Following the outbreak of the COVID-19 pandemic, assays using lateral flow, ELISA, chemiluminescence, and fluorescence technologies were developed to detect antibodies to SARS-CoV-2 antigens.1-4 Target proteins include the SARS-CoV-2 nucleocapsid (NC) phosphoprotein,5 the full-length spike glycoprotein,6 or the receptor-binding domain (RBD) of the spike protein.7 The underlying premise is that such antibody assays can be used to evaluate the epidemiology of COVID-19 and, potentially, identify individuals and communities who would be protected from reinfection and have higher rates of herd immunity, respectively. However, it remains unclear which antibodies are actually protective or how long antibody levels are sustained.8 Notably, the majority of tests currently in use have reported sensitivities and specificities ranging from 96% to 98%, which, while acceptable, are especially concerning when evaluating individuals from regions with <5% prevalence of disease.9,10 Under those circumstances, a positive result has a high probability to be falsely positive, especially in a situation of low population prevalence. Since the majority of FDA approved serological assays only detect 1 viral target per test, even a true positive result has limited meaning, as only certain antibodies to SARS-CoV-2 appear to have viral neutralizing activity.11 Thus, simply considering a patient as SARS-CoV-2 antibody positive provides, at best, incomplete and at worst, misleading information regarding the clinical implications of those antibodies.
The limitations of the current assays to detect antibodies to SARS-CoV-2 become even more apparent when considering the spectrum of clinical responses to COVID-19. Clearly, certain patient characteristics are associated with an increased risk of morbidity and mortality.12,13 For example, recipients of solid organ allografts who become infected with SARS-CoV-2 are at especially high risk for adverse consequences compared to matched controls.14,15 This is presumably a consequence of being immunosuppressed and possessing other underlying health issues.16 Knowing the SARS-CoV-2 antibody status of patients before and after transplant could affect patient care.
Transplant candidates and recipients are routinely monitored to evaluate their HLA antibody status pretransplantation and posttransplantation. The majority of histocompatibility laboratories around the world use a Luminex platform to detect and identify the HLA antibodies these patients possess.17 The HLA Luminex-based test is a multiplex, solid-phase antibody detection assay where up to 100 different microparticles coated with individual class I or class II HLA alleles, respectively, are simultaneously tested with patient sera. Due to its sensitivity, specificity, and high throughput, this multiplex bead assay revolutionized the field of organ transplantation.18 In a similar manner, a Luminex-based multiplex test designed to identify antibody responses to unique SARS-CoV-2 targets could dramatically enhance our understanding of the immune response to COVID-19. In this study, we describe the development and validation of a multiplex high-throughput solid phase assay that simultaneously determines the presence/absence of antibodies to 5 SARS-CoV-2 viral proteins, specifically, the full spike protein, 3 individual domains of the spike protein (S1, S2, and receptor binding domain), and the nucleocapsid protein. As the impact of COVID-19 among transplant candidates and recipients is only beginning to emerge, assays such as the one described here that can easily be performed in HLA/transplant laboratories will be an invaluable tool to further our understanding.
MATERIALS AND METHODS
A prototype multiplex bead-based kit (LabScreen COVID PLUS; Cat#: LSCOV01001) for the Luminex platform (Luminex, Corp., Austin, TX) was developed by One Lambda Inc., Inc. (West Hill, CA). The assay includes 4 distinct fragments of SARS-CoV-2 Spike proteins, namely (1) full spike extracellular domain; (2) spike S1; (3) spike, RBD; and (4) spike S2; The fifth target is the SARS-CoV-2 NC protein. The amino acid sequence location of each SARS-CoV-2 protein utilized is shown in Figure S1, SDC, https://links.lww.com/TP/C47. Additionally, the kit incorporates Spike S1 fragments from 6 other coronaviruses, namely HCoV-229E, HCoV-HKU1, HCoV-NL63, HCoV-OC43, MERS-CoV, and SARS-CoV. Each of the above proteins were conjugated to individual beads. Additionally, a positive control bead, coated with human IgG, and a negative control bead, coated with human albumin were included. In total, the assay comprised 13 individually protein coated beads. The viral proteins were obtained either from commercial sources namely, Sino Biologicals (Wayne, PA), Euprotein, Inc. (North Brunswick, NJ), RayBiotech (Peachtree Corners, GA), or prepared in house (One Lambda Inc., Inc. West Hills, CA). The source and catalog numbers for each of these proteins is shown in Table S1, SDC, https://links.lww.com/TP/C47. The SARS-CoV-2 Full Spike, Spike S2, and SARS-CoV Spike S1were produced in-house based on published sequences.19,20
Polyclonal Antisera for Protein Validation
Polyclonal antisera were used to validate expression of the above proteins after their conjugation to individual beads. Antisera were purchased from a single commercial source, Sino Biologicals (Wayne, PA). The polyclonal antibodies used for bead validation were: SARS-CoV-2 NC Protein (Cat.#: 40588-T62, 1:1000 dilution), SARS-CoV-2 Spike (Cat#: 40589-T62; 1:1000 dilution), SARS-CoV-S2 (Cat#: 40590-T62; 1:1000 dilution), SARS-CoV S1 (Cat#: 40150-T62, 1:1000 dilution), SARS-CoV NC Protein (Cat#: 40143-T62, 1:1000 dilution), MERS-S1 (Cat#: 40069-T52, 1:1000 dilution), MERS NC Protein (Cat#: 40068-RP02, 1:1000 dilution), MERS Spike (CAT#: 40069-T30, 1:1000 dilution), MERS S2 (Cat#: 40070-T60, 1:1000 dilution), and HCoV-HKU1 S1 (CAT#: 40021-RP01, 1:1000 dilution).
