T cell immunity to natural SARS-CoV-2 infection may be more robust and longer lived than Ab responses. Accurate assessment of T cell responses is critical for understanding the magnitude and longevity of immunity across patient cohorts, and against emerging variants. By establishing a simple, accurate, and rapid whole blood test, natural and vaccine-induced SARS-CoV-2 immunity was determined. Cytokine release in whole blood stimulated with peptides specific for SARS-CoV-2 was measured in donors with previous PCR-confirmed infection, suspected infection, or with no exposure history (n = 128), as well as in donors before and after vaccination (n = 32). Longitudinal assessment of T cell responses following initial vaccination and booster vaccination was also conducted (n = 50 and n = 62, respectively). Cytokines were measured by ELISA and multiplex array. IL-2 and IFN-γ were highly elevated in PCR-confirmed donors compared with history-negative controls, with median levels ∼33-fold and ∼48-fold higher, respectively. Receiver operating curves showed IL-2 as the superior biomarker (area under the curve = 0.9950). Following vaccination, all donors demonstrated a positive IL-2 response. Median IL-2 levels increased ∼32-fold from prevaccination to postvaccination in uninfected individuals. Longitudinal assessment revealed that T cell responses were stable up to 6 mo postvaccination. No significant differences in cytokine production were observed between stimulations with Wuhan, Delta, or Omicron peptides. This rapid, whole blood–based test can be used to make comparable longitudinal assessments of vaccine-induced T cell immunity across multiple cohorts and against variants of concern, thus aiding decisions on public health policies.

Understanding the longevity of the adaptive immune response to SARS-CoV-2 is critical for devising public health policies to prevent reinfection. The ability to accurately measure the protective endurance of both memory B and T cells is fundamental to this understanding. Concerns about rapidly waning Ab levels after viral clearance have been raised (1). However, from what is known of viral immunology specifically regarding immune protection against coronaviruses, T cell responses may be more robust and longer lived (25). Initial studies revealed that virus-specific T cell responses developed in nearly all individuals with confirmed SARS-CoV-2 infection with responses persisting for at least 6 mo postinfection (611).

Once vaccination programs were established, vaccine-induced T cell responses could be investigated. Early studies demonstrated that SARS-CoV-2 vaccines were efficient at generating broad, protective T cell responses (1215). At the time of writing, 11.4 billion SARS-CoV-2 vaccine doses have been administered globally, but little is known about the longevity of the T cell responses they induce. Simple methods for accurately screening viral-specific T cell responses at a population level are needed to assess long-term vaccine efficacy. With the emergence of new SARS-CoV-2 variants, it is imperative to understand how mutations may affect the efficacy of the vaccine-induced T cell response, given data describing the enhanced escape capabilities these variants have from the humoral response (16, 17).

Traditional methods of measuring T cell responses are time-consuming, difficult to standardize, and require technical knowledge and specialized equipment. Moreover, current commercial tests such as ELISpot and flow cytometry require the isolation of PBMCs from whole blood and solely measure T cell production of IFN-γ, although other cytokines may provide a better indication of antiviral responses (10). Simple, “rapid” tests using whole blood, similar to those routinely used for diagnosis of tuberculosis, provide an alternative approach to measure viral-specific T cell responses (1823). In this study, we developed and optimized an in vitro whole blood stimulation assay to determine the most accurate biomarkers for identifying the presence of SARS-CoV-2–specific T cells in naturally infected individuals and in a cohort of individuals before and after vaccination. The high specificity and sensitivity of the test distinguished individuals with either natural and/or vaccine-induced T cell immunity from those individuals with little or no measurable immunity to SARS-CoV-2. This test was then used to study longitudinal T cell responses postvaccination, including boosters, and assess responses against the Delta and Omicron SARS-CoV-2 variants.

