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      Point-of-Care Diagnostic Tests for Detecting SARS-CoV-2 Antibodies: A Systematic Review and Meta-Analysis of Real-World Data

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          SARS-CoV-2 is responsible for a highly contagious infection, known as COVID-19. SARS-CoV-2 was discovered in late December 2019 and, since then, has become a global pandemic. Timely and accurate COVID-19 laboratory testing is an essential step in the management of the COVID-19 outbreak. To date, assays based on the reverse-transcription polymerase chain reaction (RT-PCR) in respiratory samples are the gold standard for COVID-19 diagnosis. Unfortunately, RT-PCR has several practical limitations. Consequently, alternative diagnostic methods are urgently required, both for alleviating the pressure on laboratories and healthcare facilities and for expanding testing capacity to enable large-scale screening and ensure a timely therapeutic intervention. To date, few studies have been conducted concerning the potential utilization of rapid testing for COVID-19, with some conflicting results. Therefore, the present systematic review and meta-analysis was undertaken to explore the feasibility of rapid diagnostic tests in the management of the COVID-19 outbreak. Based on ten studies, we computed a pooled sensitivity of 64.8% (95%CI 54.5–74.0), and specificity of 98.0% (95%CI 95.8–99.0), with high heterogeneity and risk of reporting bias. We can conclude that: (1) rapid diagnostic tests for COVID-19 are necessary, but should be adequately sensitive and specific; (2) few studies have been carried out to date; (3) the studies included are characterized by low numbers and low sample power, and (4) in light of these results, the use of available tests is currently questionable for clinical purposes and cannot substitute other more reliable molecular tests, such as assays based on RT-PCR.

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          Performance of VivaDiag COVID‐19 IgM/IgG Rapid Test is inadequate for diagnosis of COVID‐19 in acute patients referring to emergency room department

