Long-lasting T-cell response to SARS-CoV-2 antigens after vaccination—a prospective cohort study of HCWs working with COVID-19 patients

Abstract

Background

Vaccination against SARS-CoV-2 reduces the risk of hospitalisation and death, but vaccine-induced IgG antibodies against the spike protein (IgG S) decline over time. Less is known about the nature of the vaccine-induced T-cell response to SARS-CoV-2 antigens.

Methods

IgG antibodies against nucleocapsid protein (IgG N), IgG S, and T-cell response towards SARS-CoV-2 antigens were determined in samples taken between November 2020 and November 2021 from a cohort of healthcare workers at an Infectious Diseases Department. RT-PCR screening for SARS-CoV-2 was encouraged once every four weeks in addition to testing when symptomatic or identified through contact tracing. Vaccination data were collected at the end of the study.

Results

At inclusion, T-cell response to SARS-CoV-2 antigens was found in 10/15 (66.7%) of participants with a previous/current COVID-19 infection and in 9/54 (16.7%) of participants with no prior/current history of COVID-19 infection. All participants with complete follow-up (n = 59) received two doses of a SARS-CoV-2 vaccine during the study. All participants demonstrated detectable IgG (S) antibodies at the end of the study, in median 278 days (IQR 112) after the second vaccine dose. All but four participants displayed T-cell responses towards SARS-CoV-2 antigens. IgG S antibody levels correlated with time since the second vaccine dose. In addition, previous COVID-19 infection and the strength of the S1 T-cell response correlated with IgG S antibody levels. However, no correlation was demonstrated between the strength of the T-cell response and time since the second vaccine dose.

Conclusion

COVID-19 vaccination induces robust T-cell responses that remain for at least nine months.

Keywords:

• SARS-CoV-2
• COVID-19
• IgG
• T-cell response
• HCWs

Introduction

In less than a year from the start of the COVID-19 pandemic, highly effective vaccines have been developed that substantially reduce the risk of hospitalisation and death. The emergence of SARS-CoV-2 variants that can partially evade vaccine-induced immune responses combined with waning antibody titres has raised concerns about the persistence of humoral and cellular immune responses over time. Natural infection or vaccination with mRNA or adenoviral vectors elicit antibodies against the SARS-CoV-2 spike protein and SARS-CoV-2–specific T-cell responses. The spike-antibody titre has been reported to be correlated with the risk of symptomatic infection [1]. However, the antigenic drift in the omicron variant has enabled the virus to spread rapidly, even in the vaccinated population [2]. Also, the kinetics of vaccine-induced antibodies compared to natural infection antibodies appear different. An Israeli study reported a monthly decline of up to 38% of vaccine-induced antibodies compared to a <5% decline per month after infection [3]. A growing body of evidence indicates that T-cells may protect against SARS-CoV-2 infection. A recently published study reported a higher frequency of cross-reactive and nucleocapsid-specific IL-2-secreting memory T-cells among uninfected SARS-CoV-2 exposed individuals than infected exposed individuals [4]. A second-generation SARS-CoV-2 T-cell vaccine that triggers both CD4⁺ and CD8⁺ T-cell responses well exceeding that of prior vaccine technologies is undergoing clinical trials [5].

New variants of concern have emerged every few months and rapidly spread worldwide. There is a need for complementary non-spike targets for variants that evade vaccine-induced spike antibodies to assess immunity. Nevertheless, studies on the long-term persistence of T cell responses after infection, vaccination, or a combination are lacking. The individual’s history of prior infections, type of vaccine, and the interval between doses may be crucial factors for a long-term T-cell response [6].

Healthcare workers (HCWs) are at the frontline of the pandemic and consistently risk exposure to SARS-CoV-2 regardless of the overall community transmission. To ensure adequate protection against infection, assessment of T-cell responses and individualised booster doses or T-cell vaccines may be part of future vaccine strategies.