This study received ethics approval from the Wales Research Ethics Committee 5 (IRAS nos. 286991 and 305040). To investigate T cell responses in naturally infected individuals, participants from across Wales and England were recruited to the project between June and December 2020. All participants gave written, informed consent prior to inclusion. At the time of blood sample collection, corresponding details of prior test results for SARS-CoV-2 infection, confirmed by using PCR with reverse transcription from a nose and throat swab performed by public health bodies at accredited laboratories, were obtained via questionnaire. Those who had a positive PCR test were assigned to the “Confirmed infection (PCR positive)” cohort (n = 29). Participants were also asked if they had ever tested positive for SARS-CoV-2 via a lateral flow test, and to note any COVID-19–related symptoms they had experienced as well as record any close contact they had with persons who had been confirmed positive by PCR. Those who had a positive lateral flow test, experienced COVID-19–like symptoms, or had close contact with a positive individual were assigned to the “Suspected infection/exposure” cohort (n = 30). All other individuals were assigned to the “History negative” cohort (n = 69). None of the participants was hospitalized due to COVID-19 at any time before or during the study. Of this cohort, 80 were female, 36 were male, and 12 were unknown. Age and ethnicity are unknown.

To investigate T cell responses in vaccinated individuals, 50 participants from across Wales, England, and the United States were recruited to the project between December 2020 and July 2021. All participants gave written, informed consent prior to inclusion. Of this cohort, 32 were female and 18 were male; 9 were aged 18–29 y, 7 were aged 30–39 y, 8 were aged 40–49 y, 11 were aged 50–59 y, 2 were aged 60–69 y. and 13 were unknown; 42 were of white ethnicity, 1 was of Asian (Chinese) ethnicity, and 7 were unknown. At the time of blood sample collection, details of prior test results for SARS-CoV-2 infection and details of SARS-CoV-2 vaccinations including date and vaccine manufacturer were obtained via questionnaire. The breakdown of vaccinations received are as follows: Pfizer-BioNTech = 39; Oxford/AstraZeneca (ChAdOx1-S) = 10; Johnson and Johnson/Janssen = 1.

A final cohort of donors was recruited between October 2021 and February 2022 from Wales, England, and the United States to investigate T cell responses following booster vaccinations (n = 62). Of this cohort, 44 were female and 18 were male; 10 were aged 18–29 y, 14 were aged 30–39 y, 13 were aged 40–49 y, 10 were aged 50–59 y, 14 were aged 60–69 y, and 1 was aged 70+ y; 60 were of white ethnicity and 2 were of Asian/Asian British ethnicity. Donors received one of four different combinations of vaccine type. The breakdown of vaccines received are as follows: Pfizer-BioNTech first/second/third doses = 25; Pfizer-BioNTech first/second doses/Moderna third dose = 14; Oxford/AstraZeneca (ChAdOx1-S) first/second doses/Pfizer-BioNTech third dose = 9; Oxford/AstraZeneca (ChAdOx1-S) first/second doses/Moderna third dose = 11; 3 donors did not state the vaccine manufacturer.

Of this cohort, 30 donors were measured for responses against the SARS-CoV-2 variants of concern, that is, Delta and Omicron.

The ancestral SARS-CoV-2 peptide pool consisted of 470 15-mer peptides overlapping by 11 aa, covering the entire proteome of the nucleocapsid phosphoprotein (Miltenyi Biotec, Bergisch Gladbach, Germany), membrane glycoprotein (Miltenyi Biotec), and the spike (S1 and S2) protein (JPT Peptide Technologies, Berlin, Germany). All peptides were purified by HPLC. Previous studies confirmed that stimulation with either 0.1 µg/ml/peptide or 1.0 µg/ml/peptide resulted in the same T cell responses (20, 24). From these studies, a midrange concentration of 0.5 µg/ml/peptide was chosen as the final concentration for all stimulations. The SARS-CoV-2 Delta and Omicron peptide pools consisted of the same peptides covering the nucleocapsid phosphoprotein and membrane glycoprotein but contained peptides from the mutated spike proteins (S1 and S2) of Delta (spike B.1.617.2) and Omicron (spike B.1.1.529), respectively (JPT Peptide Technologies).