          To the Editor, From late December 2019, coronavirus infectious disease (COVID‐19) epidemics spread from Wuhan, China, to all over the world, including Italy. 1 , 2 , 3 To date, real‐time reverse transcription‐polymerase chain reaction (RT‐PCR) in respiratory samples is the current gold standard method for the diagnosis of COVID‐19. 4 , 5 However, molecular testings are time consuming and require specialized operators, factors that limit their use in real life when the rapid diagnosis is required for fast intervention decisions. Recently, an easy to perform serological assay has been assessed 6 to differentiate COVID‐19 positive patients from negative subjects. We herein report results of a real‐life study performed in an emergency room department of a tertiary hospital in Northern Italy to validate VivaDiag COVID‐19 IgM/IgG Rapid Test lateral flow immunoassay (LFIA) for the rapid diagnosis of COVID‐19. Overall 110 subjects were tested for COVID‐19‐specific serological assay at Fondazione IRCCS Policlinico San Matteo. In detail, we enrolled 30 healthy volunteers with documented negative results for COVID‐19 RT‐PCR in respiratory samples (M 11/F 19; median age, 38.5; range, 25‐69 years). Ten of them (33.3%) had been infected in the past with one of the common OC43, 229E, HKU1, and NL63 coronavirus. Thirty COVID‐19‐positive patients (25 M/5 F; median age, 73.5; range, 38‐86 years) admitted to the Infectious Diseases Department or at the Intensive Care Unit were tested as positive controls. Finally, the performance of VivaDiag COVID‐19 IgM/IgG Rapid Test LFIA was tested in 50 patients at their first access at emergency room department with fever and respiratory syndrome (34 M/16 F; median age, 61.50; range, 33‐97 years) in comparison with results of nasal swab molecular screening. 5 VivaDiag COVID‐19 IgM/IgG from VivaChek was performed according to manufacturer's instruction by adding 10 µL of serum or whole blood sample into the sample port followed by adding 2 to 3 drops (70‐100 µL) of dilution buffer. 6 After about 15 minutes, results were read. Respiratory samples (FLOQSwabs; Copan Italia, Brescia, Italy) were collected from all the patients. Total nucleic acids (DNA/RNA) were extracted from 200 µL of UTM using the QIAsymphony instrument with QIAsymphony DSP Virus/Pathogen Midi Kit (complex 400 protocols) according to the manufacturer's instructions (QIAGEN; Qiagen, Hilden, Germany). Specific real‐time RT‐PCR targeting RNA‐dependent RNA polymerase and E genes were used to detect the presence of SARS‐CoV‐2 according to the WHO guidelines 7 and Corman et al 5 protocols. In the cohort of patients admitted to the emergency room department, data from serological tests were compared to molecular results to define specificity, sensitivity, positive predictive value (PPV), and negative predictive value (NPV) of the rapid serological test. As expected, all 30 COVID‐19 negative volunteers were negative for both immunoglobulin G (IgG) and immunoglobulin M (IgM) using the VivaDiag COVID‐19 IgM/IgG Rapid Test. No cross‐reactivity was detected in the 10 subjects with previous coronaviruses infection, supporting the high specificity of the VivaDiag COVID‐19 IgM/IgG Rapid Test LFIA. Serum samples were obtained at a median 7 days (interquartile range, 4‐11) after the first COVID‐19 positive result from 30 hospitalized patients. A total of 19 of 30 (63.3%) were positive for both IgM and IgG, 5 of 30 (16.7%) were negative for both IgG and IgM, 5 of 30 (16.7%) were weakly positive for both IgM and IgG, and only 1 of 30 (3.3%) was positive for IgM and negative for IgG. Thus, the sensitivity of the rapid assay was suboptimal (data not are shown). A possible explanation is the low antibody titers or a delayed humoral response. 6 Focusing on acute patients enrolled from the emergency room department, 12 of 50 (24%) were negative for COVID‐19 by real‐time RT‐PCR. Of these, 1 (8.3%) showed a positive results for the VivaDiag COVID‐19 IgM/IgG Rapid Test, while the other 11 of 12 (91.7%) tested negative. On the other side, 38 patients were positive for COVID‐19 by real‐time RT‐PCR. Of these, only 7 (18.4%) showed a positive or weak positive serology for IgM and/or IgG, while the other 31 of 38 (81.6%) tested negative for the rapid serology assay (Table 1). Thus, the sensitivity of the VivaDiag COVID‐19 IgM/IgG Rapid Test was 18.4%, specificity was 91.7%, while NPV was 26.2%, and PPV was 87.5% in patients enrolled from emergency room department. In contrast with the high levels of sensitivity reported in the previous study, 6 VivaDiag COVID‐19 IgM/IgG Rapid Test revealed a very poor sensitivity (less than 20%). Indeed, the majority of patients that tested positive for COVID‐19 by real‐time RT‐PCR would have been identified as negative using only the rapid serological assay, leading to a misdiagnosis of COVID‐19 disease in the vast majority of patients. On the basis of our results, VivaDiag COVID‐19 IgM/IgG Rapid Test LFIA is not recommended for triage of patients with suspected COVID‐19. Table 1 Characteristics and VivaDiag COVID‐19 IgM/IgG Rapid Test results of 50 consecutive patients referred to the emergency room department Patient Sex Age Result of COVID‐19 real‐time RT‐PCR on NS VivaDiag COVID‐19 IgM/IgG Rapid Test IgM IgG 1 M 33 neg − − 2 M 51 pos − − 3 M 51 pos − − 4 M 38 pos − − 5 F 80 pos − − 6 F 64 neg − − 7 M 81 neg − − 8 M 76 pos +/− − 9 M 33 pos − − 10 M 37 neg − − 11 F 45 pos − − 12 M 53 pos − − 13 M 66 neg − − 14 M 78 pos − − 15 F 97 pos − − 16 M 38 pos − − 17 M 72 pos − − 18 M 56 pos − − 19 M 80 pos − +/− 20 M 72 pos − − 21 F 55 pos − − 22 M 82 pos − − 23 M 47 pos + +/− 24 F 63 pos − − 25 F 80 pos +/− − 26 M 59 pos − − 27 M 66 pos − − 28 M 39 pos − − 29 F 78 neg − − 30 M 71 neg − − 31 F 46 neg − − 32 F 51 pos − − 33 F 75 pos − − 34 F 82 pos + +/− 35 F 51 pos +/− +/− 36 M 84 pos − − 37 M 50 pos − − 38 M 50 pos + +/− 39 F 72 neg − − 40 M 54 neg − − 41 F 64 neg + − 42 M 64 pos − − 43 M 70 pos − − 44 M 56 pos − − 45 M 68 pos − − 46 F 36 pos − − 47 M 60 pos − − 48 M 66 pos − − 49 M 54 neg − − 50 M 56 pos − − Abbreviations: −, negative result; +, positive result; +/−, weakly positive result; COVID‐19, coronavirus infectious disease 2019; IgG, immunoglobulin G; IgM, immunoglobulin M; NS, nasopharyngeal swab; RT‐PCR, reverse transcription‐polymerase chain reaction. John Wiley & Sons, Ltd. This article is being made freely available through PubMed Central as part of the COVID-19 public health emergency response. It can be used for unrestricted research re-use and analysis in any form or by any means with acknowledgement of the original source, for the duration of the public health emergency. MEMBERS OF THE SAN MATTEO PAVIA COVID‐19 TASK FORCE R. Bruno, M. Mondelli, E. Brunetti, A. Di Matteo, E. Seminari, L. Maiocchi, V. Zuccaro, L. Pagnucco, B. Mariani, S. Ludovisi, R. Lissandrin, A. Parisi, P. Sacchi, S. F. A. Patruno, G. Michelone, R. Gulminetti, D. Zanaboni, S. Novati, R. Maserati, P. Orsolini, and M. Vecchia (ID Staff); M. Sciarra, E. Asperges, M. Colaneri, A. Di Filippo, M. Sambo, S. Biscarini, M. Lupi, S. Roda, T. C. Pieri, I. Gallazzi, M. Sachs, and P. Valsecchi (ID Resident); S. Perlini, C. Alfano, M. Bonzano, F. Briganti, G. Crescenzi, A. G. Falchi, R. Guarnone, B. Guglielmana, E. Maggi, I. Martino, P. Pettenazza, S. Pioli di Marco, F. Quaglia, A. Sabena, F. Salinaro, F. Speciale, and I. Zunino (ECU Staff Emergency Care Unit); M. De Lorenzo, G. Secco, L. Dimitry, G. Cappa, I. Maisak, B. Chiodi, M. Sciarrini, B. Barcella, F. Resta, L. Moroni, G. Vezzoni, L. Scattaglia, E. Boscolo, C. Zattera, M. F. Tassi, V. Capozza, D. Vignaroli, and M. Bazzini (ECU Resident Emergency Care Unit); G. Iotti, F. Mojoli, M. Belliato, L. Perotti, S. Mongodi, and G. Tavazzi (Intensive Care Unit); G. Marseglia, A. Licari, and I. Brambilla (Pediatric Unit); D. Barbarini, A. Bruno, P. Cambieri, G. Campanini, G. Comolli, M. Corbella, R. Daturi, M. Furione, B. Mariani, R. Maserati, E. Monzillo, S. Paolucci, M. Parea, E. Percivalle, A. Piralla, F. Rovida, A. Sarasini, and M. Zavattoni (Virology Staff); G. Adzasehoun, L. Bellotti, E. Cabano, G. Casali, L. Dossena, G. Frisco, G. Garbagnoli, A. Girello, V. Landini, C. Lucchelli, V. Maliardi, S. Pezzaia, and M. Premoli (Virology Technical staff); A. Bonetti, G. Caneva, I. Cassaniti, A. Corcione, R. Di Martino, A. Di Napoli, A. Ferrari, G. Ferrari, L. Fiorina, F. Giardina, A. Mercato, F. Novazzi, G. Ratano, B. Rossi, I. M. Sciabica, M. Tallarita, and E. Vecchio Nepita (Virology Resident); M. Calvi and M. Tizzoni (Pharmacy Unit); and C. Nicora, A. Triarico, V. Petronella, C. Marena, A. Muzzi, and P. Lago (Hospital Management). CONFLICT OF INTERESTS The authors declare that there are no conflict of interests. AUTHOR CONTRIBUTIONS IC, FN, FG, FS, MS, SP, RB, FM, FB, and the other members of the San Matteo Pavia COVID‐19 Task Force listed reviewed and approved the manuscript. IC and FN discussed results, data analysis, and wrote the paper. FG, FS, and MS collected the samples. SP, RB, and FM discussed results. FB conceived the study.
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            Comparison of Abbott ID Now and Abbott m2000 Methods for the Detection of SARS-CoV-2 from Nasopharyngeal and Nasal Swabs from Symptomatic Patients

