Nucleocapsid Antibodies as an Optimal Serological Marker of SARS‐CoV‐2 Infection: A Longitudinal Study at the Thomayer University Hospital

ABSTRACT

Background

The longitudinal study was conducted over the initial 2 years of the COVID‐19 pandemic, spanning from June 2020 to December 2022, in healthcare workers (HCWs) of the Thomayer University Hospital. A total of 3892 blood samples were collected and analyzed for total nucleocapsid (N) antibodies. The aim of the study was to evaluate the dynamics of N antibodies, their relationship to the PCR test, spike (S) antibodies, interferon‐gamma, and prediction of reinfection with SARS‐CoV‐2.

Methods

Blood collections were performed in three rounds, along with questionnaires addressing clinical symptoms of past infection, PCR testing, and vaccination. Antibody measurements included total N antibodies (Roche Diagnostics) and postvaccination S antibodies (Euroimmun). Cellular immunity was tested by interferon‐gamma release assay (Euroimmun).

Results

At the end of the study, 35.9% of HCWs were positive for N antibodies, and 39.5% of HCWs had either known PCR positivity or N antibodies or both. Ten percent of participants had no knowledge of a COVID‐19 infection and 35% of positive individuals exhibited no symptoms. The values of positive antibodies decrease over a period of 6 months to 1 year, depending on the initial value, and their dynamics are highly variable. The study also demonstrated that the highest levels of spike antibodies and interferon‐gamma occur during so‐called hybrid immunity.

Conclusion

Nucleocapsid antibodies proved valuable in monitoring SARS‐CoV‐2 infection dynamics, and they may detect cases of SARS‐CoV‐2 infection missed by PCR tests. The study identified distinct patterns in antibody dynamics and protection of hybrid immunity during reinfection.

The study reveals the diagnostic significance of nucleocapsid antibodies and the strategy for testing past COVID‐19 infection in the posvaccination era. The results demonstrate that hybrid immunity acquired through infection and vaccination is the most effective protection against further reinfection of SARS‐CoV‐2.

1. Introduction

Severe acute respiratory syndrome coronavirus 2 (SARS‐CoV‐2) is a positive single‐stranded RNA virus with four structural components: the spike (S), nucleocapsid (N), envelope (E), and membrane (M) proteins. These components are vital for the virus's structure, function, and interaction with host cells. The receptor‐binding domain in the S1 subunit of the spike protein mediates specific cell entry and viral fusion by binding to human angiotensin converting enzyme 2 [1]. The nucleocapsid (N) protein is crucial for the transcription and replication of SARS‐CoV‐2, binding and packaging viral RNA into a ribonucleoprotein complex [2]. It also modulates host immune responses and influences viral pathogenesis. Zhao et al. [3] reported that the N protein has a dual role in innate immune responses, i.e., low doses suppress, while high doses promote type I interferon signaling and inflammatory cytokines.

Among the SARS‐CoV‐2 antigens, the N protein has high immunogenicity and is prevalently present in viral particles [4]. Detecting this protein in patient blood samples indicates acute viral infection [5]. Serological testing is critical for diagnosing and monitoring the virus's spread, with specific guidelines necessary for accurate results. To confirm past infection, determining antibodies against nucleocapsid is preferable, while for vaccination response confirmation, a test demonstrating antibodies against spike protein is necessary [6]. Although both proteins can accumulate mutations, the spike protein, as the primary target of host immune responses, tends to accumulate more mutations, making it a less stable diagnostic tool. Mutations in the nucleocapsid protein do not affect diagnostic test performance [7]. The selection of a specific immunoglobulin isotype for N antibodies assays lacks sufficient evidence, making total N antibodies preferable for monitoring of past infections [8]. Unlike the recommended nasopharyngeal swab for PCR in acute infection, venous blood collection for the determination of postacute antibodies is easy, standardized, and offers higher sample stability.

Advanced laboratory techniques, including cellular assays, have been implemented for COVID‐19 diagnostics. The interferon‐gamma (IFN‐gamma) T‐cell release assay detects a response to the S protein, which is generally low in the acute phase of infection but increase over time [9].