Serum samples from both presumed negative patients (ie, pre-COVID-19), and confirmed COVID-19 positive patients were used in this study. Samples were provided by Emory University and the University of Washington. Initially, 96 serum samples from candidates on the UNOS waitlist during a pre-COVID-19 time period (October 2018) were randomly selected. Subsequently, serum from 42 convalescent patients with confirmed SARS-CoV-2 infection (none of them transplant candidates or recipients) were also selected for testing. SARS-Cov-2 infection was confirmed either by PCR testing (University of Washington or Emory University) or by a SARS-CoV-2 antibody testing by an internally developed and validated ELISA assay (Emory University). Last, archived serum samples from 51 randomly selected candidates on the UNOS waitlist in May 2020 and distinct from the original 96 used for test verification, were selected for testing. Georgia recorded its first cases of COVID-19 on March 3, 2020 and the first State-wide peak occurred on April 7 (https://dph.georgia.gov/covid-19-daily-status-report). These 51 samples also were tested for SARS-CoV-2 RBD antibodies by ELISA at Stanford University.
This study was exempt from ethics approval by the Institutional Review Board at Emory University Hospital.
SARS-CoV-2 Antibody Detection in Patient Sera
Antibody detection on antigen-coated microparticles was performed as follows: 5 µl of viral antigen coated beads were admixed with 20 μl of a 1:10 dilution, in PBS, of patient or control serum that was pretreated with 10 mmol/L EDTA. The serum/bead admixture was incubated in the dark at 20 °C–25 °C for 30 min with gentle rotation followed by 3 sequential washes, each using 200-μl wash buffer (WB) (OLI Cat. # LSPWABUF). Following incubation, 100 µl of a pretitered PE-conjugated anti-human IgG (Jackson ImmunoResearch, West Grove, PA: CAT#: 109-116-170) was added. Next, the beads were vortexed and incubated, in the dark, for 30 min at 20 °C–25 °C with gentle shaking. Finally, the beads were washed twice with 200-μl WB. Microparticles were resuspended in 80 μl of 1X PBS and analyzed on a Luminex FLEXMAP 3D instrument (Luminex Corp. Austin, TX). Final analysis of mean fluorescence intensity (MFI) was performed using Microsoft Excel.
Calculation of Cutoff, Sensitivity, Specificity, and Accuracy
To determine the positive/negative cutoff values for each recombinant protein, we used the Mean +3×SD formula based on the mean MFI values of 96 COVID-19 negative samples collected before 2019. The 3×SD result obtained was considered as the cutoff value for each antigen.
Validation of SARS-CoV2-coated Microparticles
Microparticles coated with the SARS-CoV-2 proteins, Full Spike, S1, S2, RBD and NC as well as non-SARS-CoV-2 proteins from community coronaviruses (CoV-229E-S1, HCoV-HKU1-S1, HCoV-NL63-S1, HCoV-OC43-S1), and novel coronaviruses (MERS-S1 and SARS-S1) were evaluated with commercial polyclonal antibodies in a matrix fashion as shown in Figure 1 (SARS-CoV-2 proteins) and Figure S2, SDC, https://links.lww.com/TP/C47 (non-SARS-CoV-2 proteins). For the SARS-CoV-2 proteins, antisera specific for the NC showed positive reactivity with the NC-coated bead while failing to react with the other 4 SARS-CoV-2 proteins. The antisera directed against the Full Spike protein reacted with the Full Spike, S1, and RBD coated beads and was nonreactive with the S2 and NC. This reactivity pattern was in agreement with the manufactures published specifications for this antibody. The antisera directed against the SARS-CoV-2 S2 protein showed positive reactivity against both the Full Spike and S2 proteins as anticipated based on the manufacturers published specifications for this antibody. The antisera specific for the SARS-CoV-1 (SARS-S1) protein showed cross-reactivity against SARS-CoV-2 Full Spike, S1, and RBD. The antisera specific for SARS NC protein also showed cross-reactivity with the SARS-CoV-2 NC protein consistent with the published specifications for this antibody.
