Participants were included from two clinical studies. The first clinical study comprised a cross-sectional observational study in Dutch symptomatic pertussis cases (Specifieke Kinkhoest Immuniteit; SKI; 2008–2012) (Han et al., 2013; van Twillert et al., 2014). This study was approved by the accredited Review Board STEG, followed by management of the METC UMC Utrecht (CCMO nr: NL16334.040.07). The second clinical study was a longitudinal observational study in Dutch symptomatic pertussis cases (Immfact; 2014–2020). This study was approved by the accredited Medical Research Ethics Committee “METC Noord-Holland” (Alkmaar, the Netherlands) followed by management of the METC MEC-U (Utrecht, the Netherlands) (CCMO nr: NL4679.094.13). All participants provided written informed consent for the collection of samples, the usage of a completed questionnaire regarding clinical symptoms and vaccination history, and the subsequent immunological analyses. Informed written consent for minor participants was provided by both parents or guardians of participants. These studies were conducted in compliance with the principles of the Declaration of Helsinki.

Participants from the SKI study consisted of 58 clinically symptomatic (ex) pertussis patients who donated blood at a single known, either early or late, time point after their laboratory confirmed diagnosis. For analysis, subjects were classified into a young group (median age of 12 years; range 11–15 years, n = 21), referred to as youngsters (Y), and an adult group referred to as (A) (median age of 38 years; range 25–56 years, n = 37). Youngsters and adults were subcategorized according to the time elapsed between their date of clinical pertussis diagnosis and date of blood sampling (τ, in months), indicated as sampled in early phase (τ1 < 4 months, or in late phase τ2 ≥ 8 months, no upper limit for inclusion). Late phase cases were excluded on serological criteria, if having an IgG plasma level specific for Pertussis Toxin (PT) ≥ 62.5 IU/ml, indicative for an additional non-diagnosed exposure (de Melker et al., 2000; de Greeff et al., 2010). Participants of the Immfact study consisted of 16 clinically symptomatic pertussis patients who donated blood at two longitudinal time points after their laboratory confirmed diagnosis. Here, youngsters (Y) with a whole cell priming background (median age of 16 years, range 12–23 years, n = 9) and older adults (O) (median age of 71 years, range 60–78, n = 7) were sampled at τ1 ≤ 3 months post diagnosis (early phase) as well as at 8 ≤ τ2 ≤ 12 months post diagnosis (late phase). For details of cohorts, see Supplementary Figure S1. All participants from both clinical cohorts and of all age groups were primed during their first year of life and according to their birth cohort with multiple doses of the Dutch whole cell vaccine (in use until the year 2005 when the Dutch National Immunization Programme switched to an acellular primary pertussis vaccine for infants). As by their birth cohort and the Dutch national immunization programme, participants did not receive any pertussis booster dose(s), except for three younger participants from the Immfact cohort. At the age of 4-years these participants received an acellular pertussis booster dose at school entrance, which was programmatically introduced in the Netherlands in 2001. The Dutch National Immunization Programme does not include further adolescent or adult acellular pertussis booster doses. Maternal pertussis immunization was only introduced in December 2019.

P.69 pertactin (PRN) was recombinantly expressed and purified from E. coli as described previously (Hijnen et al., 2005). PT and Filamentous Hemagglutinin (FHA) were both obtained from Kaketsuken, Japan. Presence of E. coli LPS or B. pertussis LOS was ruled out (<0.015 EU/ml) based on Limulus Amebocyte Lysate (LAL) testing. Pools of synthetic immunogenic Bp peptides were purchased; Bp132, a peptide pool consisting of 132 immunogenic peptides derived from PT, FHA, PRN and Fimbriae 2/3 (Bancroft et al., 2016) (Pepscan); PT peptide pool, consisting of 18-mers spanning the PT S1 subunit with 12 amino acids overlap (in-house synthesis, P. Hoogerhout); FHA peptide pool and PRN peptide pool, consisting of only the FHA or PRN-specific peptides from the Bp132 peptide pool, respectively (Pepscan).