The study involved healthcare professionals from the Thomayer University Hospital in Prague, which faced extraordinary challenges during the COVID‐19 pandemic, adapting its operations to manage patient influx, control virus spread, and to protect staff. The article aims to provide a comprehensive analysis of the effectiveness of nucleocapsid antibodies as a serological marker within the broader context of diagnostic methods. Measurement of postvaccination antibodies and cellular immunity was included as a secondary aim of the study immediately after their commercial availability.

2. Materials and Methods

2.1. Study Period: The COVID‐19 Pandemic in the Czech Republic

The study spanned two and a half years, from June 2020 to December 2022, covering three waves of the COVID‐19 pandemic in the Czech Republic: the original Wuhan, Alpha, Delta, and Omicron variants. The first cases appeared in March 2020, vaccinations began in December 2020, and mandatory testing ended in February 2022. By late 2022, the Czech Republic recorded over 4.5 million infections and 41,000 deaths. More than 5 million people were fully vaccinated, and 1.8 million received partial vaccination among the country's population of over 10 million inhabitants [10].

2.2. Study Design and Participation

Healthcare workers (HCWs) from the Thomayer University Hospital participated in the study. Each participant received information about the study's purpose, signed an informed consent, completed a questionnaire on demographics, previous PCR test results, clinical symptoms, vaccination history, and underwent blood collection. The study comprised several phases: the first round of sampling occurred at the onset of the COVID‐19 pandemic (June 30, 2020 to September 25, 2020), followed by a second round 3 months later (December 11, 2020 to February 8, 2021), after the initial peak of the pandemic in the Czech Republic. A third round was conducted 7 months postvaccination (September 13, 2021 to November 12, 2021). Subsequently, a year‐long observational study continued (November 13, 2021 to December 31, 2022). Collaboration with the Institute of Health Information and Statistics (IHIS) of the Czech Republic ensured the accurate assignment of all new infection confirmed by PCR to participants for 2022.

2.3. PCR Testing

PCR testing was not part of this study but was considered the primary standard diagnostic method for confirming acute COVID‐19 infection. At the beginning and culmination of the pandemic, testing of all HCWs by PCR was not possible for ethical, economic, and logistical reasons. The Thomayer University Hospital had implemented PCR testing rules since the pandemic's onset. All symptomatic HCWs and all those who had been in contact with a PCR positive person were tested. All the others were assumed negative and there was no reason to test them. The PCR results were recorded using personal questionnaires and verified in the IHIS database.

2.4. Detection of Antibodies and Interferon‐Gamma

Blood samples were analyzed for antibodies and IFN‐gamma. The semiquantitative measurement of total N protein antibodies was determined by electrochemiluminescence Elecsys anti‐SARS‐CoV‐2 assay on a Cobas e602 analyzer system (all from Roche Diagnostics, Rotkreuz, CH), following the manufacturer's instructions. A positive response was defined by a cutoff index (COI) of > 1, with a sensitivity of 97.2% (95.4%–98.8%) and a specificity of 99.8% (99.3%–100%) [11].

In the third round, all HCWs were tested for postvaccination S antibodies and IFN‐gamma as a T‐lymphocyte response to in vitro S protein stimulation. Spike IgG levels were measured using the anti‐SARS‐CoV‐2 QuantiVac ELISA IgG (Euroimmun, Lübeck, DE), with a positive response defined by a cutoff of 25.6 BAU/mL, showing a sensitivity of 100% (96.8%–100%) and a specificity of 98.3% (95.0%–99.6%) [12]. IFN‐gamma levels were determined using the Quant‐T‐cell SARS‐CoV‐2 stimulation tube and Quant‐T‐cell ELISA (Euroimmun, Lübeck, DE), with a positive response defined by a cutoff of 100 IU/L, with a sensitivity of 89.4% (81.5%–94.3%) and specificity of 84.6% (66.5%–93.8%) [13]. Samples not meeting the criteria for negative (background) and positive (stimulation) controls could not be evaluated, but the number of invalid results was reported.