            LETTER The ID Now COVID-19 (IDNCOV) assay performed on the ID Now instrument (Abbott Diagnostics, Inc., Scarborough, ME) is a rapid diagnostic test that can be performed in a point-of-care setting equivalent to Clinical Laboratory Improvement Amendments (CLIA)-waived testing. The assay utilizes isothermal amplification and can reportedly deliver results in approximately 5 to 13 min. As this assay could provide significant improvements to workflow in our hospital system, we sought to compare the performance of this test with our current coronavirus disease 2019 (COVID-19) assay, the Abbott RealTime SARS-CoV-2 (severe acute respiratory syndrome coronavirus 2) (ACOV) assay performed on the Abbott m2000 system (Abbott Molecular Inc., Des Plaines, IL). We compared the results from 524 paired foam nasal swabs (NS) tested on IDNCOV with nasopharyngeal swabs (NPS) placed in viral transport media tested on ACOV collected consecutively from symptomatic patients meeting current criteria for a diagnosis of COVID-19 (1). Five locations were included in the evaluation including three emergency departments (ED) and two immediate care centers (IMCC). IMCC A and ED 2 were experienced users of the IDNow platform. The other sites were new users of the platform and received training specifically for the IDNCOV. All ACOV testing was performed by one central clinical laboratory, and all NPS were heat inactivated for 30 min at 60°C prior to testing. NS were tested directly on the IDNCOV from IMCCs, and the tests were performed on-site. NS from the EDs were transported to the clinical microbiology laboratory in sterile transport containers (urine cups or conical tubes) and tested by laboratory personnel at each separate location. Statistical analysis was performed using SPSS v.26. The overall positivity rate in this sample collection was 35%, ranging from 22% to 60% among the five sites. Overall agreement was 75% positive agreement (95% confidence interval [95% CI], 67.74%, 80.67%) and 99% negative agreement (95% CI, 97.64%, 99.89%) between IDNCOV and ACOV for all specimens tested. Agreement at individual sites varied (Table 1). Two subjects tested positive on IDNCOV that were initially negative on ACOV. In case one, a repeat sample was positive on ACOV (repeat IDNCOV was not performed), and the case was resolved as a true positive result. For case two, all repeat testing (both IDNCOV and ACOV) was negative and was resolved as a likely false-positive result. This sample was collected during the first day of testing and could have been operator error. TABLE 1 Agreement between ACOV and IDNCOV Site Total no. of samples tested No. of samples with the following result a : % Positivity Positive agreement (95% CI) Negative agreement (95% CI) Performance agreement (kappa) (95% CI) A+/IND+ A+/IND− A−/IND+ A−/IND− IMCC A 208 33 13 1 161 22 71.74 (56.32, 83.54) 99.38 (96.09, 99.97) 0.783 (0.779, 0.788) IMCC B 125 39 17 0 69 44 69.64 (55.74, 80.84) 100.0 (93.43, 100.0) 0.711 (0.706, 0.717) ED 1 105 26 11 0 68 35 70.27 (52.83, 83.56) 100.0 (93.33, 100.0) 0.751 (0.744, 0.757) ED 2 31 12 3 0 16 50 80.0 (51.37, 94.69) 100 (75.92, 100.0) 0.803 (0.792, 0.814) ED 3 55 29 3 1 22 60 90.63 (73.83, 97.55) 95.65 (76.03, 99.77) 0.852 (0.844, 0.861) Total 524 139 47 2 336 35 74.73 (67.74, 80.67) 99.41 (97.64, 99.89) a Positive (+) and negative (−) results by ACOV (A) and IDNCOV (IND) are shown. Fleiss kappa analysis comparing the performance at each of the sites demonstrated that strength of agreement between the sites (Table 1) was rated as good to very good with comparable standard errors. We interpret this to mean that a site’s ability to run the test (or lack of experience) did not necessarily contribute to the variability in positivity that was found in this evaluation. Compared to the ACOV cycle numbers (CN) (which are similar but not directly comparable to cycle thresholds from other reverse transcription-PCR [RT-PCR] assays due to the unique ACOV assay design), a significant proportion, but not all, discordant samples exhibited at higher cycle numbers (Fig. 1). The mean CN for concordant positive samples was 12.71 (95% CI, 11.76, 13.67), ranging from 2.99 to 31.01, with a standard deviation of 5.5. The mean CN for discordant samples (ACOV positive [ACOV+]/IDNCOV negative [IDNCOV−]) was 21.07 (95% CI, 19.55, 22.60), ranging from 6.79 to 30.63, with a standard deviation of 5.1. These differences are statistically different (P = 6.75e−16). The stated limit of detection in the published instructions for use is 100 copies/ml for ACOV (2) and approximately 3,225 copies/ml when calculated based on the published genomes/reaction for IDNCOV (3). Based on the distribution of cycle numbers seen in Fig. 1 and performance agreement among the sites, negative results on IDNCOV are likely related to both a higher limit of detection on IDNCOV and preanalytical sampling error. FIG 1 Boxplot of cycle numbers of concordant and discordant paired results. Distribution of cycle numbers from IDNCOV-positive/ACOV-positive samples (including a single data point [CN 31.01] outlier beyond the standard error) compared to INDCOV-negative/ACOV-positive samples. Overall, the ID Now COVID-19 assay demonstrated significantly different performance characteristics compared to the Abbott RealTime SARS-CoV-2 assay.
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              Rapid point-of-care testing for SARS-CoV-2 in a community screening setting shows low sensitivity

