From 19 November 2020 to 9 February 2021, pre-vaccination blood was collected from a group of 19 people who use drugs (PWUD). All except one patient were HIV seronegative, and seen subjects received replacement therapy with methadone or buprenorphine. None of the patients presented detectable levels of anti-SARS-CoV-2 RBD or nucleocapsid IgG antibodies in the serum and had no history of COVID-19 infection. The post-vaccination cohort consisted of 25 PWUD donors. For this group, blood collection was performed in November 2021, 10–30 weeks after the participants had received their second dose of (mRNA) vaccines Moderna/mRNA-1273 or Pfizer/BioNTech BNT162b2. One of these donors received only one dose of mRNA vaccine but was diagnosed with SARS-CoV-2 46 days before blood sampling. One patient of this group had HIV and was on antiretroviral therapy, whereas eight subjects from this group belonged to the pre-vaccination cohort. Patients on medication-assisted rehabilitation were predominantly prescribed methadone or buprenorphine. See Supplementary Tables S1 , S2 for a summary of the clinical information and characteristics of all study participants. In addition, we tested 12 cryopreserved PBMC samples previously bio-banked from non-exposed seronegative healthy donors (HD) and five samples collected from COVID-19 convalescent donors. A cohort of 21 healthy subjects who also received two doses of mRNA vaccines was used as a control for post-vaccination studies.

This study and all biobanking procedures carried on at Oslo University Hospital were approved by the Norwegian Regional Ethics Committee (approval nos. 166545 and 135924) and the Data Protection Officer at the Oslo municipality. Informed consent was obtained from all participants.

PBMC and serum samples were collected at a single time point for each patient ( Supplementary Table S2 ).

PBMCs were isolated from the whole blood using either Cellular Preparation Tubes (CPT) tubes (BD vacutainer, # 362782) or Lymphoprep™ (Serumwerk Berburg, # 1858). CPT tubes were spun at 1,600g for 25 min at RT and plasma collected and stored at −20°C for future analysis. Lymphoprep™ isolation was performed according to the manufacturer’s instructions. Briefly, a 1:1 mix of blood and PBS (35 mL) was layered onto 10–15 mL Lymphoprep™ solution in a 50-mL Falcon tube. Tubes were then centrifuged for 25 min at 800g and PBMC collected, washed in PBS (Gibco, # 10010-015), and spun three more times (400g, 7 min, 4°C) before being counted and resuspended in Fetal bovine serum (FBS) (Gibco, # 10270-106) complemented with 10% Dimethyl sulfoxide (DMSO). After overnight pre-chilling at −80°C in Mr. Frosty (Nalgene™, # 5100-0001), cells were transferred to liquid nitrogen for long-term storage.

IgG anti-RBD (BAU/mL) and IgG anti-nucleocapsid serum antibodies were quantified using a multiplexed bead-based assay as previously described (40).

To test SARS-CoV-2-specific reactivity of T cells from pre-vaccinated donors, PBMC from each patient were distributed on a 96-well round bottom cell culture plate at a concentration of 3×10⁵ cells per well (~5×10⁴ and ~1×10⁵ CD8⁺ and CD4⁺ T cells, respectively) and stimulated with a pool of SARS-CoV-2 peptides (one peptide per well at 1.5 µg/mL: final volume = 200 µL per well; for the list of the 9–10-mer peptides used in these studies, see Supplementary Methods ) (41, 42). At day 3, supernatants (SN) were collected for cytokine analysis (ELISA), after which methyl-³H thymidine (Hartmann Analytic GmbH, 0.02 mCi/mL) was added for cell proliferation assessment. After a 5- to 6-day incubation period, cells were harvested and counted using a PerkinElmer MicroBeta2 2450 Microplate Scintillation Counter. Proliferation data were calculated by subtracting the background signal (unstimulated cells). IFNγ ELISA was performed on supernatants using the following reagents: IFNγ monoclonal antibody (2G1, Invitrogen), Human IFNγ ELISA Standard Recombinant Protein (eBioscience), biotinylated IFNγ monoclonal antibody (B133.5, Invitrogen), horseradish peroxidase substrate TMB solution (Merck), and Streptavidin-rHRP (Southern Biotech). Plates were read with a Tecan multiplate reader. IFNγ levels were calculated by subtracting the background signal from unstimulated control wells or the average signal from the six wells with peptides that gave the lowest response.