We performed a prospective cohort study of HCWs at an Infectious Diseases Department caring for COVID-19 patients from November 2020 to November 2021, i.e. during the second and third wave of COVID-19 in Sweden. SARS-CoV-2 IgG antibodies against the spike protein (IgG S) and the nucleocapsid protein (IgG N) and T-cell responses to SARS-CoV-2 peptides were assessed, and the incidence of COVID-19 was recorded. The study was not designed to evaluate vaccine efficacy but is a pragmatic investigation of the antibody, and T-cell responses to SARS-CoV-2 antigens in a cohort of HCWs vaccinated as per the national policy.

Methods

Ethics statement

The study was approved by the Institutional Review Board in Göteborg, Sweden (Dnr 2020-04674). Written informed consent was obtained from all participants.

Study design and setting

All personnel working at the Department of Infectious Diseases (ID), Västmanlands hospital Västerås, Sweden, in November 2020, were eligible to be included in the study. Information about the study was given at workplace meetings and through posters and emails. During the study, the region Västmanland experienced the second (November 2020–January 2021) and third wave (March 2021–May 2021) of COVID-19.

At the beginning of the study in late November 2020, participants completed a short questionnaire (Supplementary Material), and blood samples were obtained for serology and analysis of T-cell responses towards SARS-CoV-2 antigens. Follow-up blood samples were obtained 12 months later in November 2021. Participants were encouraged to provide a combined gargle and throat swab sample for SARS-CoV-2 screening by RT-PCR every four weeks. In addition, RT-PCR testing was required by the employer when experiencing symptoms compatible with COVID-19 or identified through contact tracing.

Vaccination was not part of the study protocol and inclusion into the study did not influence the type or timing of the vaccination. By national policy, HCWs working in close contact with COVID-19 patients were offered vaccination with BNT162b2 in January 2021 with a three-week interval between doses. Personnel without close patient contact were offered either BNT162b2, mRNA-1273, or ChAdOx1 starting in April 2021, adhering to the priority rules of the general Swedish population. The vaccination dates, but not the type of vaccine, were obtained from the participants’ medical records at the end of the study.

All data was pseudo-anonymized, and the identifying key was only accessible to the authors and study nurses.

Laboratory assays

IgG antibodies against the SARS-CoV-2 nucleocapsid

A chemiluminescent microparticle immunoassay (CMIA) (IgG assay, Abbott Core Laboratory Systems, Lake Forest, IL, USA) was used on the Architect i2000SR Plus instrument for the qualitative detection of IgG N antibodies following the manufacturer’s instruction. Sample/calibrator (S/C) relative light unit (RLU) index <0.8 was regarded as negative, S/C index 0.8–<1.4 as indeterminate, and S/C index ≥1.4 as positive.

IgG antibodies against the receptor-binding domain (RBD) of the S1 subunit of the spike protein of SARS-CoV-2

For detecting IgG antibodies against the spike protein, a CMIA (AdviseDX SARS-CoV-2 IgG II assay, Abbott Core Laboratory Systems, Lake Forest, IL, USA) was used on the Architect i2000SR Plus instrument following the manufacturer’s instructions. Zero to seven binding antibody units (BAU)/mL was considered negative, >7 BAU/mL but <14 BAU/mL indeterminate, and ≥14 BAU/mL positive. Samples collected in November 2020 were stored at −70 °C for later analysis.