Venous blood samples were collected in the SKI study using CPT Mononuclear cell preparation tubes (BD) and in the Immfact study using vacutainer blood collection tubes (BD). Peripheral blood mononuclear cells (PBMCs) were isolated using standard procedures and frozen in 10% DMSO. Samples were kept at −80°C overnight and then stored at −135°C.

PBMCs were quickly thawed at 37°C and in vitro cultured as described elsewhere (Schure et al., 2012) with minor modifications. Briefly, SKI PBMC samples were cultured for 5 days at 37°C, 5% CO₂, in 96-well U-bottom plates at 3.0 × 10⁵ viable cells in 150 µL/well in replicate wells per condition in AIM-V medium (Gibco, Invitrogen, United States) containing 5% human AB serum (Harlan, United Kingdom) (AIM-V+) only (negative control), or in the presence of 5 μg/ml of PT (heat-inactivated), FHA or PRN, or of 5 μg/ml of pokeweed mitogen (Sigma Chemicals, United States) (positive control). Alternatively, Immfact PBMC samples were labelled with CellTrace Violet (Thermofisher) at a final concentration of 0.5 µM and cultured for 6 days at 37°C, 5% CO₂, in loosely capped 5 ml tubes (Falcon, BD) at 2.0 × 10⁶ in 1 ml/tube in RPMI 1640 medium (Gibco, Invitrogen, United States) containing 5% human AB serum (Harlan, United Kingdom; RPMI+) in medium only (negative control) or in the presence of PT S1 peptide pool, Bp132, PRN peptide pool, FHA peptide pool (all at 0.1 µM per individual peptide) or anti-CD3 and CD28 antibodies (at 0.5 μg/ml and 1 μg/ml, respectively, positive control). For both SKI and Immfact samples aliquots of culture supernatants were collected on day 5 and stored at −80°C for subsequent quantification of cytokine levels. After culture, SKI and Immfact PBMCs were harvested on day 5 and day 6, respectively, for further analyses. Depending on the amount of PBMCs available per sample, one or more B. pertussis antigenic stimulations were tested apart from the negative control.

5-stimulated PBMC (SKI) samples were spun down and after collection of supernatants, cells were reconstituted in culture medium and transferred to anti-human IFNγ (Mabtech, Sweden) coated ELISpot plates (Millipore, United States) at a starting concentration of 1.4 × 10⁵ cells in 150 µL/well and three times two-fold serially diluted. Plates were incubated for approximately 20 h at 37°C, 5% CO₂, then after four washing steps and one cell-lysing step, incubated with 1 μg/ml anti-human IFNγ (Mabtech), followed by peroxidase labeled extravidin (Sigma) and BCIP/NBT (KPL, United States). After development, the plates were air-dried at room temperature prior to analysis of numbers of IFNγ spot forming cells (SFC), assessed by an automatic computer-assisted ImmunoScan-Pro reader (CTL Europe, Germany). Results are expressed as IFNγ SFC/100.000 PBMCs with a lower detection limit of 0.1 IFNγ SFC/100.000 PBMCs.