Two collections of lyophilized 15-mer peptides with 11 amino acid overlaps were used in this study: one (PepTivator® SARS-CoV-2 Prot_S; Militenyi, # 130-126-700) covered the immunodominant regions (aa 304–338, 421–475, 492–519, 683–707, 741–770, 785–802, and 885–1,273), and the other (PepTivator® SARS-CoV-2 Prot_S Complete; Miltenyi, # 130-127-953) covered the entire length of the SARS-CoV-2 S glycoprotein (aa 5-1273; Protein QHD43416.1, GenBank MN908947.3). The combination of these two pools was named “Peptivator mix” and used to stimulate cells. In addition to the Peptivator mix, a collection of 101 9/10-mer spike and non-spike peptides identified by the NEC Immune Profiler (NIP) algorithm (43) was used to test CD8⁺ T-cell response. These epitopes were selected as top binders for the most prevalent HLA types in Norway, including HLA-A*01:01, HLA-A*02:01, HLA-A*03:01, HLA-A*23:0, HLA-A*29:02 HLA-B*07:02, HLA-B*08:01, HLA-B*15:01, HLA-B*15:02, HLA-B*40:01, HLA-B*44:02, HLA-C*03:03, HLA-C*04:01, HLA-C*07:01, and HLA-C*07:02. Among these peptides, 43 derived from the spike protein and 58 from non-spike regions (see Supplementary Methods ). Additionally, a pool of 23 9/10-mer spike peptides (the NOI pool) validated in our laboratory (41, 42) was used to identify vaccine response. The list of peptides is shown in Supplementary Methods .

Quantification of T-cell activation was performed using the activation-induced marker assay (AIM) as described (41, 44–46). Briefly, stimulated PBMCs were washed in cold RPMI 1640 medium with GlutaMAX™ supplement and enriched in live cell using the magnetic column protocol according to manufacturer instructions (MACS MultiStand, # 130-042-303 with OctoMACS™ Separator). PBMCs (10 million cells per mL) were then distributed on a 96-well round bottom cell culture plate (200 μL/well) in TexMACS medium (Miltenyi, # 130-096-197) supplemented with 1 mmol/L Sodium Pyruvate (Gibco # 11360-039), 1 mmol/L MEM NEAA (Gibco # 11140-035), 50 nmol/L 1-thioglycerol (Sigma-Aldrich, # M1753), 12 μg/mL Gentamycin (VWR, # E737), and 20 U/mL IL-2 (R&D # AFL202). After 3 h incubation, cells were washed and treated (1 h) with 15-mer or 9/10-mer peptide pools (1.5 µg of peptide/mL in a final volume of 200 µL per well) in the presence of anti-CD28/CD49d co-stimulatory antibodies (BD # 347690) at a final concentration of 1:200. At this time, BV711 anti-human CD107a antibody (Clone H4A3; BioLegend, # 328640) was added for assessment of T-cell degranulation by flow. After an additional 18 h incubation with Brefeldin A/Monensin cocktail (GolgiStop 500X, Invitrogen # 00-4980-93), which improves assay sensitivity (47), cells were washed once in 1× PBS containing 5% FBS and 0.1% sodium azide, and then stained in the dark for 10 min with 0.5 μL of Fixable Near IR Live/Dead viability stain (Molecular Probes, # L34976) in a 10-μL final volume of cold PBS containing 5% FBS. Cells were then permeabilized at 4°C for 20 min in 100 μL BD Cytofix/Cytoperm solution (BD # 554714) and washed two times in 200 µL of 1× BD perm/wash solution (BD Biosciences Fixation and Permeabilization kit; # 554714). They were then stained in 20 µL of 1× BD perm/wash solution containing the following fluorochrome-conjugated antibodies: AF488 anti-human IL-2 (Clone MQ1-17H12; BioLegend, # 500314), PerCP-Cy5.5 anti-human CD8 (Clone RPA-T8; BioLegend, # 301032), PE anti-human CD137 (Clone 4B4-1; BioLegend, # 309804), PE-CF594 anti-human Granzyme B (Clone GB11; BD # 562462), PE-Cy5 anti-human (CD4 Clone RPA-T4; BD # 566925), PE-Cy7 anti-human TNF (Clone MAb11; Invitrogen, # 25-7349-82), AF647 anti-human IFNγ (BioLegend, #502516), BV510 anti-human CD40L (Biolegend, #310830), BV605 anti-human CD3 (Clone SK7; BD # 563219), and BV421 anti-human Perforin 1 (Clone dG9; BioLegend # 308122). Antibody concentrations were used according to manufacturer instructions. After a 30-min incubation in the dark, cell pellets were washed once in PBS, resuspended, and acquired on an Attune NxT Flow Cytometer (Thermo Fisher). Further replicate assays were performed similarly, but with a simplified staining panel (see Supplementary Methods ).