The T-cell response towards SARS-CoV-2 antigens

PBMC was prepared using density gradient separation according to the manufacturer’s instructions, either using BD Vacutainer CPT tubes (Becton, Dickinson and Company, Franklin Lakes NJ, USA) or pluriMate II 15 ml tubes filled with PBMC Spin media (pluriSelect Life Science, Leipzig, Germany). PBMC was washed with phosphate-buffered saline (PBS) and resuspended in serum-free AIM-V media (ThermoFisher Scientific, MA, USA) at 2.5 × 10⁶ cells per ml. The reactivity of the PBMC samples to a SARS-CoV-2 S1 (spike) scanning peptide pool and a defined peptide pool containing epitopes from the spike (S), nucleocapsid (N), membrane (M), and open reading frame (O) proteins of the virus was analysed at ABC Labs (Solna, Sweden) with the ELISpot Confirm kit: SARS-CoV-2 (T-cell, IFN-γ) (Mabtech AB, Stockholm, Sweden, exclusively licenced to ABC Labs in the Nordic countries). This ELISpot kit is similar to the commercially available ELISpot Path: Human IFN-γ (SARS-CoV-2, S1scan + SNMO) (Mabtech AB, Stockholm, Sweden). The assay was set up according to the manufacturer’s instructions. Briefly, PBMC was stimulated in IFN-γ capture ELISpot plates with (1) media + anti-CD28 antibody (control), (2) S1 peptide pool + anti-CD28 antibody, (3) SNMO peptide pool + anti-CD28 antibody or (4) anti-CD3 antibody + anti-CD28 antibody (positive control) for 18 h (37 °C, 5% CO₂). The cells were washed, and the plate was incubated for 2 h with a secondary anti-IFN-γ antibody conjugated to ALP. The plate was washed, and spots were developed with BCIP/NBT-plus substrate for 10–15 min. Plates were washed and dried, and spots were analysed with a Mabtech ASTOR ELISpot reader (Mabtech AB, Stockholm, Sweden). Criteria for a positive T-cell response were set in relation to the negative control (media). A well with seven or more spot-forming units (SFU) after deduction of the number of spots in the negative control well (background), and with more than two times the number of SFU in the negative control well was regarded as positive. A normalisation method to the positive control (anti-CD3 antibody) was applied to account for differences in sampling quality and enable comparison of T-cell response between individuals and between T-cell response and antibody levels. The sample SFU/well was divided by the SFU/well for the positive control times 1000. Wells that did not meet the criteria for a positive response were set at 0 SFU/well.

Detection of SARS-CoV-2 in gargle water/throat swab samples by real-time reverse transcription-polymerase chain reaction (RT-PCR)

The combination of gargle water and throat swab was chosen to enable self-sampling. Before implementation, the combined sample was evaluated by the Department of Laboratory Medicine, Västmanlands hospital Västerås, and the performance was deemed equivalent to nasopharyngeal swabs. Participants were provided with a barcode label to mark the sample. After analysis, participants could retrieve the RT-PCR results from their medical records. Following regional policy, all positive RT-PCR results were conveyed to the COVID-19 contact tracing team for further action. Participants were reminded by email to provide monthly samples. At the end of the study, the results of all RT-PCR tests provided by each participant (whether taken as part of the study or due to symptoms or contact tracing) were obtained from the laboratory linked to the participants’ study identification number.

All RT-PCR samples were analysed in the clinical workflow at the Department of Laboratory Medicine, Västmanlands hospital Västerås, Sweden. During the study period, several different RT-PCR methods were used in parallel to cope with the significant number of samples. Most samples were analysed with Alinity m Resp-4-Plex assay (Abbott Core Laboratory Systems, Lake Forest, IL, USA), but Allplex 2019-nCoV assay (Seegene, Seoul, South Korea) after extraction with GXT96 X3 Extraction Kit (Hain Lifesience, Nehren, Germany), Allplex SARS-CoV-2/FluA/FluB/RSV Assay (Seegene, Seoul, South Korea) after extraction with GXT96 X3 Extraction Kit (Hain Lifesience, Nehren, Germany), Abbott RealTime SARS-CoV-2 Assay (Abbott Core Laboratory Systems, Lake Forest, IL, USA), Alinity m SARS CoV-2 (Abbott Core Laboratory Systems, Lake Forest, IL, USA), Xpert Xpress SARS-CoV-2 (Cepheid, Sunnyvale, CA, USA), and Xpert^® Xpress SARS-CoV-2/Flu/RSV (Cepheid, Sunnyvale, CA, USA) were also used during the study period.

Statistics

Jamovi version 2.2.5.0 (https://www.jamovi.org.) and STATA version 16.1 (StataCorp, TX, USA) were used for the statistical analyses. Continuous data were assessed for normality. Non-parametric statistical analyses were performed using one-way ANOVA (Kruskal-Wallis) or repeated-measures ANOVA. Mann–Whitney U-test was used to test for association between the magnitude of the T-cell response to S1 antigens and previous COVID-19 at the study end.

