Table 1 [Tab. 1] summarizes a selection of these kits.

While a positive rRT-PCR result for SARS-CoV-2 with clinical manifestation of the disease confirms a COVID-19 case, a negative rRT-PCR does not necessarily preclude infection in the presence of suggestive symptoms. Early studies from China showed that rRT-PCR detected a lower number of COVID-19 cases than chest computerized tomography (CT) [46], [47]. A similar experience was observed with MERS [48]. PCR sensitivity is reported to fade during the course of the infection, being more than 80% during the first 3 days of infection and reaching 45% after day 15; therefore, multiple testing is recommended [49], [50], [51]. The reasons for low PCR sensitivity could be either preanalytical or analytical. Preanalytical causes of low sensitivity range from the type of sample and improper sampling procedures to bad storage and transport [52]. Analytical reasons are mainly attributed to variation between diagnostic assays, which may have been inadequately validated due to the emergency release and approval of the assays [52].

Several studies have compared commercially available assays. A comparison between multiple commercially available kits (from Cepheid, DiaSorin, Hologic Panther, and Roche Cobas) and the protocol designed by the Centers for Disease Control and Prevention (CDC) for SARS-CoV-2 detection based on the N1 and N2 regions showed that all of the assays were 100% specific, and the Cepheid kit had 100% agreement with the CDC protocol [53]. The Abbott Real-Time SARS-CoV-2 assay was found to detect more positive cases than the modified CDC 2019-nCoV RT-PCR [54]. The Cobas SARS-CoV-2 Test detected 4.2% more positive cases than a laboratory-developed assay based on the CDC protocol targeting the N1 and N2 regions of the viral genome [55]. The Cobas 6800 SARS-CoV-2 Test showed 96.4% agreement with the Panther Fusion SARS-CoV-2 Assay [56]. The Panther Fusion SARS-CoV-2 Assay showed 98.3% overall agreement with a laboratory-developed assay detecting the E gene of the virus [57]. The Roche Cobas assay showed very good agreement with the Cepheid assay on samples with high cycle thresholds [58].

It is worth noting, however, that a positive PCR result does not necessarily indicate the presence of infectious virus particles. A study suggested that a cycle threshold (Ct) equal to or more than 24 after 8 days from symptoms onset might predict the presence of noninfectious virus particles [59]. Another study failed to isolate virus particles in culture from samples containing 100,000 virus particles per milliliter of sputum after 10 days from symptom onset, and therefore suggested the discharge of a patient with these criteria [60]. These studies provide a guide for infection control and help in reducing the hospitalization period.

3.2 Droplet digital polymerase chain reaction (ddPCR)

Droplet digital PCR (ddPCR) is quantitative PCR utilizing standard PCR components with two main differences: first, the PCR reaction is performed in thousands of individually fractionated reaction droplets and second, the reaction results is read at an endpoint, unlike the real-time PCR [61]. These two features provide the ddPCR with multiple advantages over the real-time PCR such as: the direct and reproducible quantification of the target without the need for a standard curve, the potential for greater target multiplexing, and due to dilution, negation of the effect of PCR inhibitors, allowing the amplification of target with extremely low starting quantities [61].

In a comparison between real-time PCR and ddPCR for the detection of SARS-CoV-2, the ddPCR detected SARS-CoV-2 in 26 samples that were negative by real-time PCR and had improved sensitivity, negative predictive value, negative likelihood ratio, and accuracy [62]. Moreover, the ddPCR can also be used for the evaluation of disease progression through the monitoring of viral load in patient samples [63]. Two droplet digital PCR assays have been approved by the FDA for the detection of SARS-CoV-2 (Table 1 [Tab. 1]).

3.3 Loop-mediated isothermal amplification (LAMP) assays

The loop-mediated isothermal amplification (LAMP) assay was developed in 2000 to overcome the shortcomings of other nucleic acid amplification methods such as the need for complicated instruments for target amplification and detection of the product, as for PCR; the use of expensive modified nucleotides, as for strand displacement amplification; or compromised specificity due to low amplification temperature, as in self-sustained sequence replication and sequence-based amplification [64]. The LAMP assay utilizes 4–6 primers and a DNA polymerase with strand displacement activity. The primers work in sequential order during the LAMP cycles to produce an amplicon that makes a loop structure at its end due to the included complementary sequence in the design of the primers. This loop structure is used as a template with multiple amplification starting points which leads to the accumulation of large amounts of the target that can be detected by simple colorimetric methods [64]. These features make the LAMP assay rapid, sensitive, target-specific, and instrument-independent [65]. However, the primer design in LAMP is more complicated than in PCR, multiplexing of the target is limited, and the final product is not suitable for downstream amplification [65].

In combination with reverse transcription, the LAMP assay can also be used to detect RNA targets [64]. Numerous studies have described the design of LAMP assays for the detection of SARS-CoV-2, which have proven to be highly sensitive and have a similar specificity to that of RT-PCR [66], [67]. The limit of detection of the LAMP assay has been reported as approximately 100 RNA copies per reaction, and the turnaround time was less than 30 minutes [68], [69]. However, there are only two FDA-approved kits for diagnostic detection of SARS-CoV-2 based on LAMP technology. The AQ-TOP COVID-19 Rapid Detection Kit is a LAMP-based assay targeting the ORF1ab of SARS-CoV-2. The assay measures the fluorescence generated by fluorescence resonance energy transfer probes, which are integrated into the amplified products and, therefore, the results are given in real-time. The assay can be used on different thermocycler instruments and delivers positive results in 15 minutes, while it takes 30 minutes for a negative result. Despite the short time to result, this assay cannot be considered as rapid because it does not include RNA extraction, which must be performed separately.

The ID NOW COVID-19 assay from Abbott is another FDA-approved test based on LAMP technology (see Rapid Point-of-Care Assays below).

3.4 Rapid Point-of-Care Assays (POCT)

The need for prompt identification of SARS-CoV-2-infected individuals is crucial to put patients on appropriate management regimens and to reduce the transmission risk. Two rapid diagnostic assays were approved by the FDA for diagnostic use under emergency authorization (Table 1 [Tab. 1]). The ID NOW COVID-19 assay from Abbott employs the isothermal nucleic acid amplification technique directed towards a unique region in the RdRp region of the genome [70]. The test results can be delivered within 5–13 minutes according to the manufacturer’s instructions; therefore, the assay is very convenient as a point-of-care (POC) assay. The assay has 100% specificity and negative agreement with other rRT-PCR assays [71], [72], [73]. However, the assay detected 33%–45% less positive cases than the Xpress Xpert assay depending on the type of sample [70]. It also detected a lower number of positive samples when compared to the Abbott Real-Time SARS-CoV-2 assay and the Roche Cobas SARS-CoV-2 assay, especially in samples with a low viral load [54], [72]. Its overall sensitivity was estimated to be approximately 71.7% [73].