A single 10-ml sodium heparin vacutainer (BD Biosciences) tube of blood was collected from each participant and processed in the laboratory within 24 h of blood draw. Whole blood samples (1 ml) were aliquoted into sterile T332 Micrewtubes (Simport Scientific, Saint-Mathieu-de-Beloeil, QC, Canada) containing pre-aliquoted peptides. Additionally, one tube contained the T cell mitogen PHA-L (Sigma-Aldrich, St. Louis, MO; final concentration 50 µg/ml) as a positive control, and another contained 50 µl of PBS (Sigma-Aldrich) (negative control). Samples were incubated at 37°C, 5% CO2 for 18–22 h. Tubes were then centrifuged at 2000 × g for 3 min before harvesting ∼200–300 µl of plasma from the top of each blood sample. Plasma samples were stored at −20°C for up to 6 wk prior to analysis by ELISA or Luminex xMAP array.

IFN-γ protein was measured by a commercially available IFN-γ ELISA MAX Deluxe kit (BioLegend, San Diego, CA), following the manufacturer’s instructions with a few modifications, including an additional point on the standard curve (1000 pg/ml), a 1-h incubation for standards, samples, and blanks, and a pre-read step (at 450 nm with just the tetramethylbenzidine substrate) to standardize development of the assay. When standard 2 reached 0.1 OD, stop solution was added and the plate was read at 450 nm. The amount of IFN-γ in each sample was analyzed using the Gen5 software eight-point standard curve. To calculate the T cell response to SARS-CoV-2, the amount of IFN-γ in the control (PBS only) sample was subtracted from the corresponding value for the SARS-CoV-2 peptide-stimulated sample and reported as pg/ml plasma. In the absence of a response to the peptides, the amount of IFN-γ was calculated as below the lower limit of detection. Therefore, a value equal to half the lowest value on the standard curve was given for that sample. The cutoff for determining a positive response was set by Youden’s index (J) (see Statistical analysis), which determined the optimal cutoff value of >22.1 pg/ml.

Protein levels of IL-2, TNF-α, MIP-1β, MCP-1, IL-1β, IL-4, IL-5, IL-6, IL-7, IL-8, IL-10, IL-12p70, IL-13 IL-17A, G-CSF, and GM-CSF were measured by commercially available human cytokine 17-plex assays and human cytokine Th1/Th2 assays (Bio-Rad, Hercules, CA). These cytokines were chosen as a broad Th1, Th2, Th17 screening panel, as previous studies have shown that in COVID-19 patients, there is an elevation in not only Th1 cytokines but also in Th2 and Th17 cytokines (20). Additionally, the IL-6–driven cytokine storm has been well characterized in severe COVID-19 disease (25, 26). Therefore, this large panel of cytokines was chosen to give the greatest chance of identifying the best or most accurate biomarkers for SARS-CoV-2 responses. A summary of cytokine production during viral infection has been well described by Shah et al. (27).

The mean fluorescence intensity (MFI) of each cytokine bead set was measured on a Bio-Plex 200 instrument (Bio-Rad). Cytokine concentration was calculated from control curves of standards provided in the kit. In the absence of a response to the peptides, the amount of cytokine was considered as below the lower limit of detection, and a value equal to half the lowest value on the standard curve was given for that sample. When a response was out of range above the upper limit of detection, this was recorded as double the amount of the highest value on the standard curve. The cutoff for determining a positive response was set by Youden’s index. The IL-2 optimal cutoff was determined as >18.8 pg/ml. Optimal cutoff values for IL-13 and IL-10 were >2.3 and >7.8 pg/ml, respectively.