Responding T cells were identified using the “OR” Boolean gating (FlowJo’s Boolean analysis tools) that were found within regions double positive for the following marker pairs: CD137⁺ CD154⁺, CD154⁺ IFNγ ⁺, CD154⁺ IL-2⁺, CD154⁺ TNF⁺, TNF⁺ CD107a⁺, TNF⁺ GZMB⁺, TNF⁺ IFNγ⁺, or TNF⁺ Perforin⁺ (for gating strategy, see Supplementary Figure S2 ). Unstimulated background was subtracted to identify the frequency of specific activated T cells. Further in-depth functional profiling of SARS-CoV-2-specific T cells was performed using FlowJo’s Boolean analysis tools (AND, OR, and NOT), and data were presented with the Simplified Presentation of Incredible Complex Evaluations (SPICE) 6.1 software. Generated data were used to visualize antigen-specific T-cell polyfunctionality (48, 49), where activated cells expressed combinations of CD137, CD154, IFNγ, IL-2, and TNF (for the CD4+ Th cell subset) and CD107a, GZMB, IFNγ, Perforin, and TNF (for the CD8+ CTL cell subset). The frequency of all possible permutations above unstimulated background was calculated for responding specific T cells. For CTL, we performed the analysis on IFNγ+ or TNF+ cells (as subsets of CTL constitutively express GZMB and Perforin). Median frequencies of each population for each cohort were represented in a SPICE pie chart. A permutation test from the SPICE software was used to compare the distribution of cell populations for each pair of pie charts. Values of p < 0.05 were considered statistically significant. A detailed list of the possible permutations of polyfunctional populations can be found in Supplementary Table S3 .

Statistical analyses were performed using GraphPad Prism V.9.3.1 (GraphPad software). Two-tailed Mann–Whitney U test, Spearman’s rank correlation coefficient, Principal Component Analysis (PCA) plots loading, and Pearson correlation matrix plots were used as indicated.

In 2020, public health concerns during the height of the COVID-19 pandemic led us to collect blood samples to investigate humoral responses in PWUD. We found that only 4 of 99 PWUD developed an IgG anti-RBD response in the months before vaccines were available (not shown), a rate that was similar to what we have found in the general population (39). Of the 36 PWUD donors who participated in the current study, 28 used illegal or prescribed opioids, 19 were in the pre-vaccination group, and 25 were in the post-vaccination group. Both pre- and post-vaccination samples were available for eight donors, one of whom received only one dose of vaccine but was diagnosed with SARS-CoV-2 46 days before blood was sampled (see Materials and Methods, Supplementary Tables S1, S2 ). We first tested T-cell responses in unexposed PWUD and healthy donors (HD) who were not vaccinated and were seronegative for anti-RBD and anti-nucleocapsid antibodies. In these groups, a response would likely be the result of cross-reactivity of T cells generated from prior exposure to endemic seasonal HCoV. We tested T-cell response to single peptides derived from spike (N = 43) or non-spike proteins (ORF1ab, ORF3A, Envelope; N = 57) in three sub-cohorts: PWUD (N = 19), HD (N = 12), and, as positive controls, COVID-19 Wuhan convalescent patients (N = 5; see Materials and methods and Supplementary Methods for peptide list). Overall, we found that proliferation ( Figure 1 ) and IFNγ secretion ( Supplementary Figure S1A ) were both significantly increased in convalescent donors and that although the proliferative response was similar between PWUD and HD donors, PWUD showed a slight decrease in IFNγ secretion ( Figure 1 , Supplementary Figure S1A ). In all the cohorts, the level of the proliferative response and IFNγ secretion was comparable notwithstanding the type of stimulus (spike vs. non-spike peptides; Figure 1B , Supplementary Figure 1B ). As expected, donors in the PWUD and HD cohorts presented with variable degrees of proliferative and secretory response ( Figure 1C , Supplementary Figure 1C ), but there was no significant difference between groups ( Figure 1D ; 621 vs. 746.5 CPM median values, p = 0.98 and Supplementary Figure S1D ; 13.2 vs. 22.8 pg/mL median values, p = 0.42) indicating that PWUD do not possess T-cell cross-reactivity generated from potential prior exposures to endemic seasonal HCoV.