Pre-COVID-19 Samples Versus SARS-CoV-2 Proteins
Figure 2A shows data from a representative experiment, wherein archived serum samples, collected from 96 randomly selected and pre-COVID-19 (October, 2018) kidney transplant candidates were tested with the COVID Plus assay. As the data illustrate, with the exception of the S1 spike protein, all samples had MFI levels ≤5000. For the Full Spike protein, the average value was 675 ± 816; range 44–4540 MFI. For the spike S2 protein, the average value was 138 ± 97; range 42–908 MFI. For the spike RBD protein, the average value was 224 ± 363; range 23–3000 MFI. For the NC protein, the average value was 388 ± 441; range 89–2593 MFI. For the S1 protein, background levels were higher but all samples showed MFI values <7500. For the S1 spike, the average background value was 1226 ± 1411; range 133–7439 MFI. Based on these data, responses <5000 MFI for the Full Spike, S2, RBD, and NC-coated beads were considered negative, while values <7500 MFI were considered negative for S1. Figure 2B displays the responses for the 96 pre-COVID-19 samples against a panel of other coronavirus targets including SARS-CoV and MERS. The data demonstrate that while most individuals have significant IgG responses to community coronaviruses, all pre-COVID-19 samples tested negative against the SARS-CoV-2 targets as well as to SARS and MERS S1 proteins.
SARS-CoV-2 Positive Samples Versus SARS-CoV-2 Proteins
Sera from 42 nontransplanted patients each confirmed positive for SARS-CoV-2 infection by either RT-PCR positivity or SARS-CoV-2 antibody testing (ELISA, Emory University)18 were tested with the COVID-Plus assay. As shown in Figure 3, serum from all 42 patients tested positive with at least 3 of the 5 SARS-CoV-2 proteins. Positivity was determined as an MFI value >3 SDs above the mean of the negative controls. Interestingly, the level of responses differed from patient to patient. Specifically, MFI values ranged from 10 000 to 60 000 MFI for the Full Spike protein, 5000–50 000 MFI for S1, 2000–8000 for S2, 2000–30 000 for RBD, and 3000–15 000 for the NC protein. All 42 patients had a >5000 MFI response to the Full Spike, while 33/42, 38/42, and 39/42 patients had >5000 MFI response to S2, RBD, and NC, respectively. For S1, 38/42 samples showed a response >7500 MFI.
Differential Responsiveness of RT-PCR Positive Patients
The spectrum of response to SARS-CoV-2 proteins is illustrated in Figure 4. In Figure 4A, the semiquantitative MFI differences between and within individuals is plotted. Figure 4B illustrates the proportional difference among the COVID-19 positive patients. The data show that, on average, the Full Spike protein constituted the majority of the response in any given individual. However, the proportional differences between the 3 components of the Full Spike S1, S2, and RBD varied greatly between individuals. Of note, the proportional response to the RBD protein varied from 2% to 26% of the total MFI response. For cumulative MFI values <40 000, the RBD response ranged between 2% and 10% with a mean response proportion of 5.5%. However, for patients with a cumulative MFI value >40 000, the RBD response ranged between 12% and 26% with an average of 20%.
Screening of Serum Samples From Transplant Candidates Collected Post-COVID-19 Onset
As seen in Figure 5, 3 of 51 serum samples collected in May, 2020 from renal transplant candidates tested positive for at least 1 SARS-CoV-2 protein. Serum from 2 patients reacted with all 5 proteins (subjects #14 and #51), while 1 subject (#8) reacted only with the Full Spike protein. Interestingly, these same samples were tested for RBD antibodies by ELISA methodology. While patients 14 and 51 were positive, patient 8 was negative. To better understand these Luminex positive reactions, historic sera from these 3 subjects, dating back to pre-CoVID-19 time periods, were tested for reactivity in the COVID-Plus assay. Subjects #14 and #51 initially showed a distinct pattern of negative reactivity until May and April of 2020, respectively (Figure 6A and B). The positive results in April and May are consistent with COVID-19 infection during a period of relatively high virus prevalence in Georgia. In contrast, subject #8 showed reactivity only against the Full Spike protein and this reactivity appeared long before the global appearance of COVID-19 (Figure 7).