GraphPad Prism version 9.0.1 was used for all statistical analyses of datasets. Significance was determined using either a Kruskal–Wallis test with Dunn’s multiple comparison test, a one-way ANOVA with Tukey’s multiple comparison test, paired t tests for matched individuals, or unpaired t tests for unmatched cohorts; a p value <0.05 was considered significant. Receiver operating characteristic (ROC) curves and corresponding sensitivity and specificity values were generated from the ROC analysis on GraphPad Prism. As a summary measure of the ROC curves, Youden’s index (J) was used to enable the selection of a threshold value (cutoff point for positivity) for that marker, while also determining both sensitivity and specificity of each biomarker (28, 29). Pearson R2 values and associated p values were calculated by simple linear regression on GraphPad Prism.

Blood samples from 128 participants were stimulated overnight with a mega pool of 470 SARS-CoV-2 peptides and the level of IFN-γ was measured in the plasma. Significant differences were observed in the magnitude of IFN-γ responses between individuals with previous PCR-confirmed infection and suspected infection (median IFN-γ = 188.8 and 58.7 pg/ml, respectively) and individuals with no history of exposure (median IFN-γ = 3.9 pg/ml) (Fig. 1A). In cohorts of PCR-positive individuals and those with suspected infection/exposure, 97% of individuals (28/29) and 80% of individuals (24/30), respectively, demonstrated a positive response. This was in comparison with 22% (15/69) of donors from the cohort of history-negative individuals. A positive response (>22.1 pg/ml) was determined by Youden’s index (see Materials and Methods).

FIGURE 1.

Measurement and diagnostic accuracy of cytokine production to determine SARS-CoV-2–specific T cell responses in naturally infected and history-negative individuals.

(A) IFN-γ and (B) IL-2 production in response to overnight stimulation with SARS-CoV-2 peptide mega pool. Results show median with 95% confidence interval. ****p < 0.0001. ns, not significant. (C and D) Receiver operating characteristic curves defining sensitivity and specificity readouts for (C) IL-2 (n = 49) and (D) IFN-γ (n = 59). Area under the curve (AUC) and associated p value are indicated.

FIGURE 1.

Measurement and diagnostic accuracy of cytokine production to determine SARS-CoV-2–specific T cell responses in naturally infected and history-negative individuals.

(A) IFN-γ and (B) IL-2 production in response to overnight stimulation with SARS-CoV-2 peptide mega pool. Results show median with 95% confidence interval. ****p < 0.0001. ns, not significant. (C and D) Receiver operating characteristic curves defining sensitivity and specificity readouts for (C) IL-2 (n = 49) and (D) IFN-γ (n = 59). Area under the curve (AUC) and associated p value are indicated.

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Additional SARS-CoV-2–induced cytokines were measured by a multiplex array to determine whether other biomarkers could more accurately distinguish natural infection from non-infection. A clear distinction in the magnitude of IL-2 responses was observed between individuals with previous PCR-confirmed infection and suspected infection (median IL-2 = 103.5 and 49.6 pg/ml, respectively) and individuals with no known history of exposure (median IL-2 = 3.1 pg/ml) (Fig. 1B). From the cohort of previously infected donors, all 29 individuals demonstrated a positive IL-2 response, in marked contrast to only 1 out of 69 from the history-negative cohort. In the cohort with suspected infection/exposure, 77% of individuals demonstrated a positive response (>18.8 pg/ml).

ROC curves were generated for IL-2 and IFN-γ. For IFN-γ, the area under the curve (AUC) value was 0.9465 with a p value <0.0001 (Fig. 1C). For IL-2, the AUC value was 0.9950 (p < 0.0001) (Fig. 1D). Additionally, Youden’s index was used to measure the potential effectiveness of each biomarker. For IL-2, a sensitivity of 99% and a specificity of 100% was achieved. For IFN-γ, a sensitivity of 80% and specificity of 97% was attained.

A significant difference was also observed for IL-13 between the PCR-positive and history-negative groups (p = 0.0193; data not shown). However, only 42% of the PCR-positive cohort recorded a result above the limit of detection, and consequently IL-13 was nowhere nearly as accurate as IL-2 or IFN-γ at differentiating between cohorts (AUC = 0.6669; data not shown). No other cytokine examined could discriminate between previously infected and history-negative individuals (IL-6, TNF-α, MIP-1β, IL-12p70, IL-4, IL-5, IL-10, G-CSF, IL-17A, IL-7, IL-8, IL-1β, MCP-1, GM-CSF; data not shown).