CellTrace-labelled day 0 and day 6-stimulated Immfact PBMCs were washed and labelled at 4°C for cell surface and intracellular markers with the following antibodies: anti-CD3-Alexa700 (clone SK7), anti-CD4-BV711 (clone OKT-04), anti-CD8-BV785 (clone RPA-T8), anti-CD45RO-PerCP-Cy5.5 (clone UCHL1), anti-FoxP3-Alexa647 (clone 259D), anti-PD-1-BV605 (clone EH12.2H7), anti-CD127-BV650 (clone A019D5), anti-CTLA-4-PE (clone L3D10), anti-TIGIT-PE/efluor-610 (clone MBSA43), anti-Helios-PE-Cy7 (clone 22F6), anti-CD27-BUV395 (clone L128), anti-CD25-BUV737 (clone 2A3) and Live-dead ZOMBIE NEAR IR-APC-Cy7. Acquisition was performed on a BD LSR Fortessa X-20 (BD Biosciences, Franklin Lakes, NJ, United States). Subsets of CD4⁺ T cells were identified based on combined marker expression on gated CD3⁺CD4⁺ cells. Frequencies and phenotypes of proliferating antigen-specific CD4⁺ T cells were determined by analyzing sequential halving of the CellTrace Violet fluorescence intensity combined with performing surface phenotypic and intracellular marker labelling. Percentages of proliferated CD4⁺ T cells found after medium stimulation conditions were subtracted from percentages of proliferated CD4⁺ T cells after Bp-antigen stimulation, when comparing proliferative CD4⁺ T-cell responses between groups. For both replication index calculations (by proliferation modeling) as well as gating analyses, FlowJo software was used (Tree Star, Ashland, OR, United States).

Concentrations of the cytokines IL-2, IFNγ, TNF-α, IL-5, IL-13, IL-17A and IL-10 in day 5 culture supernatants were determined in pg/mL according to manufacturer’s instructions using a commercial multiplex bead-based immunoassay kit (Bio-Rad, United States) (SKI samples) or using the LEGENDplex Human Th cytokines kit (BioLegend) (Immfact samples), and expressed in [pg/ml]. Measurements and data analysis were performed with a Bio-Plex 200, using Bio-Plex Manager software or flow cytometry (FACS Canto II), respectively.

Dimensionality-reduced analyses (viSNE) of flow cytometry data were performed in Cytobank (www.Cytobank.org). Prior to the viSNE plots, CD4⁺ proliferating events were exported and pooled per culture condition, age group and timepoint. The number of cells included in the analysis was 2.3 × 10⁵ and was equal for youngsters and older adults.

For data analysis and visualization of data, GraphPad Prism (GraphPad Software version 8.4.1) was used. Statistical significance of differences was analyzed with the nonparametric Mann-Whitney t-test, when comparing stimulated and unstimulated conditions of samples, and the Wilcoxon matched pairs signed-rank test, when comparing longitudinal samples from individuals. For all analyses, p values <0.05 were considered statistically significant.

Production of IFNγ is an important parameter for an effective immune response to a Bp infection (Mills et al., 1993; Ryan et al., 1997; Hafler and Pohl-Koppe, 1998). We first set out to explore IFNγ production in total PBMC cultures stimulated with Bp protein antigens to compare these responses between youngsters and adults. To determine the impact of age on functionality of Bp-specific responses, we studied IFNγ responses in early and late phase post-infection PBMC samples from youngsters and adult pertussis patients of the SKI cohort with a similar whole cell pertussis vaccine priming background. Production of IFNγ of 5-day cultured PBMCs stimulated with protein antigens Pertussis Toxin (PT), filamentous hemagglutinin (FHA) and pertactin (PRN) were determined by ELISpot. Geomean number of IFNγ SFC ranged between 11–118 per 1 × 10⁵ PBMCs, depending on the stimulation, age and timepoint. The frequency of IFNγ-producing cells in response to PT was significantly lower in adults compared to youngsters in the early phase (Figure 1A). Waning of the PT response from the early to late phase was observed in youngsters but not in adults. In the early phase response to FHA, adults showed comparable median frequencies of IFNγ-producing cells (Figure 1A), which did not decline over time. In response to PRN, no age-related differences in numbers or patterns over time were found for IFNγ-producing cells (Figure 1A). Similar to findings on the number of IFNγ-producing cells, the level of IFNγ measured in culture supernatants was significantly lower in PT-stimulated samples of adults compared to youngsters in the early phase of the response (Figure 1B). In summary, these data suggest age-related differences in IFNγ responses to PT in the early phase after clinical pertussis infection.