The Accula SARS-CoV-2 Test is another FDA-approved rapid test that can yield results within 30 minutes [74]. The assay employs RT-PCR techniques targeting the nucleocapsid gene of the virus; however, it reads the result via the lateral flow technique [74]. Similar to the Abbott ID NOW COVID-19 assay, the Accula SARS-CoV-2 showed 100% negative agreement when compared to assays targeting the E gene, while it had only 68.0% positive agreement especially in samples with a low viral load [74].

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4 Immunological tests

As mentioned, the false-negative RT-PCR results for SARS-CoV-2, despite the presence of suggestive clinical disease manifestations, constitute a major challenge for the medical staff and increase the load on infection control officers and isolation rooms in hospitals. The humoral immune response against SARS-CoV-2 may help in confirming the diagnosis. Immunoglobulin M (IgM) and IgA are reported to appear within 5 days from symptom onset in 85.4% and 92.7% of patients, respectively [50], [51]. IgG antibodies are detected in 79% of the patients 14 days post-symptom onset [50], [51]. Combining both molecular and immunological tests was found to significantly increase the positive detection rate by more than 96% [50], [51], [75], [76].

Detection of SARS-CoV-2 antibodies has a number of advantages in addition to improving the total detection rate in COVID-19 patients. Antibody detection can help in estimating the actual disease prevalence in the population. Immunological assays detecting antibodies can be performed on a large scale, allowing for mass testing of the population. Furthermore, the detection of antibodies in COVID-19 patients may help in predicting disease prognosis. Patients with severe symptoms have been reported to have higher IgM antibody levels for a longer period of time [75], [77].

Immunological tests can detect either virus antigens or the antibodies against these antigens produced as an immune response to infection.

4.1 Antigen (Ag)-based immunoassays

This type of assay looks for virus proteins in respiratory samples from patients (such as nasopharyngeal swabs, oropharyngeal swabs, or nasal swabs) using an antibody specific to a certain target and different methods to detect the antigen-antibody (Ag-Ab) complexes, such as the immunofluorescence or lateral flow assays.

Multiple Ag-based immunoassays received the FDA approval under the emergency authorization such as the Sofia 2 SARS Antigen FIA, BinaxNOW COVID-19 Ag Card, and BD Veritor System for Rapid Detection of SARS-CoV-2 (Table 2 [Tab. 2]; complete list: [78]). These assays are designed as POC tests providing results within 15 minutes. Many of these assays have not been validated by enough studies yet; however, they will have an important utility in the rapid diagnosis of patients admitted to the emergency department in hospitals. A recent comparison of the BD Veritor SARS-CoV-2 and the Sofia 2 SARS Antigen FIA tests, relative to the PCR results, found that these assays had 98.1% overall percent agreement with each other. The Veritor test showed 81.8% to 87.5% agreement with the PCR results, improving at increasing days from symptom onset [79]. Another study estimated the analytical sensitivity of the Abbott BinaxNOW COVID-19 Ag Card to be 40,000 to 80,000 copies/swab [80].

4.2 Antibody (Ab)-based immunoassays

A humoral immune response including IgM, IgG, and IgA antibodies can be measured by serological tests such as enzyme immunoassays (EIA). Numerous forms of EIA are available on the market. The FDA approved three types of such assays for testing of patients with COVID-19: the rapid immunoassays, Chemiluminescence immunoassays (CLIA), and the enzyme-linked immunosorbent assays (ELISA).

4.2.1 Rapid immunoassays

Some rapid immunoassays employ lateral flow technology to detect the presence of IgM, IgG, or most commonly IgG and IgM antibodies simultaneously, against SARS-CoV-2. These assays are based on the liquid chromatography principle whereby the antibodies in patient samples migrate in flow with the buffering reagents on a nitrocellulose membrane. Detection components are immobilized on certain pre-marked locations on the nitrocellulose membrane. The rapid immunoassays can deliver results within 15 minutes and are simple, can be performed without the need for expensive equipment, and show good performance in comparison to conventional immunoassays [81]. The multiple advantages of rapid immunoassays encourage their use in the diagnosis of SARS-CoV-2 in clinical laboratories, especially in countries with limited resources [82]. Many commercially available rapid immunoassay kits received FDA approval for the detection of SARS-CoV-2 antibodies under emergency authorization (Table 2 [Tab. 2]). The rapid immunoassays are usually highly specific; however, they only have acceptable sensitivity [83], [84], [85], [86], [87].

4.2.2 Enzyme immunoassays (EIA)

The most commonly used EIAs in clinical laboratories are the enzyme-linked immunosorbent assay (ELISA) and the Chemiluminescence immunoassay (CLIA). ELISA and CLIA are sensitive assays and can be used to test a large number of samples in one run, which makes them suitable for laboratories with high throughput requirements. They are also available in fully automated system formats and in manual formats. Some CLIAs have an advantage over ELISA in terms of turnaround time and analytical sensitivity. These assays can be designed to detect total SARS-CoV-2 antibodies or antibodies against one of its protein components such as the S protein or the nucleocapsid protein. It has been reported that the EIAs that detect anti-nucleocapsid protein antibodies provide better sensitivity than those that detect anti-spike protein antibodies [88]. However, since the nucleocapsid protein is one of the conserved proteins among coronaviruses, further studies are required to investigate whether this preferential sensitivity from anti-nucleocapsid protein antibodies is not due to cross-reactivity with other coronaviruses.

The enzyme immunoassays, in general, have shown very good performance in the detection of antibodies after day 10 of infection, with compromised sensitivity during days 5–9 of the infection [89], [90]. This is not due to a technical problem in the established assays but rather due to the immunological window required until antibodies are produced, which differs from patient to patient.

4.3 Virus neutralization assay (VNA)

While other immunoassays detect the total antibody spectrum against a virus or certain of its components, the virus neutralization assay (VNA) distinguishes the fraction of antibodies that interfere with the binding of the virus to its cellular receptor, inhibits cell entry, and neutralizes its infectivity; hence called neutralizing antibodies. In case of SARS-CoV-2, the neutralizing antibodies are directed against the receptor-binding domain (RBD) in the spike protein of the virus. The VNA is very helpful during the emergence of a new virus when no commercial assays are available to detect the presence of virus-specific antibodies. It is also helpful in determining whether a convalescent plasma contains protective neutralizing antibodies against the virus [91].