2.5. Statistical Analysis

Univariate analysis was performed on measurement characteristics for positive or negative outcomes, with categorical values presented as numbers and percentages and continuous values as medians and interquartile ranges. Regression analysis and Pearson's correlation coefficient (r) were used to estimate the relationship between days after a positive PCR and N antibody values. A chi‐square test for independence was used to determine if differences between the groups of HCWs with new infections were significant. The statistical analyses and plots were generated using MS Excel.

2.6. Ethics Statement

The study received approval from the Ethics Committee of the Institute for Clinical and Experimental Medicine and the Thomayer University Hospital in accordance with the Declaration of Helsinki (code G‐20‐60) on June 18, 2020. Participation in the study was voluntary. Adhering to guidelines for information provision, all participants provided written informed consent. Handling of all personal data obtained from IHIS has been in accordance with the General Data Protection Regulation (Regulation EU).

3. Results

3.1. The Progression of the COID‐19 Pandemic at the Thomayer University Hospital

The study, conducted during the first 2 years of the COVID‐19 pandemic at the Thomayer University Hospital in Prague, faced an exceptional workload with increased hospitalizations for moderate and severe COVID‐19. All HCWs with confirmed PCR positive results had to isolate. By the end of 2021, the hospital had 3600 hospitalized patients and 670 positive HCWs, accounting for over a quarter of the staff (Figure 1).

FIGURE 1.

Cumulative number of hospitalized patients with COVID‐19 and healthcare workers (HCWs) with COVID‐19 confirmed by PCR at the Thomayer University Hospital. The columns represent the chronological setting of the study.

The Thomayer University Hospital has around 2500 employees, only a part of them participated in the voluntary three‐round testing. However, this number substantially increased over time, reaching a total of 1703 participants in the third round. The initial two rounds occurred before vaccination, involving 808 and 1381 HCWs, respectively. The baseline characteristics of the participants indicated that more than 80% were women aged 30–64, working as healthcare professionals (Table 1).

TABLE 1.

Demographic characteristics of the study and participants.

	1st round	2nd round	3rd round
Date	Jun 30—Sep 25, 2020	Dec 11—Feb 8, 2021	Sep 13–Nov 12, 2021
Participants	808	1381	1703
Age, years
18–29	74 (9.2%)	187 (13.5%)	168 (9.9%)
30–64	681 (84.3%)	1109 (80.3%)	1405 (82.5%)
> 65	53 (6.5%)	85 (6.2%)	130 (7.6%)
Gender
Female	713 (88.2%)	1153 (83.5%)	1418 (83.3%)
Male	95 (11.8%)	228 (16.5%)	285 (16.7%)
Profession
Healthcare professionals	600 (74.3%)	1114 (80.7%)	1458 (85.6%)
Support staff	208 (25.7%)	267 (19.3%)	245 (14.4%)
Vaccination	—	—	1578 (92.7%)

3.2. Comparison of Nucleocapsid Antibodies Testing With Official PCR Testing

One of the aims of the study was to determine and document the prevalence of N antibodies in HCWs during the acute spread of the SARS‐CoV‐2 virus. The results were subsequently compared with available information from positive PCR tests, as described in the Section 2. Antibody tests were performed independently at a different time than individual PCR tests. In the first round, the range was 13–144 days after PCR tests, with a median of 96 days. In the second round, the range was 9–311 days after PCR tests, with a median of 63 days. In the third round, the range was 7–574 days after PCR tests, with a median of 262 days.

The number of HCWs testing positive for PCR and/or N antibodies increased consistently across the testing rounds, indicating a growing number of participants with active infection developing an immune response to the N protein. In the first round, relatively few HCWs were infected, as demonstrated by both PCR and N antibody positivity (1.7% and 1.9%). However, results in the second, resp. the third rounds uncovered a slightly higher number of tests with N antibody positivity (28.5%, resp. 35.9%) compared to PCR positivity (21.7%, resp. 29.4%) as shown in Figure 2.

FIGURE 2.

The number and percentage of HCWs in each round with negative and positive results for PCR and/or N protein antibodies out of the total HCWs.