Spearman’s rank correlation was used to test for association between normalised S1 T-cell response and IgG S levels at the study end.

Results

The participants’ characteristics are presented in Table 1. Among participants with a previous or current COVID-19 infection at the study start (as reported by the participant or based on RT-PCR at inclusion) (n = 15), IgG N was confirmed in 3/15 (20.0%), IgG S in 10/15 (66.7%) and T-cell response to either S1 or SNMO in 14/15 (93.3%) (Table 2). In contrast, among study participants with no previous confirmed COVID-19 infection (n = 54), 1/54 (1.9%) was IgG N positive, 2/54 (3.7%) were positive for IgG S and T-cell response to either S1 or SNMO was found in 10/54 (18.5%) of participants (Table 2). A T-cell response to S1 and/or SNMO among study participants with no previous confirmed SARS-CoV-2 infection was not associated with age (p = 0.973), sex (p = 0.305), years of work experience in ID (p = 0.798), years of work experience in health care (p = 0.755), reported frequency of household member with COVID-19 (p = 0.501), nor the reported frequency of having being identified as a close-contact within contact tracing for COVID-19 (p = 0.501). Fifty-six participants (five with previous COVID-19, and 51 with no previous COVID-19) were seronegative (no IgG N nor IgG S) at the study-start. Twelve seronegative individuals demonstrated T cell responses to S1 and/or SNMO antigens.

Table 1. Characteristics of study participants.

Characteristic	All participants (n = 69)	Complete follow-up (n = 59)
Age, median (IQR), years	39 (12.2)	40 (12.2)
Sex, female, n (%)	57 (82.6)	48 (81.4)
Occupation, n (%)
Nurse	21 (30.4)	17 (28.8)
Assistant nurse	13 (18.8)	10 (16.9)
Doctor	18 (21.6)	18 (30.5)
Other	17 (24.6)	14 (23.7)
Work experience years, median (IQR)
In infectious diseases	4 (11.3)	7 (13.5)
In health care	10 (17.0)	12 (20.5)
Previous confirmed COVID-19, n (%)
By RT-PCR	13 (18.8)	11 (18.6)
By serology	9 (13)	9 (15.3)
Exposure to COVID-19 outside work, n (%)
Household member with confirmed infection	6 (8.7)	6 (10.2)
Identified as a close contact through contact tracing	4 (5.8)	4 (6.8)

IQR: interquartile range; n: number.

Table 2. Prevalence of IgG (N), IgG (S), and T-cell response (S1 and SNMO) at study start (n = 69).

	Prior or current COVID-19 (n = 15)	No previous or current COVID-19 (n = 54)
Serology, n (%)
IgG N	3 (20)	1 (1.9)
IgG S	10 (66.7)	2 (3.7)
IgG N and/or S	10 (66.7)	3 (5.6%)
T-cell response, n (%)
S1+	11 (73.3)	7 (13.0)
SNMO+	14 (93.3)	8 (14.8)
S1 and/or SNMO+	14 (93.3)	10 (18.5)

n: number; +: positive; S1: spike antigens; SNMO: spike, nucleoprotein, membrane, and open reading frame antigens.

Ten study participants were lost to follow-up, but no consent was withdrawn. Participants with complete follow-up (n = 59) provided a median of 14 (range 3–18) RT-PCR tests during the study; 12 (median, range 3–13) as per the study protocol and 2 (median, range 0–7) due to symptoms or contact tracing). A positive RT-PCR test was detected in 14/59 (22.0%) participants (of which two recently had recovered from COVID-19 but remained RNA positive at inclusion). The study’s testing protocol captured around half of all new infections (7/12, 58.3%). Out of the 11 participants that tested positive for SARS-CoV-2 RNA after the study start (one was positive at inclusion) three were T-cell responsive at inclusion (one was responsive to S1 and SNMO antigens, and two were responsive only to SNMO antigens).