Conventional VNA is performed by mixing the whole virus in cell culture supernatant with the serum of a patient and measuring the virus infectivity to target cells compared to a control without patient serum. The test results are then measured by the level of damage the virus causes to the cell (cytopathic effect) [92]. However, this form of the assay should be performed in a biosafety level-3 (BLS-3) laboratory. To avoid handling infectious virus particles, a modified VNA based on a pseudovirus bearing the outer membrane of the infectious virus to be studied and all other components from other non-human pathogen virus or a mutated virus are used [93]. The readout in this case is a fluorescence such as green fluorescent protein or luciferase measurements in the infected cells. This modified assay can then be performed in a BSL-2 laboratory. The assay can also quantitatively determine the titer of antibodies by diluting the patient samples and reporting the lowest dilution of sample inhibiting virus or pseudovirus entry.

Despite the diagnostic and research utility of the VNA, it requires cell culture facility and specially trained personnel and therefore would not be convenient for commercial production.

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5 Conclusions

Prompt diagnosis of COVID-19 cases is crucial for isolation, treatment, and contact tracing. Molecular diagnostic assays constitute by far the first line of defense for the identification of infected individuals. Immunological assays are mostly negative at the beginning of symptoms and cannot therefore play a significant role in the diagnosis of infected patients, but rather in defining the immunological status of a population at risk, such as health care workers, or the general population, particularly with the reports of frequent asymptomatic infection.

We additionally urgently need to produce more sensitive assays to reduce false-negative results and their deleterious effects on the spread of the virus in the human population.

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Notes

Competing interests

The author declares that he has no competing interests.