There is only partial agreement between known PCR positivity and the presence of N antibodies, influenced by various factors with diagnostic implications. Figure 3 illustrates the distribution of positive results between PCR and N antibody tests in each round (Figure 3A). Most HCWs showed positive results in both PCR and N antibody tests. However, some were positive only in PCR or only in N antibodies. Combining 500 HCWs with positive PCR results (61 PCR pos. + 439 PCR/N Ab pos.) and 173 HCWs with positive N antibodies only, 39.5% of all tested HCWs were positive in the third round (673 out of 1703) (Figure 3A).

FIGURE 3.

Confirmation of past COVID‐19 infection using nucleocapsid antibodies and their comparison with available PCR tests: (A) number of positive results per round; (B) proportion of positive results from all rounds using different cutoff values to evaluate antibody positivity (CO ≥ 1.000 and CO ≥ 0.135).

Adjusting the N antibody positivity cutoff from the manufacturer's recommended 1.00 to our optimized 0.135 [14] increased test agreement from 63.8% to 67.9% of all positive tests. Only 4.8% of positive PCR results were found without accompanying positive N antibodies. The N antibody test identified 27.3% of HCWs among all positives who did not have a positive PCR test during the tested period (Figure 3B). Half of these individuals had negative PCR results, and the other half were untested.

Questionnaires assessed whether participants had experienced COVID‐19‐like respiratory symptoms (temperature over 37°C, sore throat, cough, difficulty breathing, loss of taste or smell) in the past year. Symptoms were compared with positive PCR and nucleocapsid antibody results (Figure 4). The highest proportion of symptomatic cases (47.7%) occurred when both tests were positive, while 16.2% remained asymptomatic. In cases where only one test was positive, the ratio of symptomatic to asymptomatic cases was balanced. Notably, 35.2% of individuals identified as positive by either of these methods showed no symptoms, highlighting significant asymptomatic cases.

FIGURE 4.

Percentage of symptomatic and asymptomatic HCWs with nucleocapsid antibodies only, PCR positivity only, or both.

3.3. Values and Dynamics of Nucleocapsid Antibodies

In the first round, only 15 HCWs exhibited positive N antibodies with very high values and a median COI of 74.88. Over time, the maximum antibody values increased; however, the median decreased on 42.95 COI with a higher number of positive samples (Table 2). This discrepancy emerged with the increase in new infections characterized by low antibody values, which depend on the time interval between exposure and testing.

TABLE 2.

Statistical data on positive nucleocapsid antibody values (cutoff index) by rounds.

Positive nucleocapsid Abs (≥ 1)
	1st round	2nd round	3rd round
HCWs (no.)	15	394	612
Min (COI)	5.80	1.01	1.00
Max (COI)	107.50	209.70	250.00
Median (COI)	74.88	42.95	18.40

HCWs underwent PCR testing when suspected of COVID‐19 or after contact with a confirmed positive individual. All PCR tests were conducted independently of this study during the pandemic. Analysis of the questionnaires revealed that 814 HCWs had tested positive for PCR before blood collection for antibody assessment. When examining the correlation between N antibody values and the time since the positive PCR test, no significant correlation was identified. The regression curve in Figure 5 demonstrates a gradual decrease and Pearson's correlation coefficient of −0.150 indicates a weak negative correlation between the two variables. The antibody responses to SARS‐CoV‐2 infections, measured as levels of N antibodies, were highly variable.

FIGURE 5.

Correlation between nucleocapsid antibody values and number of days after positive PCR test.

Nucleocapsid antibody dynamics were observed in 541 HCWs who participated in all three rounds of blood collection. Blood samples were obtained 3–7 months (2nd round) and 12–16 months (3rd round) after the initial collection. Throughout the follow‐up period, 353 HCWs remained negative, while 188 HCWs tested positive. Seven HCWs had positive antibodies from the 1st round, 132 HCWs from the 2nd round, and 49 HCWs from the 3rd round. The individual dynamics of antibody values were tracked over time.

Figure 6A shows those who have had positive N antibodies since the start of testing. The positive antibody values measured in the first round exhibited a slight decline and remained positive for another year due to the original Wuhan variant of SARS‐CoV‐2. Among these cases, we documented one reinfection between the second and third rounds, resulting in a sharp increase in antibody values. Figure 6B shows HCWs who tested negative in the first round, but had measurable N antibodies after the infection of Wuhan and Alpha variants in the second round. Only 10 out of 132 individuals with low‐positivity antibodies experienced a decline to just below the value of 1.0 COI within 6 months, while others maintained the same antibody values, and some exhibited a continued rise. In general, it can be concluded that N antibodies persist for at least 6 months to a year.