Herein, we describe the development of a novel multiplexed Luminex-based immunoassay that simultaneously and semiquantitatively assesses patient sera for antibodies to multiple SARS-CoV-2 proteins. This assay builds on >25 y of experience with the Luminex platform to detect and identify antibodies to HLA antigens present in the serum of transplant candidates and recipients. The strengths of this multiplexed, solid-phase assay compared with other platforms (eg, ELISA) include the ability to evaluate multiple viral targets simultaneously (and thereby provide internal assay controls for coronavirus specificity), the capacity to incorporate additional SARS-CoV-2 proteins in the future, a relatively short assay time, high-throughput and semiquantitative assessment of the antibody response. Furthermore, these data demonstrate that the antibody responses to common community coronaviruses do not cross react with the SARS-CoV-2 proteins in the COVID Plus assay (Figure 2). An additional benefit is that evaluation and monitoring of samples from transplant candidates and recipients for antibodies to SARS-CoV-2 is readily adaptable into the routine clinical practice of histocompatibility laboratories that support solid organ and/or stem cell transplant programs.
In the United States, the need for robust, accurate and reproducible serological tests to identify individuals who developed antibodies to SARS-CoV-2 was recognized by mid-March 2020. In response to demand, the FDA authorized emergency use authorization of a multitude of such tests to be developed, implemented and used throughout the United States. It was anticipated that the tests would identify individuals, (1) who had developed humoral immunity in response to exposure; (2) would be resistant to reinfection with SARS-CoV-2; and (3) from whom convalescent plasma could be collected and used as a therapeutic or prophylactic for patients diagnosed with COVID-19.21 However, recent studies reveal that a positive antibody test is not necessarily adequate to determine current or future immunity to SARS CoV-2. Indeed, recent data by Suthar et al22 documented the importance of appropriate timing for serological testing relative to PCR testing and/or symptom onset after infection. Additional studies by Chi et al23 revealed that although antibodies to the RBD of SARS CoV-2 are the antibodies most likely to be neutralizing, so, too, could antibodies to other viral targets. Thus, understanding which antibodies are protective and being able to detect them are both essential components of serological testing.
The genome of SARS-CoV-2, a single-stranded, enveloped RNA coronavirus, encodes 4 major structural proteins, namely spike, envelope, membrane, and nucleocapsid as well as more than a dozen nonstructural proteins.24 Based on their immunogenicity and predicted neutralizing potential, either the NC, spike (full length, S1, or S2) or the RBD are the individual designated targets for the vast majority of antibody assays in use. Sensitivity and specificity for such assays typically range from 96% to 98%, which, while reasonable, can unfortunately lead to false positive and false negative results.9,10 An additional important limitation of such single target assays is that they fail to assess the breadth of a patient’s response. In fact, recent studies reveal that patient antibody responses to SARS CoV-2 are not uniform. For example, while convalescent plasma donors and patients who have recovered from SARS-CoV-2 infection have detectable RBD antibodies that are neutralizing, RBD antibodies detected in pediatric COVID-19 patients who developed multisystem inflammatory syndrome are not neutralizing.11
The collective published data illustrate the gap in our understanding the complexities of the humoral immune response to COVID-19. Assays that interrogate a response to a single viral antigen limit our ability to understand fully the immune responsiveness to SARS-CoV-2. This is evidenced by the response of subject #8 who showed strong reactivity against the Full Spike protein but was negative for S1, S2, and RBD. The low level of NC protein reactivity also did not exceed the 3 SD cutoff to consider the reaction as positive. Since S1, S2, and RBD proteins are elements of the Full Spike protein; this reactivity could reflect responsiveness to a unique cross-reactive epitope not related to SARS-CoV-2 infection. This reactivity pattern was not observed in any other SARS-CoV-2 negative sample either pre- or post-COVID-19. Furthermore, the reactivity pattern of subject #8 extended well back into the global pre-COVID-19 era. The exact nature of this reactivity is under further investigation. When the results of all confirmed SARS-CoV-2 positive samples and negative samples obtained from samples before the COVID-19 pandemic are considered, the specificity and sensitivity of the assay are 98.6% and 100%, respectively. Excluding the data from the 1 subject positive only with the Full Spike protein, assay specificity and sensitivity are both 100%.