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References

1.
Cutler D. How Will COVID-19 Affect the Health Care Economy? JAMA. 2020 Jun;323(22):2237-8. DOI: 10.1001/jama.2020.7308 Externer Link
2.
Zhu N, Zhang D, Wang W, Li X, Yang B, Song J, Zhao X, Huang B, Shi W, Lu R, Niu P, Zhan F, Ma X, Wang D, Xu W, Wu G, Gao GF, Tan W; China Novel Coronavirus Investigating and Research Team. A Novel Coronavirus from Patients with Pneumonia in China, 2019. N Engl J Med. 2020 Feb;382(8):727-33. DOI: 10.1056/NEJMoa2001017 Externer Link
3.
Wu F, Zhao S, Yu B, Chen YM, Wang W, Song ZG, Hu Y, Tao ZW, Tian JH, Pei YY, Yuan ML, Zhang YL, Dai FH, Liu Y, Wang QM, Zheng JJ, Xu L, Holmes EC, Zhang YZ. A new coronavirus associated with human respiratory disease in China. Nature. 2020 Mar;579(7798):265-9. DOI: 10.1038/s41586-020-2008-3 Externer Link
4.
Lu R, Zhao X, Li J, Niu P, Yang B, Wu H, Wang W, Song H, Huang B, Zhu N, Bi Y, Ma X, Zhan F, Wang L, Hu T, Zhou H, Hu Z, Zhou W, Zhao L, Chen J, Meng Y, Wang J, Lin Y, Yuan J, Xie Z, Ma J, Liu WJ, Wang D, Xu W, Holmes EC, Gao GF, Wu G, Chen W, Shi W, Tan W. Genomic characterisation and epidemiology of 2019 novel coronavirus: implications for virus origins and receptor binding. Lancet. 2020 Feb 22;395(10224):565-74. DOI: 10.1016/S0140-6736(20)30251-8 Externer Link
5.
Baj J, Karakuła-Juchnowicz H, Teresiński G, Buszewicz G, Ciesielka M, Sitarz E, Forma A, Karakuła K, Flieger W, Portincasa P, Maciejewski R. COVID-19: Specific and Non-Specific Clinical Manifestations and Symptoms: The Current State of Knowledge. J Clin Med. 2020 Jun;9(6):1753. DOI: 10.3390/jcm9061753 Externer Link
6.
Arons MM, Hatfield KM, Reddy SC, Kimball A, James A, Jacobs JR, Taylor J, Spicer K, Bardossy AC, Oakley LP, Tanwar S, Dyal JW, Harney J, Chisty Z, Bell JM, Methner M, Paul P, Carlson CM, McLaughlin HP, Thornburg N, Tong S, Tamin A, Tao Y, Uehara A, Harcourt J, Clark S, Brostrom-Smith C, Page LC, Kay M, Lewis J, Montgomery P, Stone ND, Clark TA, Honein MA, Duchin JS, Jernigan JA; Public Health – Seattle and King County and CDC COVID-19 Investigation Team. Presymptomatic SARS-CoV-2 Infections and Transmission in a Skilled Nursing Facility. N Engl J Med. 2020 May;382(22):2081-90. DOI: 10.1056/NEJMoa2008457 Externer Link
7.
Corman VM, Landt O, Kaiser M, Molenkamp R, Meijer A, Chu DK, Bleicker T, Brünink S, Schneider J, Schmidt ML, Mulders DG, Haagmans BL, van der Veer B, van den Brink S, Wijsman L, Goderski G, Romette JL, Ellis J, Zambon M, Peiris M, Goossens H, Reusken C, Koopmans MP, Drosten C. Detection of 2019 novel coronavirus (2019-nCoV) by real-time RT-PCR. Euro Surveill. 2020 Jan;25(3):2000045. DOI: 10.2807/1560-7917.ES.2020.25.3.2000045 Externer Link
8.
Singhal T. A Review of Coronavirus Disease-2019 (COVID-19). Indian J Pediatr. 2020 Apr;87(4):281-6. DOI: 10.1007/s12098-020-03263-6 Externer Link
9.
Zhang G, Hu C, Luo L, Fang F, Chen Y, Li J, Peng Z, Pan H. Clinical features and short-term outcomes of 221 patients with COVID-19 in Wuhan, China. J Clin Virol. 2020 Jun;127:104364. DOI: 10.1016/j.jcv.2020.104364 Externer Link
10.
Ferrari D, Motta A, Strollo M, Banfi G, Locatelli M. Routine blood tests as a potential diagnostic tool for COVID-19. Clin Chem Lab Med. 2020 Jun;58(7):1095-9. DOI: 10.1515/cclm-2020-0398 Externer Link
11.
Mardani R, Ahmadi Vasmehjani A, Zali F, Gholami A, Mousavi Nasab SD, Kaghazian H, Kaviani M, Ahmadi N. Laboratory Parameters in Detection of COVID-19 Patients with Positive RT-PCR; a Diagnostic Accuracy Study. Arch Acad Emerg Med. 2020 Apr 4;8(1):e43.
12.
Lei P, Fan B, Mao J, Wang P. Multiple parameters required for diagnosis of COVID-19 in clinical practice. J Infect. 2020 Jun;80(6):e27-8. DOI: 10.1016/j.jinf.2020.03.016 Externer Link
13.
Henry BM, de Oliveira MHS, Benoit S, Plebani M, Lippi G. Hematologic, biochemical and immune biomarker abnormalities associated with severe illness and mortality in coronavirus disease 2019 (COVID-19): a meta-analysis. Clin Chem Lab Med. 2020 Jun;58(7):1021-8. DOI: 10.1515/cclm-2020-0369 Externer Link
14.
Velavan TP, Meyer CG. Mild versus severe COVID-19: Laboratory markers. Int J Infect Dis. 2020 Jun;95:304-7. DOI: 10.1016/j.ijid.2020.04.061 Externer Link
15.
Zeng F, Li L, Zeng J, Deng Y, Huang H, Chen B, Deng G. Can we predict the severity of coronavirus disease 2019 with a routine blood test? Pol Arch Intern Med. 2020 May;130(5):400-6. DOI: 10.20452/pamw.15331 Externer Link
16.
Tan L, Wang Q, Zhang D, Ding J, Huang Q, Tang YQ, Wang Q, Miao H. Lymphopenia predicts disease severity of COVID-19: a descriptive and predictive study. Signal Transduct Target Ther. 2020 Mar;5(1):33. DOI: 10.1038/s41392-020-0148-4 Externer Link
17.
Xu H, Zhong L, Deng J, Peng J, Dan H, Zeng X, Li T, Chen Q. High expression of ACE2 receptor of 2019-nCoV on the epithelial cells of oral mucosa. Int J Oral Sci. 2020 Feb;12(1):8. DOI: 10.1038/s41368-020-0074-x Externer Link
18.
Gupta S. Tumor necrosis factor-alpha-induced apoptosis in T cells from aged humans: a role of TNFR-I and downstream signaling molecules. Exp Gerontol. 2002 Jan-Mar;37(2-3):293-9. DOI: 10.1016/s0531-5565(01)00195-4 Externer Link
19.
Pedersen SF, Ho YC. SARS-CoV-2: a storm is raging. J Clin Invest. 2020 May;130(5):2202-5. DOI: 10.1172/JCI137647 Externer Link
20.
Kushner I. The phenomenon of the acute phase response. Ann N Y Acad Sci. 1982;389:39-48. DOI: 10.1111/j.1749-6632.1982.tb22124.x Externer Link