All study participants with complete follow-up received two doses of a SARS-CoV-2 vaccine during the study. The median number of days from the second vaccine dose to the date of the blood sampling at the study end was 278 days (IQR 112, range 9–287). At the end of the study, 54/59 (91.5%) participants demonstrated a T-cell response to S1 antigens (Table 3). The 14 participants that demonstrated a T-cell response to SNMO antigens at the study start remained SNMO responsive at the study end. All but two of the 11 participants that contracted COVID-19 after inclusion were SNMO responsive at the study end. The magnitude of the T-cell response to S1 antigens reported as normalised SFU/well (median), was 21.1 (IQR 33.9) for participants with previous COVID-19 and 12.5 (IQR 16.9) for participants with no previous COVID-19 at study end (p = 0.003, Table 3). A statistically significant correlation between the magnitude of the S1 T-cell response and IgG S antibody levels was found (p = 0.02) (Figure 1), but not to time since the second vaccine dose (Figure 2).

Figure 1. Magnitude of T-cell response to S1 antigen (SFU/well) [normalised to the positive control (anti-CD3)] and IgG S antibody levels (BAU/mL) at the study end (n = 59).

Figure 2. Magnitude of T-cell response to S1 antigen (SFU/well) [normalised to the positive control (anti-CD3)] and days passed since the second vaccine dose (n = 59).

Table 3. T-cell response to S1 and SNMO antigens after vaccination for participants with previous COVID-19 (as reported by the participant and/or diagnosed during the study) and participants with no COVID-19.

	COVID-19 and vaccination (n = 24)	Vaccination only (n = 35)
S1+, n (%)	24 (100)	30 (85.7)
SFU/well*, median (IQR)	21.1 (33.9)	12.5 (16.9)
SNMO+, n (%)	24 (100)	23 (65.7)
SFU/well*, median (IQR)	20.8 (54.6)	11.7 (10.1)
T-cell responsive (S1 and/or SNMO), n (%)	24 (100)	31 (88.6)

S1: spike antigens; SNMO: spike, nucleoprotein, membrane, and open reading frame antigens; +: positive; n: number; SFU: spot forming units; IQR: interquartile range.

Magnitude of T-cell response (median SFU/well) is presented for T-cell responsive participants.

*Normalised to 1000 anti-CD3 (positive control).

All participants were positive for IgG S at the study end (median BAU 243/mL, IQR 458, range 25.7–5677). Days passed since the second vaccine dose (p = 0.015), and no positive PCR during the study (0.003) were negatively associated with IgG S levels, whereas previous COVID-19 (as reported by participants or detected by PCR at inclusion) (p = 0.533), age (p = 0.784) and sex (p = 0.757) were not correlated with IgG S levels at the end of the study. None of the 11 participants who tested positive for SARS-CoV-2 RNA during the study were positive for IgG N at the study end.

Discussion

In this pragmatic prospective cohort study of HWCs at an ID department, 93% were T-cell responsive to S1 and/or SNMO antigens a median of nine months after the second dose of SARS-CoV-2 vaccine. Our study has one of the most extended follow-up times after vaccination and confirms the results of previous studies reporting durable T-cell responses following vaccination [7]. A recently published study demonstrated that vaccine-induced cellular immunity is highly conserved for new variants, including Omicron [8]. Spike-specific CD8⁺ and CD4⁺ T-cell responses were long-lasting and cross-reactive towards different variants and are a plausible explanation for the robust vaccine protection against severe disease even in the absence of neutralising antibodies. After vaccination, testing T cell reactivity against the SNMO pool containing 47 synthetic peptides derived from the spike, nucleocapsid, membrane, ORF3a, and ORF7 proteins, did not add any additional information over using the S1 peptide pool as the stimulus in our study. In a study including recovered COVID-19 patients, B7/N105⁺ CD8⁺ T cell responses elicited by non-spike epitopes were associated with long-lasting protection against severe disease [9]. Studies comparing the clinical impact of different T-cell responses in terms of protection and perseverance over time are needed. Vaccines that can induce broader T cell responses may be an attractive strategy to fight future variants where more conserved antigens can preferentially be targeted to create long-term cross-reactive protection [10].