Next, we extended our analyses of cytokines produced in total PBMC cultures stimulated with protein antigens in youngsters and adults from the SKI cohort by measuring Th1 (IL-2, TNF-α), Th2 (IL-5, IL-13) (Figure 2), Th17 (IL-17) and regulatory (IL-10) associated cytokines (Supplementary Figure S2). Overall, adults presented with lower levels of cytokines than youngsters in both early and late phase after clinical pertussis infection. We found significantly lower levels of Th1-associated cytokines IL-2 and TNF-α, and Th2-associated cytokines IL-5 and IL-13 in PT-stimulated culture supernatants in adults compared to youngsters in the early phase (Figure 2). In FHA-stimulated samples, significantly lower levels of Th2 associated IL-5 and IL-13 cytokines were found in the late phase in adults compared to youngsters (Figure 2). In addition, PRN-stimulated culture supernatants had significantly lower levels of IL-5, IL-13 and TNF-α in the late phase. Levels of IL-17A and IL-10 were comparable between the two age groups (Supplementary Figure S2). Together, these data indicate that in various phases after clinical pertussis infection, Bp specific cytokine responses, typically mediated by CD4⁺ T cells, are reduced at older age.

Longitudinal follow up confirms lower levels of Th-type cytokines after Bp infection in older adults The lower Bp-antigen specific cytokine responses found in the older age group suggested that either the proportion of Bp-antigen specific functional memory CD4⁺ T cells responding in this group was smaller, or its proliferative capacity was impaired compared to youngsters. Immunosenescence of CD4⁺ T-cell proliferation has been described at increased age (Jiang et al., 2007). To shed more light on the mechanism, we designed a flow-based assay, allowing the analysis of Bp antigen-specific proliferation of CD4⁺ T cells, as well as exploring several cellular markers linked to T-cell immunosenescence. Here, we selected paired samples obtained at fixed time points in the early and late phase post clinical pertussis from whole cell vaccine primed youngsters and older adults in the Immfact study (Supplementary Figure S1). In contrast to stimulations with whole protein antigen, we now used peptide pools of separate and combined Bp antigens of which the epitopes have previously been described as immunogenic and optimal for binding to MHC class II (Bancroft et al., 2016) and therefore allowed us to assess the response of antigen-specific CD4⁺ T cells. First, we compared cytokine production in 5 days culture supernatants of PBMCs from youngsters and older adults post clinical pertussis infection longitudinally (Figure 3, Supplementary Figure S3). In older adults, IFNγ and IL-13 levels were significantly lower in response to peptide pools Bp132, PT S1 and FHA in the late phase. In addition, lower levels of TNF-α were detected in response to Bp132 (in the late phase) and FHA (in the early phase). Stimulation with PRN peptide pools revealed lower levels of IL-13 in older adults in the early phase. Thus, despite various differences in study design and experimental approach between the SKI and Immfact samples, we observed a similar trend towards lower Th-type cytokine production in response to Bp antigens with increasing age.

We then explored CD4⁺ T-cell populations of the Immfact PBMC samples for the various flow-cytometric read-outs of our assay. To first assess whether youngsters and older adults of the Immfact cohort display differences in the frequencies of global naïve and memory CD4⁺ T-cell subsets in their peripheral blood, we analysed the CD4⁺ T-cell composition of PBMC samples prior to stimulation based on expression of CD27 and CD45RO (Figure 4A, gating strategy in Supplementary Figure S4). We found comparable percentages of central memory (T_cm, CD45RO⁺CD27⁺), effector memory (T_em, CD45RO⁺CD27⁻) and terminally differentiated (T_emra, CD45RO⁻CD27⁻) subsets between the age groups at both early and late phase, only the percentage of naïve CD4⁺ T cells was significantly reduced in older adults compared to youngsters in the late phase (Figure 4B). After 6-day stimulation with Bp peptide pools Bp132, PT S1, FHA and PRN, we found the global composition of the CD4⁺ T-cell compartment to be comparable between youngsters and older adults and between early (Figure 4C) and late phase samples (Figure 4D).