In 2021, as vaccine rollout continued, we managed to secure follow-up blood samples from 25 PWUD donors ( Supplementary Tables S1, S2 ). Due to difficulties in recalling donors for sampling, blood was drawn between 67 and 208 days after the second dose (D2) and not after the optimal 30-day interval. A similar approach was thus employed to gather blood samples from healthy donors in 2022, but due to the surge of Omicron, the likelihood of breakthrough infection (BTI) and hybrid immunity in this control cohort was significantly increased. Indeed, four of these controls experienced breakthrough infection before collection. To assess vaccine-induced T-cell activation, we challenged PBMC with commercially available 15-mer spike peptide megapool mixes (Peptivator mix; see Materials and methods) and measured by flow cytometry the increase in the frequency of T-cell populations that were double positive for different combinations of selected activation-induced markers (AIMs). Gating strategy and AIMs used in the study are reported in Supplementary Figure S2 . We assessed the magnitude of spike-specific responses by determining the frequencies of cells that were double positive over background for one or more of the following marker pairs by “OR” Boolean gating: CD137⁺ CD154⁺, CD154⁺IFNγ⁺, CD154⁺ IL-2⁺, CD154⁺ TNF⁺, TNF⁺ CD107a⁺, TNF⁺ GZMB⁺, TNF⁺ IFNγ⁺, or TNF⁺Perforin⁺ (see Supplementary Figure S2 for gates). Frequencies of responding spike-specific cells showed that the level of response in PWUD and HD donors was not significantly different ( Figure 2A ). As sampling time was variable after vaccination in PWUD, we also visualized T-cell responses vs. time post-vaccination, but found no significant correlations in PWUD or HD ( Figure 2B ). HD controls were chosen to resemble PWUD in terms of time since vaccination, but after sampling, four were found to have had breakthrough infection (BTI) with Omicron VOC (as they were positive for IgG anti-Nucleocapsid) ( Figures 2A, B ). Furthermore, when marker combinations were taken separately, we observed that the cytotoxic response in PWUD (frequency increase in TNF⁺ GZMB⁺ events) was higher in PWUD than in HD, but that for other AIM combinations, such as those that included the expression of CD154 and TNF/IFNγ co-expression, the response was higher in HD. Moreover, these results did not appear to be influenced by the four donors with BTI in the control group ( Supplementary Figure S3 ). Representation by Principal Component Analysis (PCA) showed substantial overlap between cohorts and confirmed that, with the exception of a few PWUD patients who showed enhanced cytotoxic response (increased upregulation in perforin, granzyme B, and CD107a), CD4 Th cell responses in PWUD were similar to the response of the healthy subjects ( Figure 2C ). Further analysis revealed that the pattern of Th cell polyfunctionality in PWUD was not significantly different from the one observed in HD ( Figure 2D ). More specifically, we found that among all vaccine-specific, activated (CD137⁺) Th cells, the fraction of events that were double or triple positive for TNF, IFNγ, and/or IL-2⁺ were not significantly different between PWUD and HD ( Figure 2E ; 33% 18%, 12%, and 17% vs. 25%, 25%, 13%, and 14% median frequency values for TNF⁺ IFNγ⁺, TNF⁺ IFNγ⁺ IL-2⁺, and TNF⁺ IFNγ⁻ IL-2⁺ populations in PWUD and HD, respectively; Mann–Whitney U test, p > 0.05). A similar analysis of activated CD154⁺ Th cells produced similar results (data not shown). These data were further confirmed by Pearson correlation analysis, which showed that the correlation patterns among different expression markers measured in the two cohorts were similar ( Supplementary Figure S4A ). A comprehensive representation of the polyfunctional response is shown for both cohorts in Figure 2F , where we employed the SPICE tool to visualize and compare frequencies of single, double, triple, and quadruple positive cells within the activated CD4 Th cell compartment (see Materials and methods). This analysis confirmed the polyfunctional nature of the response in PWUD and showed that although the frequency of some vaccine-specific T-cell populations was lower in PWUD, the pattern of CD4 Th cell polyfunctionality was not significantly different from the one observed in healthy controls.