These results demonstrate that IL-2 is the most accurate cytokine biomarker for distinguishing SARS-CoV-2–infected from uninfected individuals.

To investigate whether IL-2 was an accurate biomarker for identifying vaccine-induced T cell responses, 34 individuals were recruited to donate blood samples prior to SARS-CoV-2 vaccination, and then again up to 6 wk following the first and/or second vaccine doses.

A marked and consistent increase in the magnitude of IL-2 responses was observed between pre- and postvaccination in previously unexposed individuals. The median IL-2 response increased by ∼32-fold from 6.6 pg/ml prior to vaccination to 194.6 and 210.1 pg/ml after the first and second vaccine doses, respectively (Fig. 2A). Following vaccination, 100% of individuals demonstrated a positive IL-2 response (>18.8 pg/ml).

FIGURE 2.

Measurement of cytokine production to determine SARS-CoV-2–specific T cell responses in individuals pre- and postvaccination.

(A and B) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in unexposed (A) and previously infected (B) individuals pre- and postvaccination. (C) IFN-γ, (D) IL-13, and (E) IL-10 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in previously unexposed individuals pre- and postvaccination. Square symbol indicates Janssen vaccine (A only). Results show median with 95% confidence interval. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. ns, not significant.

FIGURE 2.

Measurement of cytokine production to determine SARS-CoV-2–specific T cell responses in individuals pre- and postvaccination.

(A and B) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in unexposed (A) and previously infected (B) individuals pre- and postvaccination. (C) IFN-γ, (D) IL-13, and (E) IL-10 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in previously unexposed individuals pre- and postvaccination. Square symbol indicates Janssen vaccine (A only). Results show median with 95% confidence interval. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. ns, not significant.

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In the cohort of previously infected individuals, all donors demonstrated a positive IL-2 response following the first and second doses of vaccine (Fig. 2B). Vaccination did not boost the magnitude of responses from the levels of IL-2 found prevaccination, with the median level of IL-2 produced by these donors being 204.5 pg/ml prevaccination, and 100.5 and 179.5 pg/ml after the first and second vaccine doses, respectively.

Further Th1 and Th2 cytokines were measured in a small cohort of donors (n = 13). Significant differences similar to those observed for IL-2 were detected in the magnitude of IFN-γ responses between pre- and postvaccination in previously unexposed individuals (Fig. 2C). Median IFN-γ levels increased by ∼8-fold following vaccination. In the prevaccinated group, none of the donors demonstrated a positive IFN-γ response (>22.1 pg/ml), in noticeable contrast to 90 and 83% following one or two vaccine doses.

The magnitude of the IL-13 response was small (<10 pg/ml), but the differences between pre- and postvaccination were significant (Fig. 2D). After the first vaccine dose, 85% of donors showed a positive IL-13 response (>2.3 pg/ml), dropping to 67% in donors having received two doses. The magnitude of the IL-10 response was again small, but a significant difference was observed between prevaccinated samples and samples taken after the second vaccine dose only (Fig. 2E). All samples gave a positive IL-10 result following two vaccinations; however, so did 31% of the unvaccinated samples (>7.8 pg/ml). Cytokines TNF-α, IL-12p70, IL-4, IL-5, and GM-CSF were also measured but none was effective at differentiating unvaccinated from vaccinated individuals (data not shown).

These results confirm that IL-2 was the most accurate biomarker for distinguishing unvaccinated and vaccine-induced T cell responses.

IL-2 levels were measured in a cohort of 15 individuals at 3–4 wk postvaccination, and then again at 4–6 mo postvaccination (Fig. 3A). The mean values at each time point were remarkably similar, with the mean IL-2 values being 222.6 and 218.7 pg/ml at 4 wk and 6 mo postvaccination, respectively (p = 0.9252). Only one individual showed a decrease in IL-2 levels from a positive to a negative result and, interestingly, this individual had the lowest starting value of 19.5 pg/ml, only just above the cutoff for positivity (18.8 pg/ml).