The proportion of seronegative individuals with T-cell response before vaccination (12/56, 21.4%) was in line with two previous Swedish studies that used different methods to determine T-cell responsiveness [11, 12]. It has been described that T-cell epitopes are shared with other beta coronaviruses, and that cross-reactivity could be due to previous infections. The clinical protection of this cross-reactivity is not well known. In our study, we conclude that 3/11 participants that contracted COVID-19 during the study were T-cell responsive before their SARS-CoV-2 infection.

IgG S antibody titres decrease consistently six months after the last vaccine dose [13]. However, previous COVID-19 infection, young age, and female sex have been reported to be positively correlated with remaining humoral responses [14], which may explain why all the participants in our cohort of predominantly young female HCWs were positive for IgG S antibodies even after a median of nine months after vaccination. In addition, 41% of the participants had confirmed COVID-19 at baseline or during follow-up. A weak positive association between the magnitude of the T-cell response to S1 antigens and IgG S levels was found (Figure 1). However, the magnitude of the T-cell response to S1 antigens did not correlate to the time passed since vaccination, suggesting that a steady state of circulating memory T cells were available for recall stimulation with spike peptides. In contrast, the production of circulating IgG S levels diminished over time in line with decreased production of spike-specific IgG from plasma cells in the absence of an ongoing T cell activation. It would be interesting to stimulate memory B cells and investigate their spike-specific recall response. The IgG-N assay used in this study had insufficient sensitivity and, therefore, could not be used to distinguish individuals with previous SARS-CoV-2 infection.

This study has several limitations. The study size is small, and some participants were lost to follow-up. We intended to capture asymptomatic/pre-symptomatic infections by screening PCR every four weeks, but as participants were able to choose to provide a sample either on a workday or on a day off work, and no data on symptoms at the time of sampling were collected, we could not verify that all participants that provided a positive test as part of the study protocol were truly asymptomatic at the time of testing. T-cell analysis was performed on freshly prepared peripheral blood monocytes, but sampling conditions and transportation to the laboratory could have impacted the results. The study was not designed to evaluate vaccine efficacy, and thus, we did not collect information about the type of vaccine. The study’s strengths include the paired T-cell data pre and post-vaccination, one of the most extended follow-ups of T-cell responses after vaccination, frequent RT-PCR testing during the study, and a commercially available test for T-cell response analysis applied under real-life conditions.

To summarise, we found the T-cell test used in this study to be comparatively sensitive to previous COVID-19 infections in unvaccinated individuals. T-cell analysis was confirmatory to a positive IgG S antibody response, and the IgG S titres and the magnitude of T-cell response to the S1 antigen were also weakly correlated. Almost all participants were T-cell responsive to the S1 antigen after two doses of a SARS-CoV-2 vaccine, and the time since the last vaccine dose did not seem to influence the magnitude of the T-cell response. This is noteworthy since the T-cell response likely is important for protection against severe infections. Further studies are needed to determine how the magnitude of T-cell S1 response to SARS-CoV-2 antigen is best assessed and validly compared between sampling dates and individuals.

Acknowledgements

The authors thank the staff at the Departments of Infectious Diseases and Laboratory Medicine, Västmanlands hospital Västerås, for contributing to the study, the research nurses Marie Stenius Svensson and Angelica Norling, Centre for Clinical Research, Region Västmanland, for their skillful assistance, and AFA Försäkring, Stockholm, Sweden and Centre for Clinical Research, Region Västmanland, for financial support.

Disclosure statement

Robert Wallin has been a consultant for ABC Labs that performed CE-mark certification testing of Mabtech ELISpot Confirm kit: SARS-CoV-2 (T cell, INF-ƴ). Ola Winqvist is the Medical Director and Chief Scientific Advisor of ABC Labs.

Additional information

Funding

This work was supported by the AFA Försäkring in Sweden and Centre for Clinical Research, Region Västmanland - Uppsala university.