One recently described assay simultaneously evaluated the response to both nucleocapsid and spike proteins with a reported sensitivity of 100% and 99.9% specificity.25 However, that assay required patient samples to be diluted 1/200 before testing, which suggests that the signal-to-noise ratio of undiluted samples was less than optimal. Furthermore, the assay time was close to 20 h. In the multiplex assay described here, the responses to 5 different viral targets were simultaneously tested and the total assay time was <4 h. Furthermore, 96 samples can be assessed at the same time in a single tray.
Interestingly, the proportional responses to the 5 antigens differed among the patients. For example, some individuals displayed a higher MFI to NC versus RBD (eg, sample P18; 15 641 versus 8662, respectively), while other subjects displayed a higher response to the RBD target than to the NC (eg, sample P36; 33 491 versus 14 703, respectively). It is intriguing to speculate whether such observations would explain differences in clinical manifestations and outcomes. The differences could potentially be derived from immune deviation because of underlying disease or genetics. There may also be influences due to interindividual variations in B-cell repertoires, differences in disease progression, and the amount of time from which patients were initially diagnosed until the time they were tested for antibodies. Interrogation of antibody responses from well-characterized patients grouped according to age, severity of disease, and response to therapy will be improved by simultaneously evaluating the response to multiple SARS-CoV-2 proteins. Other important questions such as an individual’s antibody response to vaccination, viral neutralization titers and whether some antibodies enhance disease progression will be better addressed with this technology.
The semiquantitative nature of the response offered by this assay will also likely be of greater value rather than the simple “yes” or “no” result obtained from lateral flow assays. While beyond the scope of this article, the observation that 6 patients with Covid 19 (#s 5, 6, 9, 13, 14, and 18) had substantially lower levels of antibodies to SARS-CoV-2 proteins than the other 36 infected patients is intriguing and begs the question of whether patients with lower antibody levels had a different clinical course than those with higher levels. The titer of antibodies to SARS-CoV-2 correlates with their ability to limit viral infectivity. Our preliminary data suggest that due to the increased dynamic range of the Luminex FLEXMAP 3D instrument, multiplex testing for SARS-CoV-2 antibodies allows for a level of quantification not possible with lateral flow, ELISA, or chemiluminescence assays unless serial dilutions are performed for each sample.
While this first iteration of this multiplex Luminex based assay has 5 target proteins, our intention is to add additional targets to the panel. Indeed, the SARS-CoV-2 virus has 29 total proteins any of which could be immunogenic and stimulate an antibody response. Antibodies to nonstructural and accessory proteins of SARS-CoV-2, namely ORF9b and NSP5, have been identified in the convalescent sera of infected patients infected with SARS-CoV-2.26 Although viral targets not expressed on the cell surface are not typically considered to generate antibodies that are neutralizing, that is not always true. For example, for influenza, antibodies to ORF proteins confer protective immunity based on passive transfer studies and experiments with B-cell knockout mice.27,28 As the current version of COVID Plus only uses a total of thirteen individual Luminex beads, there is ample real estate for expansion. Additionally, we intend to expand the assay to include detection of IgM and IgA antibodies to SARS-CoV-2 proteins.
Limitations of this study include the source of our negative and positive control sera. Negative control samples were obtained from transplant candidates with chronic kidney disease. As such, the patients may be limited in their ability to produce antibodies. However, we observed that all the sera from this group of subjects had antibodies to community CoV and; furthermore, sera from 2 patients that had tested negative before the COVID-19 pandemic did develop antibodies to all 5 SARS-CoV-2 at a time infections were surging. With regard to sera from confirmed positive cases, we were unable to collate sample date with disease status (eg, acute versus convalescent). A final limitation to the multiplex assay (and all other antibody detection assays) is that it does not determine whether any of the detected antibodies have neutralizing capacity. Future studies will address this issue.
In conclusion, we developed a multiplex, high-throughput, sensitive, and specific assay that should provide a more comprehensive and in-depth understanding of the dynamics, biology, and repertoire of antibody responses to SARS-CoV-2. From a practical standpoint, the SARS-CoV-2 multiplex assay described here is very similar to the multiplex assays used by histocompatibility laboratories around the world to identify HLA antibodies. As such, incorporating the SARS-CoV-2 assay into their routine testing of transplant candidates and recipients would be seamless and easily adapted by HLA laboratories
The authors would like to acknowledge Drs. Sean Stowell, Scott Boyd, and Zahra Kashi for testing and/or providing COVID-19 positive samples. The authors also thank Ms. Shilpee Biwas for expert technical assistance and Dr Tina Meng for Spike protein preparations. In addition, the authors acknowledge Drs. Medhat Askar and Phillip Ruiz for helpful discussions regarding assay development.
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