FIGURE 3.

Longitudinal T cell responses following vaccination.

(A) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in matched individuals at 1 and 4–6 mo postvaccination. (B) Regression analysis of IL-2 production and the number of days postvaccination. (C) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in matched individuals at 6 mo after the second vaccination and 1 mo after the third vaccination (booster). (D) Regression analysis of IL-2 production and the number of days after booster vaccination. Dashed line indicates positivity cutoff. *p < 0.05; ns, not significant (A and C). R2 and p values are indicated; each symbol represents an individual donor (B and D).

FIGURE 3.

Longitudinal T cell responses following vaccination.

(A) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in matched individuals at 1 and 4–6 mo postvaccination. (B) Regression analysis of IL-2 production and the number of days postvaccination. (C) IL-2 production in response to overnight stimulation with the SARS-CoV-2 peptide pool in matched individuals at 6 mo after the second vaccination and 1 mo after the third vaccination (booster). (D) Regression analysis of IL-2 production and the number of days after booster vaccination. Dashed line indicates positivity cutoff. *p < 0.05; ns, not significant (A and C). R2 and p values are indicated; each symbol represents an individual donor (B and D).

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This longitudinal analysis was extended to include more individuals who donated a blood sample anywhere between 3 and 199 d postvaccination (two doses). We evaluated whether there was any correlation between the magnitude of the IL-2 response and the number of days that had passed since receiving the second vaccine. Regression analysis revealed that the IL-2 response did not significantly decrease over time (R2 = 0.0045, p = 0.6043; (Fig. 3B). This suggests that the vaccine-induced T cell response did not wane within ∼200 d of vaccination. Furthermore, when the magnitude of IL-2 from donors vaccinated with either Pfizer-BioNTech (n = 39) or Oxford/AstraZeneca (ChAdOx1-S) (n = 10) was compared, the mean response was highly comparable (215.4 and 184.3 pg/ml, respectively, p = 0.6774; data not shown), demonstrating equal efficacy of both vaccines.

To investigate the effect booster vaccination had on the magnitude of the IL-2 response, 16 individuals were recruited to donate a blood sample 6 mo after their second vaccination, and at 3–4 wk after booster vaccination (third dose). Following booster vaccinations, the IL-2 levels doubled from a mean of 77.4 pg/ml before booster to 141.8 pg/ml after booster, and, importantly, one donor who gave a negative T cell response before booster demonstrated a positive T cell response after booster (Fig. 3C). Despite the IL-2 levels increasing significantly (p = 0.0275) in the cohort overall, the effect was not as dramatic as the results seen before and after initial vaccination, with ∼30% of the donors giving either similar or slightly decreased IL-2 levels following booster vaccination.

Longitudinal analysis of responses following booster vaccination was then carried out, with 61 individuals donating blood anywhere between 20 and 131 d after booster (third dose). We again evaluated whether any correlation existed between the magnitude of the IL-2 response and the number of days that had passed since receiving the booster vaccine. Regression analysis revealed that the IL-2 response did not significantly decrease over time (R2 = 0.0001, p = 0.9261; (Fig. 3D). This suggests that the vaccine-induced T cell response following a booster did not wane within ∼130 d. Additional analysis was performed to compare the magnitude of IL-2 response between donors vaccinated with four different combinations of Pfizer-BioNTech, Oxford/AstraZeneca (ChAdOx1-S), and Moderna vaccines (see Study cohort). Statistical analysis did reveal small differences in immune responses elicited by the different combinations (p = 0.0383, data not shown). However, given the small numbers in each group, this warrants further investigation. Overall, receiving a booster from any combination of vaccines provides relatively comparable efficacy to induce T cell responses.