To investigate the response of CD8⁺ cytotoxic T lymphocytes (CTL), we stimulated the cells with the Peptivator mix or a pool of 23 9/10-mer spike peptides (the NOI pool) that were predicted by the NEC Immune profiler AI tool (see Supplementary Methods for peptide list) (43) and experimentally validated (41). We determined the frequency of spike-specific T cells after stimulation with either the Peptivator mix or the NOI pool by measuring CD8 CTL cell responses using the Boolean “Or” gating on the same double positive regions defined for the analysis of the Th cell subset. We found that the frequency of spike-specific CD8 CTL in PWUD and HD donors was not significantly different ( Figure 3A ), remained stable with time ( Figure 3B ), and was characterized by similar response patterns ( Figure 3C ). Likewise, the pattern of CD8 CTL cell polyfunctionality in PWUD was not significantly different from the one observed in healthy controls ( Figure 3D ), an observation further confirmed by Pearson correlation analysis ( Supplementary Figure S4B ), which showed overall similarity between cohorts in terms of immunophenotypic response. A more in-depth analysis of polyfunctionality revealed that among spike-specific (TNF⁺) CTL cells, the fraction of events that were double or triple positive for some markers, including IFNγ, CD107, IL-2, and/or GZMB, was similar to the HD group ( Figure 3E ; 1.8%, 2.0%, 0%, and 47% vs. 5.7%, 6.9%, 0%, and 33% median frequency values for IFNγ⁺ CD107a⁺, IL-2⁺, IL-2⁺ GZMB⁺, and GZMB⁺ populations in PWUD and HD, respectively), but other T-cell populations, such as IFNγ⁺ TNF⁺ CTL cells, appeared to be more frequent in HD ( Figure 3E ; 9.9% vs. 21% Mann–Whitney U test, ***p = 0.001). Conversely, the fraction of degranulating cells expressing TNF (TNF⁺ CD107a⁺ CD8 CTL cells) increased significantly in PWUD ( Figure 3E ; 9.6% vs. 5.9%; Mann–Whitney U test, *p < 0.05). Nevertheless, when we analyzed all permutations using the multiparameter SPICE visualization, we found that the polyfunctional response of the CD8 CTL cell fraction was not significantly different from HD and thus broadly preserved in PWUD ( Figure 3F ). Altogether, these data indicate that PWUD have a normal response to vaccination, but the frequency of some subpopulations differed from the healthy controls. Next, to further investigate the characteristics of the antigenic response to the vaccine in PWUD, we performed a direct comparison of the CD8 CTL cell response to the stimulation with 15-mer and 9/10-mer peptide pools (Peptivator mix vs. NOI pool). Biplots of T-cell responses based on the frequency of cell populations expressing different marker combinations revealed a high degree of linearity and a substantial overlap between the two cohorts in terms of response ( Figure 4 ). These results were replicated in a subgroup of donor using a simplified AIM assay ( Supplementary Methods and data not shown).

As observed for the general population, serum IgG anti-RBD level (BAU/mL) decayed with time after vaccination in PWUD ( Figure 5A ). Furthermore, we found no significant correlation between humoral and spike-specific CD4 Th cell response to the Peptivator mix ( Figure 5B ). Similarly, the humoral response did not correlate with CD8 CTL response, notwithstanding the type of stimulation ( Figure 5C ).