FIGURE 6.

Monitoring the dynamics of nucleocapsid antibodies in individual HCWs over time: (A) positivity at the beginning (1st round) (n = 7) (B) positivity after 3–7 months from the beginning of the study (2nd round) (n = 132).

3.4. The Effect of Nucleocapsid Antibodies as a Marker of Infection and Vaccination on Spike Antibody Levels

In the third round of the study, 1578 HCWs (92.7% of all participants) received two doses of Comirnaty (Pfizer, BioNTech), administered in the range of 7–273 days (median: 229 days) before the final blood collection. Since postvaccination are exclusively produced S antibodies, the diagnostic approach for past infection, involving the detection of N antibodies, remains consistent. S antibodies are generated in response to both virus infection and vaccination. Therefore, in the third round of the study, measurements were taken for both N and S antibodies. Of all participants, 1627 HCWs (95.5%) tested positive for S antibodies, including both vaccinated individuals and those previously infected.

The results were categorized into four groups based on vaccination status and COVID‐19 infection. Samples from HCWs who had experienced previous infection showed the lowest median S antibody value at 79.0 BAU/mL. Those vaccinated without prior infection had a median value of 142.5 BAU/mL, which was twice as high as the COVID‐19‐infected group. Significantly higher levels, around 500 BAU/mL, were observed in individuals who experienced both infection and vaccination (Figure 7).

FIGURE 7.

Levels of IgG antibodies against spike protein in four different HCWs groups: Unvaccinated with negative nucleocapsid antibodies, unvaccinated with positive nucleocapsid antibodies, vaccinated with negative nucleocapsid antibodies, and vaccinated with positive nucleocapsid antibodies.

3.5. The Effect of Nucleocapsid Antibodies as a Marker of Infection and Vaccination on IFN‐Gamma Levels

Cellular immunity was assessed by measuring IFN‐gamma production following blood stimulation with the S protein in vitro. Positive cellular responses were noted postinfection and postvaccination. Sixteen participants were excluded due to low cell responses to a non‐specific positive control, and 18 due to high basal IFN‐gamma levels (negative control). Among the remaining 1569 HCWs, 1510 (96.2%) exhibited a positive cellular response.

A distribution of results very similar to S antibodies was observed when HCWs were categorized into four groups based on N antibody positivity and vaccination status. IFN‐gamma levels were slightly lower postinfection but higher postvaccination. Following combined stimulation from natural infection and vaccination, IFN‐gamma levels peaked at 1500 IU/L (Figure 8).

FIGURE 8.

Levels of interferon‐gamma in four different HCWs groups: Unvaccinated with negative nucleocapsid antibodies, unvaccinated with positive nucleocapsid antibodies, vaccinated with negative nucleocapsid antibodies, and vaccinated with positive nucleocapsid antibodies (IFN‐gamma levels could not be determined at 34 HCWs due to non‐compliance with negative or positive control criteria).

3.6. New Infection and Reinfection During the Following Year

One year after the last blood collection, we retrospectively analyzed new COVID‐19 infections and reinfections among participants. A positive PCR test between January and December 2022 indicated a new infection with the Omicron variant. Among unvaccinated HCWs without N antibodies, 56% had new infections (Table 3). Conversely, the vaccinated group previously recovered from COVID‐19 infection had the lowest reinfection rate at 18.6%. This group showed high postvaccination antibody levels and robust cellular immunity at the study's end.

TABLE 3.

HCWs with new COVID‐19 infection over the following year.

Group	Total HCWs (n)	New infections (Observed)	New infections (%)	Chi‐square test statistic	p
Negative HCWs	50	28	56.0
HCWs with COVID‐19	84	33	39.3
Vaccinated HCWs	1041	427	41.0
HCWs with COVID‐19 + vaccination	528	98	18.6
Total	1703	586		90.12	2.06 × 10−19

The p‐value is extremely low (much < 0.05), indicating that the differences between the groups are highly significant.