Blood samples from 30 donors were stimulated with peptides from the ancestral (Wuhan) virus as well as peptides from the mutated Delta and Omicron viruses. IL-2 levels following stimulations showed no significant differences (p = 0.0535), with mean values of 100.3, 127.6, and 114.3 pg/ml achieved for Wuhan, Delta, and Omicron peptides, respectively (Fig. 4A). Additional analysis for IFN-γ revealed a similar pattern, with mean IFN-γ levels of 228.7, 237.6, and 257.0 pg/ml for Wuhan, Delta, and Omicron peptides, respectively (p = 0.3209; (Fig. 4B). These highly comparable measurements from both cytokines indicate that the T cell response induced by the currently available vaccines are unaffected by mutations in the spike protein.

FIGURE 4.

Measurement of cytokine production against SARS-CoV-2 variants.

(A) IL-2 and (B) IFN-γ production in response to overnight stimulation with SARS-CoV-2 peptide pools containing peptides from spike proteins from different variants. Results show mean ± SEM. ns, not significant.

FIGURE 4.

Measurement of cytokine production against SARS-CoV-2 variants.

(A) IL-2 and (B) IFN-γ production in response to overnight stimulation with SARS-CoV-2 peptide pools containing peptides from spike proteins from different variants. Results show mean ± SEM. ns, not significant.

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Simple, rapid, and functional assays that accurately detect SARS-CoV-2–specific T cell responses are essential for comparing T cell immunity across multiple population cohorts and for long-term assessments of vaccine efficacy. The results show that measuring plasma IL-2 from SARS-CoV-2 peptide-stimulated whole blood accurately distinguishes between COVID-19 convalescents and uninfected healthy blood donors with a high degree of sensitivity and specificity, with plasma IFN-γ also demonstrating high levels of accuracy. Plasma IL-2 was also an excellent biomarker for distinguishing vaccinated and unvaccinated individuals. These observations are in line with findings from Bertoletti and colleagues (22) and Smith and colleagues (23), among others, who also observed that IL-2 and IFN-γ provided the most accurate readout of whole-blood based assays.

There are several benefits of using the mega peptide pools rather than whole viral proteins. First, the peptides span the entire immunogenic region of the viral proteome and include both class I and class II HLA peptides, so they will capture responses from both CD4+ and CD8+ T cells. Additionally, the peptide pools are specifically designed to provide HLA-independent stimulation, which is of importance due to the unavailable information on the HLA type of the donors in this study, as it is known that HLA genotype may influence the severity and progression of COVID-19 (30). Furthermore, the nature of the peptides is such that the APCs can be loaded directly with these peptides without the need for lysosomal or proteasomal processing. One potential limitation of using commercially available overlapping peptide pools is that it may not produce as much T cell activity compared with using larger experimentally derived mega pools of peptides (31). However, as experimentally derived mega peptide pools are not commercially available, our current mega pool of peptides still yields fantastic results as they cover the major immunodominant regions of the spike, matrix, and nucleoprotein viral proteins. Additionally, T cell responses in SARS-CoV-2 are estimated to recognize even more epitopes per donor than seen in the context of other RNA viruses, meaning that even if the response measured is not a maximal one, we are still likely to pick up some responses in our donors, indicating that false negatives are unlikely.

Although most cytokine production is assumed to come from T cells, we cannot rule out the possible contribution from other cell types such as NK cells or myeloid cells. NK cells have been shown to produce IFN-γ when SARS-CoV-2–specific peptides bind to NKG2D receptors (32). However, NK cells are not widely regarded as large producers of IL-2, which is produced primarily by activated CD4+ T cells. DCs have also been shown to produce small amounts of IL-2, but this is in response to LPS and is not peptide induced (33). Therefore, we, similar to several others, refer to this method of whole blood cytokine release via mega peptide pools as a “T cell test,” although we do acknowledge the caveat described above (6, 34).