              Objective With the current SARS-CoV2 outbreak, countless tests need to be performed on potential symptomatic individuals, contacts and travellers. The gold standard is a quantitative polymerase chain reaction (qPCR)–based system taking several hours to confirm positivity. For effective public health containment measures, this time span is too long. We therefore evaluated a rapid test in a high-prevalence community setting. Study design Thirty-nine randomly selected individuals at a COVID-19 screening centre were simultaneously tested via qPCR and a rapid test. Ten previously diagnosed individuals with known SARS-CoV-2 infection were also analysed. Methods The evaluated rapid test is an IgG/IgM–based test for SARS-CoV-2 with a time to result of 20 min. Two drops of blood are needed for the test performance. Results Of 49 individuals, 22 tested positive by repeated qPCR. In contrast, the rapid test detected only eight of those positive correctly (sensitivity: 36.4%). Of the 27 qPCR-negative individuals, 24 were detected correctly (specificity: 88.9%). Conclusion Given the low sensitivity, we recommend not to rely on an antibody-based rapid test for public health measures such as community screenings.

                Author and article information

                J Clin Med
                J Clin Med
                Journal of Clinical Medicine
                18 May 2020
                May 2020
                : 9
                : 5
                [1 ]AUSL–IRCCS di Reggio Emilia, Servizio di Prevenzione e Sicurezza negli ambienti di Lavoro (SPSAL), I-42122 Reggio Emilia (RE), Italy
                [2 ]ASL Foggia, Dipartimento di Prevenzione, Servizio Prevenzione e Sicurezza Ambienti Lavoro, Piazza Pavoncelli 11, I-41121 Foggia (FG), Italy; dott.pietro.ferraro@
                [3 ]Department of Medicine and Surgery, School of Medicine, University of Parma, 43123 Parma (PR), Italy; gualerzi@
                [4 ]Department of Medicine and Surgery, School of Occupational Medicine, University of Parma, I-43123 Parma (PR), Italy; silvia.ranzieri@
                [5 ]Cardiac Intensive Care Unit, The Heart Institute, Cincinnati Children’s Hospital Medical Center, 3333 Burnet Avenue, Cincinnati, OH 45229-3026, USA; brandon.henry@
                [6 ]Department of Industrial Pharmacy, Sechenov First Moscow State Medical University (Sechenov University), 119991 Moscow, Russia; younisbensaid@ (Y.B.S.); osipova-mma@ (N.V.P.)
                [7 ]Executive Director of the Union “National Pharmaceutical Chamber”, 125009 Moscow, Russia; rapalatainfo@
                [8 ]Laboratory for Industrial and Applied Mathematics (LIAM), Department of Mathematics and Statistics, York University, 4700 Keele Street, Toronto, ON M3J 1P3, Canada; wujh@
                [9 ]School of Medicine, Vita-Salute San Raffaele University, 20132 Milan (MI), Italy; signorelli.carlo@
                Author notes
                [* ]Correspondence: mricco2000@ (M.R.); bragazzi@ (N.L.B.)

                Equal contribution.

                © 2020 by the authors.

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