21.
Aggarwal S, Garcia-Telles N, Aggarwal G, Lavie C, Lippi G, Henry BM. Clinical features, laboratory characteristics, and outcomes of patients hospitalized with coronavirus disease 2019 (COVID-19): Early report from the United States. Diagnosis (Berl). 2020 May;7(2):91-6. DOI: 10.1515/dx-2020-0046 Externer Link
22.
Du W, Yu J, Wang H, Zhang X, Zhang S, Li Q, Zhang Z. Clinical characteristics of COVID-19 in children compared with adults in Shandong Province, China. Infection. 2020 Jun;48(3):445-52. DOI: 10.1007/s15010-020-01427-2 Externer Link
23.
Wang L. C-reactive protein levels in the early stage of COVID-19. Med Mal Infect. 2020 Jun;50(4):332-4. DOI: 10.1016/j.medmal.2020.03.007 Externer Link
24.
Li H, Xiang X, Ren H, Xu L, Zhao L, Chen X, Long H, Wang Q, Wu Q. Serum Amyloid A is a biomarker of severe Coronavirus Disease and poor prognosis. J Infect. 2020 Jun;80(6):646-55. DOI: 10.1016/j.jinf.2020.03.035 Externer Link
25.
Ji M, Yuan L, Shen W, Lv J, Li Y, Li M, Lu X, Hu L, Dong W. Characteristics of disease progress in patients with coronavirus disease 2019 in Wuhan, China. Epidemiol Infect. 2020 May;148:e94. DOI: 10.1017/S0950268820000977 Externer Link
26.
Liu F, Li L, Xu M, Wu J, Luo D, Zhu Y, Li B, Song X, Zhou X. Prognostic value of interleukin-6, C-reactive protein, and procalcitonin in patients with COVID-19. J Clin Virol. 2020 Jun;127:104370. DOI: 10.1016/j.jcv.2020.104370 Externer Link
27.
Aziz M, Fatima R, Assaly R. Elevated interleukin-6 and severe COVID-19: A meta-analysis. J Med Virol. 2020 Nov;92(11):2283-5. DOI: 10.1002/jmv.25948 Externer Link
28.
Ulhaq ZS, Soraya GV. Interleukin-6 as a potential biomarker of COVID-19 progression. Med Mal Infect. 2020 Jun;50(4):382-3. DOI: 10.1016/j.medmal.2020.04.002 Externer Link
29.
Zhang J, Yu M, Tong S, Liu LY, Tang LV. Predictive factors for disease progression in hospitalized patients with coronavirus disease 2019 in Wuhan, China. J Clin Virol. 2020 Jun;127:104392. DOI: 10.1016/j.jcv.2020.104392 Externer Link
30.
Zhang Y, Zhong Y, Pan L, Dong J. Treat 2019 novel coronavirus (COVID-19) with IL-6 inhibitor: Are we already that far? Drug Discov Ther. 2020;14(2):100-2. DOI: 10.5582/ddt.2020.03006 Externer Link
31.
Toniati P, Piva S, Cattalini M, Garrafa E, Regola F, Castelli F, Franceschini F, Airò P, Bazzani C, Beindorf EA, Berlendis M, Bezzi M, Bossini N, Castellano M, Cattaneo S, Cavazzana I, Contessi GB, Crippa M, Delbarba A, De Peri E, Faletti A, Filippini M, Filippini M, Frassi M, Gaggiotti M, Gorla R, Lanspa M, Lorenzotti S, Marino R, Maroldi R, Metra M, Matteelli A, Modina D, Moioli G, Montani G, Muiesan ML, Odolini S, Peli E, Pesenti S, Pezzoli MC, Pirola I, Pozzi A, Proto A, Rasulo FA, Renisi G, Ricci C, Rizzoni D, Romanelli G, Rossi M, Salvetti M, Scolari F, Signorini L, Taglietti M, Tomasoni G, Tomasoni LR, Turla F, Valsecchi A, Zani D, Zuccalà F, Zunica F, Focà E, Andreoli L, Latronico N. Tocilizumab for the treatment of severe COVID-19 pneumonia with hyperinflammatory syndrome and acute respiratory failure: A single center study of 100 patients in Brescia, Italy. Autoimmun Rev. 2020 Jul;19(7):102568. DOI: 10.1016/j.autrev.2020.102568 Externer Link
32.
Johnson ED, Schell JC, Rodgers GM. The D-dimer assay. Am J Hematol. 2019 Jul;94(7):833-9. DOI: 10.1002/ajh.25482 Externer Link
33.
Wilbur J, Shian B. Diagnosis of deep venous thrombosis and pulmonary embolism. Am Fam Physician. 2012 Nov;86(10):913-9.
34.
Garcia-Olivé I, Sintes H, Radua J, Abad Capa J, Rosell A. D-dimer in patients infected with COVID-19 and suspected pulmonary embolism. Respir Med. 2020 Aug;169:106023. DOI: 10.1016/j.rmed.2020.106023 Externer Link
35.
Chen R, Sang L, Jiang M, Yang Z, Jia N, Fu W, Xie J, Guan W, Liang W, Ni Z, Hu Y, Liu L, Shan H, Lei C, Peng Y, Wei L, Liu Y, Hu Y, Peng P, Wang J, Liu J, Chen Z, Li G, Zheng Z, Qiu S, Luo J, Ye C, Zhu S, Zheng J, Zhang N, Li Y, He J, Li J, Li S, Zhong N; Medical Treatment Expert Group for COVID-19. Longitudinal hematologic and immunologic variations associated with the progression of COVID-19 patients in China. J Allergy Clin Immunol. 2020 Jul;146(1):89-100. DOI: 10.1016/j.jaci.2020.05.003 Externer Link
36.
Demelo-Rodríguez P, Cervilla-Muñoz E, Ordieres-Ortega L, Parra-Virto A, Toledano-Macías M, Toledo-Samaniego N, García-García A, García-Fernández-Bravo I, Ji Z, de-Miguel-Diez J, Álvarez-Sala-Walther LA, Del-Toro-Cervera J, Galeano-Valle F. Incidence of asymptomatic deep vein thrombosis in patients with COVID-19 pneumonia and elevated D-dimer levels. Thromb Res. 2020 Aug;192:23-6. DOI: 10.1016/j.thromres.2020.05.018 Externer Link
37.
Zhang L, Yan X, Fan Q, Liu H, Liu X, Liu Z, Zhang Z. D-dimer levels on admission to predict in-hospital mortality in patients with Covid-19. J Thromb Haemost. 2020 Jun;18(6):1324-9. DOI: 10.1111/jth.14859 Externer Link
38.
Lin C, Xiang J, Yan M, Li H, Huang S, Shen C. Comparison of throat swabs and sputum specimens for viral nucleic acid detection in 52 cases of novel coronavirus (SARS-Cov-2)-infected pneumonia (COVID-19). Clin Chem Lab Med. 2020 Jun;58(7):1089-94. DOI: 10.1515/cclm-2020-0187 Externer Link
39.
Wang X, Tan L, Wang X, Liu W, Lu Y, Cheng L, Sun Z. Comparison of nasopharyngeal and oropharyngeal swabs for SARS-CoV-2 detection in 353 patients received tests with both specimens simultaneously. Int J Infect Dis. 2020 May;94:107-9. DOI: 10.1016/j.ijid.2020.04.023 Externer Link