4. Discussion

The COVID‐19 pandemic, caused by SARS‐CoV‐2, has led to extensive research on the virus's immune responses and related laboratory diagnostics. The nucleocapsid protein is a crucial component of SARS‐CoV‐2, eliciting a significant humoral immune response, which confirms its potential as an important serological marker for infection [15].

The Roche Elecsys immunoassay tuned for the detection of high avidity antibodies is most capable of accurately documenting past SARS‐CoV‐2 infections [16]. Our study, spanning two and a half years and covering the Wuhan, Alpha, and Delta variants, confirmed that total nucleocapsid antibodies measured by electrochemiluminescence are an ideal marker for longitudinally monitoring SARS‐CoV‐2 infection. Results showed a cumulative increase in participants developing an immune response against the N protein across three rounds (1.9%, 28.5%, and 35.9% of all HCWs). Similar studies reported a large variation in positive N antibody percentages among HCWs, ranging from 8.4% to 34.3% [17, 18, 19]. This variability could be due to differences in study design, geographical location, and preventive measures during follow‐up periods.

The study highlights a partial correlation between officially performed PCR tests and nucleocapsid antibody positivity, emphasizing the diagnostic significance of this phenomenon. Nucleocapsid antibodies can identify SARS‐CoV‐2 infections that PCR tests might miss. HCWs with positive N antibodies and false‐negative PCR results could be due to low viral load, timing, or sample collection quality [20]. HCWs with positive PCR and negative nucleocapsid antibodies were rare. Evidence suggests a genetic basis (HLA‐B*15:01) in asymptomatic SARS‐CoV‐2 infection, indicating situations where PCR is positive before antibody production, and some individuals might clear subclinical infection before seroconversion [21]. It has been published that asymptomatic cases might produce only spike IgG antibodies but not nucleocapsid IgG antibodies [22]. With many people vaccinated and possessing positive S antibodies, assessing total N antibodies is most advantageous, capturing a broader spectrum of asymptomatic individuals than IgG subclass‐specific N antibodies.

The study tracked N antibody values over three rounds, showing an increase in maximum values and a decrease in median values over time. This discrepancy was due to a higher number of recent infections with initially low antibody levels and fewer older or repeated infections. No significant correlation was found between N antibody values and time since the positive PCR test, indicating highly individualized antibody responses. N antibodies persisted for at least 6 months to a year, highlighting their diagnostic value for past infections. Other studies confirm these dynamics, showing antibody waning from 120 days onward [23] and persistence for up to 18 months postinfection [24].

The study included 92.7% of participants who had received two doses of the Comirnaty vaccine about 6 months before the final blood collection. Spike antibody and IFN‐gamma levels increased both after infection and vaccination, reaching the highest values with hybrid immunity. The synergy between antibodies and the cellular immune response highlights the effectiveness of hybrid immunity in providing superior protection compared to immunity from infection or vaccination alone. Additionally, memory T cells from previous coronavirus infections produce IFN‐gamma, playing an important role in rapid virus removal from the body [25].

One year after the last blood collection, reinfections were monitored, revealing higher rates of new infections in unvaccinated individuals with negative nucleocapsid antibodies. Furthermore, it was demonstrated that hybrid immunity provides robust protection against reinfection. Studies on the Omicron variant reported the long‐term effectiveness of hybrid immunity, particularly in protecting against severe disease [26]. However, complete protection against symptomatic infection is limited, and waning immunity is a reality with current vaccines [27].

Given the lingering health consequences of COVID‐19 in some individuals, SARS‐CoV‐2 remains a significant viral pathogen. Although people without hybrid immunity might be more likely to contract COVID‐19, those with hybrid immunity are not immune to the disease. Therefore, everyone, regardless of their COVID‐19 history, should take steps to minimize their chances of SARS‐CoV‐2 infection, such as staying up to date with vaccinations.

5. Conclusions

In conclusion, this study offers a thorough insight into nucleocapsid antibodies in SARS‐CoV‐2 infection. It underscores their crucial diagnostic value, dynamics over time, and correlation with vaccination and cellular immunity. These findings are pivotal for developing effective diagnostic and monitoring approaches for COVID‐19 infection.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.