T cell responses in previously unexposed individuals developed early after just one vaccination and were maintained after a second dose, a similar finding to that observed by De Rosa et al. (35). Interestingly, the amount of IL-2 in individuals with a previous positive PCR result following vaccination was not amplified and was comparable to those with no history of SARS-CoV-2 infection, a finding that was not observed using ELISPOT (36, 37).

Positive IL-2 responses were detectable up to 200 d after initial vaccination, and up to 130 d after booster vaccination, in keeping with observations that T cell responses are robust. In SARS-CoV-1– and MERS-CoV–infected individuals, T cell responses were detected several years postinfection (35). It was also reassuring that no significant differences were observed between T cell responses induced by either Pfizer-BioNTech or Oxford/AstraZeneca (ChAdOx1-S) after the initial two doses of vaccines. Conversely, some disparities in T cell responses were observed with different combinations of initial and booster vaccinations. However, this is a relatively small subset of individuals and does not take into account other variables such as age, which is known to influence T cell responses. Consequently, it is difficult to draw any firm conclusions from this analysis as it currently stands, yet it does suggest further investigation is warranted to understand the best combination for certain individuals to achieve the most efficacious combination possible. This may have more of a profound impact on vulnerable cohorts of individuals who may respond differentially to mRNA vaccines versus adenovirus vaccines for example.

The lack of any dramatic increase in the magnitude of IL-2 production following a booster vaccination was somewhat unanticipated, given the evidence demonstrating that a booster with either Moderna or Pfizer resulted in an ∼12-fold increase in Ab levels (38). One donor who gave a negative IL-2 response before booster did demonstrate a considerable increase in the magnitude levels of IL-2 production, taking them above the cutoff for positivity. This may suggest that boosters may be of higher importance for those individuals who have a very low or undetectable immune response, highlighting the need for population screening for T cell immunity to identify those individuals more “at risk” from severe infection.

Highly comparable T cell responses between the ancestral virus and Delta and Omicron variants confirm what is already known about T cell responses against these variants. In-depth immunological analyses have revealed that there are high levels of cross-reactivity between ancestral SARS-CoV-2–specific memory T cells and multiple viral variants (3941). Given the simplicity of this whole blood test, it is reassuring to see that results generated in the present study mimic those findings. Because of the large number of epitopes that both CD4+ and CD8+ T cells have been shown to recognize, it is unlikely that mutations of a few key viral epitopes will lead to SARS-CoV-2 being able to escape T cell recognition (31). However, should a variant of concern arise that somehow interferes with T cell activation due to mutations affecting multiple T cell epitopes, we predict that our test would be able to detect this.

Reliable biomarkers are integral to assess vaccine efficacy over time. Unlike other immunoassays that detect Ag-specific T cell responses such as ELISPOT and flow cytometry, this virus-specific cytokine release assay is simple to perform and can be employed across multiple laboratories for large-scale epidemiological studies in conjunction with testing for neutralizing Abs (42). The IL-2 release assay is a promising tool for use in multicenter clinical trials for the development of new vaccines or treatments against novel SARS-CoV-2 variants as they arise globally. The adaptive immune response is a major determinant for clinical outcome after SARS-CoV-2 infection. The strong and robust T cell response elicited from vaccinations contributes to protection against hospitalization or death (43). Continued analyses on T cell responses such as those described in the present study will aid decision making by health policy makers. This will include decisions concerning vaccine booster requirements and monitoring of SARS-CoV-2 infection in immune-suppressed and other at-risk populations, especially from new variants that are widely predicted to arise.

We thank Dr. Hanne M. Hoff (Crondall New Surgery, Farnham, Surrey, U.K.) for organizing participant cohorts and blood sample acquisition.

This work was supported by two research grants awarded to InBio: Innovate UK Grant 66476 and Innovate UK/Small Business Research Initiative Grant 10008511.

Abbreviations used in this article

     
  • AUC

    area under the curve

  •  
  • ROC

    receiver operating characteristic

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M.A.O., R.T.M., B.R.S., M.D.B., and N.F.B. are employees of InBio. M.D.C. is a co-owner and employee of InBio.

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