40.
Pasomsub E, Watcharananan SP, Boonyawat K, Janchompoo P, Wongtabtim G, Suksuwan W, Sungkanuparph S, Phuphuakrat A. Saliva sample as a non-invasive specimen for the diagnosis of coronavirus disease 2019: a cross-sectional study. Clin Microbiol Infect. 2021 Feb;27(2):285.e1-4. DOI: 10.1016/j.cmi.2020.05.001 Externer Link
41.
Chen H, Wu R, Xing Y, Du Q, Xue Z, Xi Y, Yang Y, Deng Y, Han Y, Li K, Luan Y, Zhang Y, Wei X, Yu T, Li H, Zhu L, Su S, Lian H, Lu L, Tan C, Zheng H, Chen B, Yu P, Guo Y, Ma C. Influence of Different Inactivation Methods on Severe Acute Respiratory Syndrome Coronavirus 2 RNA Copy Number. J Clin Microbiol. 2020 Jul;58(8). DOI: 10.1128/JCM.00958-20 Externer Link
42.
Pan Y, Long L, Zhang D, Yuan T, Cui S, Yang P, Wang Q, Ren S. Potential False-Negative Nucleic Acid Testing Results for Severe Acute Respiratory Syndrome Coronavirus 2 from Thermal Inactivation of Samples with Low Viral Loads. Clin Chem. 2020 Jun;66(6):794-801. DOI: 10.1093/clinchem/hvaa091 Externer Link
43.
Pastorino B, Touret F, Gilles M, Luciani L, de Lamballerie X, Charrel RN. Evaluation of Chemical Protocols for Inactivating SARS-CoV-2 Infectious Samples. Viruses. 2020 Jun;12(6). DOI: 10.3390/v12060624 Externer Link
44.
Liu R, Han H, Liu F, Lv Z, Wu K, Liu Y, Feng Y, Zhu C. Positive rate of RT-PCR detection of SARS-CoV-2 infection in 4880 cases from one hospital in Wuhan, China, from Jan to Feb 2020. Clin Chim Acta. 2020 Jun;505:172-5. DOI: 10.1016/j.cca.2020.03.009 Externer Link
45.
WHO. Laboratory testing for 2019 novel coronavirus (2019-nCoV) in suspected human cases. 2020. Available from: https://www.who.int/publications/i/item/laboratory-testing-for-2019-novel-coronavirus-in-suspected-human-cases-20200117 Externer Link
46.
Ai T, Yang Z, Hou H, Zhan C, Chen C, Lv W, Tao Q, Sun Z, Xia L. Correlation of Chest CT and RT-PCR Testing for Coronavirus Disease 2019 (COVID-19) in China: A Report of 1014 Cases. Radiology. 2020 Aug;296(2):E32-40. DOI: 10.1148/radiol.2020200642 Externer Link
47.
Fang Y, Zhang H, Xie J, Lin M, Ying L, Pang P, Ji W. Sensitivity of Chest CT for COVID-19: Comparison to RT-PCR. Radiology. 2020 Aug;296(2):E115-7. DOI: 10.1148/radiol.2020200432 Externer Link
48.
Al-Tawfiq JA, Memish ZA. Diagnosis of SARS-CoV-2 infection based on CT scan vs RT-PCR: reflecting on experience from MERS-CoV. J Hosp Infect. 2020 Jun;105(2):154-5. DOI: 10.1016/j.jhin.2020.03.001 Externer Link
49.
Long C, Xu H, Shen Q, Zhang X, Fan B, Wang C, Zeng B, Li Z, Li X, Li H. Diagnosis of the Coronavirus disease (COVID-19): rRT-PCR or CT? Eur J Radiol. 2020 May;126:108961. DOI: 10.1016/j.ejrad.2020.108961 Externer Link
50.
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, Fu Y, Ge S, Liu L, Zhang J, Xia N, Zhang Z. Antibody Responses to SARS-CoV-2 in Patients With Novel Coronavirus Disease 2019. Clin Infect Dis. 2020 Nov;71(16):2027-34. DOI: 10.1093/cid/ciaa344 Externer Link
51.
Guo L, Ren L, Yang S, Xiao M, Chang, Yang F, Dela Cruz CS, Wang Y, Wu C, Xiao Y, Zhang L, Han L, Dang S, Xu Y, Yang QW, Xu SY, Zhu HD, Xu YC, Jin Q, Sharma L, Wang L, Wang J. Profiling Early Humoral Response to Diagnose Novel Coronavirus Disease (COVID-19). Clin Infect Dis. 2020 Jul;71(15):778-85. DOI: 10.1093/cid/ciaa310 Externer Link
52.
Lippi G, Simundic AM, Plebani M. Potential preanalytical and analytical vulnerabilities in the laboratory diagnosis of coronavirus disease 2019 (COVID-19). Clin Chem Lab Med. 2020 Jun;58(7):1070-6. DOI: 10.1515/cclm-2020-0285 Externer Link
53.
Lieberman JA, Pepper G, Naccache SN, Huang ML, Jerome KR, Greninger AL. Comparison of Commercially Available and Laboratory-Developed Assays for In Vitro Detection of SARS-CoV-2 in Clinical Laboratories. J Clin Microbiol. 2020 Jul;58(8):e00821-20. DOI: 10.1128/JCM.00821-20 Externer Link
54.
Moore NM, Li H, Schejbal D, Lindsley J, Hayden MK. Comparison of Two Commercial Molecular Tests and a Laboratory-Developed Modification of the CDC 2019-nCoV Reverse Transcriptase PCR Assay for the Detection of SARS-CoV-2. J Clin Microbiol. 2020 Jul;58(8):e00938-20. DOI: 10.1128/JCM.00938-20 Externer Link
55.
Pujadas E, Ibeh N, Hernandez MM, Waluszko A, Sidorenko T, Flores V, Shiffrin B, Chiu N, Young-Francois A, Nowak MD, Paniz-Mondolfi AE, Sordillo EM, Cordon-Cardo C, Houldsworth J, Gitman MR. Comparison of SARS-CoV-2 detection from nasopharyngeal swab samples by the Roche cobas 6800 SARS-CoV-2 test and a laboratory-developed real-time RT-PCR test. J Med Virol. 2020 Sep;92(9):1695-8. DOI: 10.1002/jmv.25988 Externer Link
56.
Craney AR, Velu PD, Satlin MJ, Fauntleroy KA, Callan K, Robertson A, La Spina M, Lei B, Chen A, Alston T, Rozman A, Loda M, Rennert H, Cushing M, Westblade LF. Comparison of Two High-Throughput Reverse Transcription-PCR Systems for the Detection of Severe Acute Respiratory Syndrome Coronavirus 2. J Clin Microbiol. 2020 Jul;58(8):e00890–20. DOI: 10.1128/JCM.00890-20 Externer Link
57.
Hogan CA, Sahoo MK, Huang C, Garamani N, Stevens B, Zehnder J, Pinsky BA. Comparison of the Panther Fusion and a laboratory-developed test targeting the envelope gene for detection of SARS-CoV-2. J Clin Virol. 2020 Jun;127:104383. DOI: 10.1016/j.jcv.2020.104383 Externer Link
58.
Broder K, Babiker A, Myers C, White T, Jones H, Cardella J, Burd EM, Hill CE, Kraft CS. Test Agreement between Roche Cobas 6800 and Cepheid GeneXpert Xpress SARS-CoV-2 Assays at High Cycle Threshold Ranges. J Clin Microbiol. 2020 Jul 23;58(8):e01187-20. DOI: 10.1128/JCM.01187-20 Externer Link
59.
Bullard J, Dust K, Funk D, Strong JE, Alexander D, Garnett L, Boodman C, Bello A, Hedley A, Schiffman Z, Doan K, Bastien N, Li Y, Van Caeseele PG, Poliquin G. Predicting Infectious Severe Acute Respiratory Syndrome Coronavirus 2 From Diagnostic Samples. Clin Infect Dis. 2020 Dec 17;71(10):2663-6. DOI: 10.1093/cid/ciaa638 Externer Link
60.
Wölfel R, Corman VM, Guggemos W, Seilmaier M, Zange S, Müller MA, Niemeyer D, Jones TC, Vollmar P, Rothe C, Hoelscher M, Bleicker T, Brünink S, Schneider J, Ehmann R, Zwirglmaier K, Drosten C, Wendtner C. Virological assessment of hospitalized patients with COVID-2019. Nature. 2020 May;581(7809):465-9. DOI: 10.1038/s41586-020-2196-x Externer Link
61.
Taylor SC, Laperriere G, Germain H. Droplet Digital PCR versus qPCR for gene expression analysis with low abundant targets: from variable nonsense to publication quality data. Sci Rep. 2017 May;7(1):2409. DOI: 10.1038/s41598-017-02217-x Externer Link
62.
Suo T, Liu X, Feng J, Guo M, Hu W, Guo D, Ullah H, Yang Y, Zhang Q, Wang X, Sajid M, Huang Z, Deng L, Chen T, Liu F, Xu K, Liu Y, Zhang Q, Liu Y, Xiong Y, Chen G, Lan K, Chen Y. ddPCR: a more accurate tool for SARS-CoV-2 detection in low viral load specimens. Emerg Microbes Infect. 2020 Dec;9(1):1259-68. DOI: 10.1080/22221751.2020.1772678 Externer Link
63.
Yu F, Yan L, Wang N, Yang S, Wang L, Tang Y, Gao G, Wang S, Ma C, Xie R, Wang F, Tan C, Zhu L, Guo Y, Zhang F. Quantitative Detection and Viral Load Analysis of SARS-CoV-2 in Infected Patients. Clin Infect Dis. 2020 Jul;71(15):793-8. DOI: 10.1093/cid/ciaa345 Externer Link
64.
Notomi T, Okayama H, Masubuchi H, Yonekawa T, Watanabe K, Amino N, Hase T. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res. 2000 Jun;28(12):E63. DOI: 10.1093/nar/28.12.e63 Externer Link
65.
Kashir J, Yaqinuddin A. Loop mediated isothermal amplification (LAMP) assays as a rapid diagnostic for COVID-19. Med Hypotheses. 2020 Aug;141:109786. DOI: 10.1016/j.mehy.2020.109786 Externer Link
66.
Park GS, Ku K, Baek SH, Kim SJ, Kim SI, Kim BT, Maeng JS. Development of Reverse Transcription Loop-Mediated Isothermal Amplification Assays Targeting Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2). J Mol Diagn. 2020 Jun;22(6):729-35. DOI: 10.1016/j.jmoldx.2020.03.006 Externer Link
67.
Baek YH, Um J, Antigua KJC, Park JH, Kim Y, Oh S, Kim YI, Choi WS, Kim SG, Jeong JH, Chin BS, Nicolas HDG, Ahn JY, Shin KS, Choi YK, Park JS, Song MS. Development of a reverse transcription-loop-mediated isothermal amplification as a rapid early-detection method for novel SARS-CoV-2. Emerg Microbes Infect. 2020 Dec;9(1):998-1007. DOI: 10.1080/22221751.2020.1756698 Externer Link
68.
Lu R, Wu X, Wan Z, Li Y, Jin X, Zhang C. A Novel Reverse Transcription Loop-Mediated Isothermal Amplification Method for Rapid Detection of SARS-CoV-2. Int J Mol Sci. 2020 Apr;21(8). DOI: 10.3390/ijms21082826 Externer Link
69.
Yan C, Cui J, Huang L, Du B, Chen L, Xue G, Li S, Zhang W, Zhao L, Sun Y, Yao H, Li N, Zhao H, Feng Y, Liu S, Zhang Q, Liu D, Yuan J. Rapid and visual detection of 2019 novel coronavirus (SARS-CoV-2) by a reverse transcription loop-mediated isothermal amplification assay. Clin Microbiol Infect. 2020 Jun;26(6):773-9. DOI: 10.1016/j.cmi.2020.04.001 Externer Link
70.
Basu A, Zinger T, Inglima K, Woo KM, Atie O, Yurasits L, See B, Aguero-Rosenfeld ME. Performance of Abbott ID Now COVID-19 Rapid Nucleic Acid Amplification Test Using Nasopharyngeal Swabs Transported in Viral Transport Media and Dry Nasal Swabs in a New York City Academic Institution. J Clin Microbiol. 2020 Jul 23;58(8):e01136-20. DOI: 10.1128/JCM.01136-20 Externer Link
71.
Cradic K, Lockhart M, Ozbolt P, Fatica L, Landon L, Lieber M, Yang D, Swickard J, Wongchaowart N, Fuhrman S, Antonara S. Clinical Evaluation and Utilization of Multiple Molecular In Vitro Diagnostic Assays for the Detection of SARS-CoV-2. Am J Clin Pathol. 2020 Jul;154(2):201-7. DOI: 10.1093/ajcp/aqaa097 Externer Link
72.
Smithgall MC, Scherberkova I, Whittier S, Green DA. Comparison of Cepheid Xpert Xpress and Abbott ID Now to Roche cobas for the Rapid Detection of SARS-CoV-2. J Clin Virol. 2020 Jul;128:104428. DOI: 10.1016/j.jcv.2020.104428 Externer Link
73.
Mitchell SL, George KS. Evaluation of the COVID19 ID NOW EUA assay. J Clin Virol. 2020 Jul;128:104429. DOI: 10.1016/j.jcv.2020.104429 Externer Link
74.
Hogan CA, Garamani N, Lee AS, Tung JK, Sahoo MK, Huang C, Stevens B, Zehnder J, Pinsky BA. Comparison of the Accula SARS-CoV-2 Test with a Laboratory-Developed Assay for Detection of SARS-CoV-2 RNA in Clinical Nasopharyngeal Specimens. J Clin Microbiol. 2020 Jul;58(8):e01072-20. DOI: 10.1128/JCM.01072-20 Externer Link
75.
Xie J, Ding C, Li J, Wang Y, Guo H, Lu Z, Wang J, Zheng C, Jin T, Gao Y, He H. Characteristics of patients with coronavirus disease (COVID-19) confirmed using an IgM-IgG antibody test. J Med Virol. 2020 Oct;92(10):2004-10. DOI: 10.1002/jmv.25930 Externer Link
76.
Xu Y, Xiao M, Liu X, Xu S, Du T, Xu J, Yang Q, Xu Y, Han Y, Li T, Zhu H, Wang M. Significance of serology testing to assist timely diagnosis of SARS-CoV-2 infections: implication from a family cluster. Emerg Microbes Infect. 2020 Dec;9(1):924-7. DOI: 10.1080/22221751.2020.1752610 Externer Link
77.
Hou H, Wang T, Zhang B, Luo Y, Mao L, Wang F, Wu S, Sun Z. Detection of IgM and IgG antibodies in patients with coronavirus disease 2019. Clin Transl Immunology. 2020 May;9(5):e01136. DOI: 10.1002/cti2.1136 Externer Link
78.
US Food & Drug Administration. In Vitro Diagnostics EUAs [last accessed 2020 Dec 21]. Available from: https://www.fda.gov/medical-devices/coronavirus-disease-2019-covid-19-emergency-use-authorizations-medical-devices/vitro-diagnostics-euas Externer Link
79.
Young S, Taylor SN, Cammarata CL, Varnado KG, Roger-Dalbert C, Montano A, Griego-Fullbright C, Burgard C, Fernandez C, Eckert K, Andrews JC, Ren H, Allen J, Ackerman R, Cooper CK. Clinical Evaluation of BD Veritor SARS-CoV-2 Point-of-Care Test Performance Compared to PCR-Based Testing and versus the Sofia 2 SARS Antigen Point-of-Care Test. J Clin Microbiol. 2020 Dec;59(1):e02338-20. DOI: 10.1128/JCM.02338-20 Externer Link
80.
Perchetti GA, Huang ML, Mills MG, Jerome KR, Greninger AL. Analytical Sensitivity of the Abbott BinaxNOW COVID-19 Ag Card. J Clin Microbiol. 2021 Feb;59(3):e02880-20. DOI: 10.1128/JCM.02880-20 Externer Link
81.
Li Z, Yi Y, Luo X, Xiong N, Liu Y, Li S, Sun R, Wang Y, Hu B, Chen W, Zhang Y, Wang J, Huang B, Lin Y, Yang J, Cai W, Wang X, Cheng J, Chen Z, Sun K, Pan W, Zhan Z, Chen L, Ye F. Development and clinical application of a rapid IgM-IgG combined antibody test for SARS-CoV-2 infection diagnosis. J Med Virol. 2020 Sep;92(9):1518-24. DOI: 10.1002/jmv.25727 Externer Link
82.
Candel FJ, Vinuela-Prieto JM, González Del Castillo J, Barreiro García P, Fragiel Saavedra M, Hernández Píriz A, Jiménez Virumbrales D, Canora Lebrato J, García de Casasola G, Gil Prieto R, San-Román Montero J, Ortega Anselmi J, Barba Martín R, Prados Roa F, Marco Martínez J, Zapatero Gaviria A. Utility of lateral flow tests in SARS-CoV-2 infection monitorization. Rev Esp Quimioter. 2020 Aug;33(4):258-66. DOI: 10.37201/req/052.2020 Externer Link
83.
Dellière S, Salmona M, Minier M, Gabassi A, Alanio A, Le Goff J, Delaugerre C, Chaix ML; Saint-Louis CORE (COvid REsearch) group. Evaluation of the COVID-19 IgG/IgM Rapid Test from Orient Gene Biotech. J Clin Microbiol. 2020 Jul;58(8). DOI: 10.1128/JCM.01233-20 Externer Link
84.
Choe JY, Kim JW, Kwon HH, Hong HL, Jung CY, Jeon CH, Park EJ, Kim SK. Diagnostic performance of immunochromatography assay for rapid detection of IgM and IgG in coronavirus disease 2019. J Med Virol. 2020 Nov;92(11):2567-72. DOI: 10.1002/jmv.26060 Externer Link
85.
Ong DSY, de Man SJ, Lindeboom FA, Koeleman JGM. Comparison of diagnostic accuracies of rapid serological tests and ELISA to molecular diagnostics in patients with suspected coronavirus disease 2019 presenting to the hospital. Clin Microbiol Infect. 2020 Aug;26(8):1094.e7-1094.e10. DOI: 10.1016/j.cmi.2020.05.028 Externer Link
86.
Hoffman T, Nissen K, Krambrich J, Rönnberg B, Akaberi D, Esmaeilzadeh M, Salaneck E, Lindahl J, Lundkvist Å. Evaluation of a COVID-19 IgM and IgG rapid test; an efficient tool for assessment of past exposure to SARS-CoV-2. Infect Ecol Epidemiol. 2020;10(1):1754538. DOI: 10.1080/20008686.2020.1754538 Externer Link
87.
Kang K, Huang L, Ouyang C, Du J, Yang B, Chi Y, He S, Ying L, Chen G, Wang J. Development, performance evaluation, and clinical application of a Rapid SARS-CoV-2 IgM and IgG Test Kit based on automated fluorescence immunoassay. J Med Virol. 2021 May;93(5):2838-47. DOI: 10.1002/jmv.26696 Externer Link
88.
Burbelo PD, Riedo FX, Morishima C, Rawlings S, Smith D, Das S, Strich JR, Chertow DS, Davey RT Jr, Cohen JI. Detection of Nucleocapsid Antibody to SARS-CoV-2 is More Sensitive than Antibody to Spike Protein in COVID-19 Patients [Preprint]. medRxiv. 2020 Apr 24. DOI: 10.1101/2020.04.20.20071423 Externer Link
89.
Van Elslande J, Houben E, Depypere M, Brackenier A, Desmet S, André E, Van Ranst M, Lagrou K, Vermeersch P. Diagnostic performance of seven rapid IgG/IgM antibody tests and the Euroimmun IgA/IgG ELISA in COVID-19 patients. Clin Microbiol Infect. 2020 Aug;26(8):1082-7. DOI: 10.1016/j.cmi.2020.05.023 Externer Link
90.
Kohmer N, Westhaus S, Rühl C, Ciesek S, Rabenau HF. Clinical performance of different SARS-CoV-2 IgG antibody tests. J Med Virol. 2020 Oct;92(10):2243-7. DOI: 10.1002/jmv.26145 Externer Link
91.
Jawhara S. Could Intravenous Immunoglobulin Collected from Recovered Coronavirus Patients Protect against COVID-19 and Strengthen the Immune System of New Patients? Int J Mol Sci. 2020 Mar;21(7):2272. DOI: 10.3390/ijms21072272 Externer Link
92.
Anderson DE, Tan CW, Chia WN, Young BE, Linster M, Low JH, Tan YJ, Chen MI, Smith GJD, Leo YS, Lye DC, Wang LF. Lack of cross-neutralization by SARS patient sera towards SARS-CoV-2. Emerg Microbes Infect. 2020 Dec;9(1):900-2. DOI: 10.1080/22221751.2020.1761267 Externer Link
93.
Nie J, Li Q, Wu J, Zhao C, Hao H, Liu H, Zhang L, Nie L, Qin H, Wang M, Lu Q, Li X, Sun Q, Liu J, Fan C, Huang W, Xu M, Wang Y. Establishment and validation of a pseudovirus neutralization assay for SARS-CoV-2. Emerg Microbes Infect. 2020 Dec;9(1):680-6. DOI: 10.1080/22221751.2020.1743767 Externer Link
94.
European Commission. COVID-19 In Vitro Diagnostic Devices and Test Methods Database [last accessed 2020 Dec 21]. Available from: https://covid-19-diagnostics.jrc.ec.europa.eu/devices/7?marking=&principle=NucleicAcid-PCR%20based&format=&manufacturer=&text_name=#form_content Externer Link