Additionally, regulatory T cells (Tregs) have been suggested to accumulate with increasing age (Gregg et al., 2005; Lages et al., 2008; Elyahu et al., 2019). Therefore, we analysed the frequency of Tregs, defined as either CD127^-CD25⁺ or FoxP3⁺CD25⁺ CD4⁺ T cells prior to stimulation (Seddiki et al., 2006; Yu et al., 2012; Rodríguez‐Perea et al., 2016). Tregs were in the range of 5–8% of the CD4⁺ T cells and frequencies were comparable between youngsters and older adults (Supplementary Figure S5. Altogether, these analyses show that youngsters and older adults from the Immfact cohort do not differ in the proportions of memory T-cell subsets (based on both phenotypes) before and after Bp-peptide stimulation, nor in ex vivo Tregs.

Next, we investigated whether age had an impact on proliferative responses of antigen-specific CD4⁺ T cells in youngsters and older adults. To determine frequencies of proliferating CD4⁺ T cells, PBMCs were CellTrace labelled, stimulated with Bp antigen peptide pools for 6 days, and assessed for CellTrace Violet diminution (Figure 5A). Our data showed a general trend of lower frequencies of CD4⁺ T cells proliferating to Bp antigen in the older adults, which was significant for the Bp132 and PT S1 peptide pool early after clinical pertussis infection (Figure 5B). In the late phase, no age-related differences were found. We did observe waning of proliferative responses in youngsters in time, as frequencies of proliferated CD4⁺ T cells to Bp132 and PT S1 peptide pools were significantly lower in the late phase post infection compared to the early phase (Figure 5B). Patterns of Bp-specific CD4⁺ T-cell proliferation in older adults in response to FHA or PRN were comparable to those observed in youngsters. The proliferative response was further investigated by calculating the replication index (fold-expansion of proliferating cells). The replication index was comparable between youngsters and older adults (Figure 5C), suggesting that responding CD4⁺ T cells undergo equal rounds of cell divisions in adults compared to young individuals. Thus, these data show that the magnitude of the proliferative response of the CD4⁺ T-cell compartment to Bp antigens Bp132 and PT S1 declined with age but rounds of divisions in response to all Bp antigens per cell were similar. This suggests maintenance of a smaller memory pool of Bp-specific CD4⁺ T cells for recall responses as a hallmark of immunosenescence.

Next, we assessed whether lower cytokine production and a lower proliferative response were accompanied by an altered phenotypical profile of responding CD4⁺ T cells. We hypothesized that antigen-specific, proliferating CD4⁺ T cells of older adults express higher levels of co-inhibitory receptors CTLA-4, PD-1, and TIGIT and lower levels of the activation marker CD25 (the IL-2-alpha receptor). In contrast, we found that expression of CTLA-4 by proliferated CD4⁺ T cells in the early phase in youngsters was significantly higher compared to older adults in response to Bp132, whereas the percentage of CTLA-4⁺ proliferating CD4⁺ T cells varied but was comparable between the two age groups (Supplementary Figures S6A,B). Expression of the activation marker CD25 on proliferated CD4⁺ T cells was comparable between youngsters and older adults both in the early and late phase, as well as were frequencies of CD25⁺ proliferated CD4⁺ T cells, both in the early and late phase (Supplementary Figures S6C,D). Likewise, analysis of expression and frequencies of PD-1⁺ (Supplementary Figure S7A–B) and TIGIT⁺ (Supplementary Figure S7C–D) cells within proliferating CD4⁺ T cells did not reveal age-related differences.

Lastly, we applied dimensionality reduction analyses (viSNE) to proliferated CD4⁺ T cells of pooled datafiles of youngsters and older adults in the early phase to reveal any differences based on combined marker expression between proliferated CD4⁺ T cells of youngsters and older adults (Figures 6A,B). Cluster analysis showed a potentially interesting cluster, cluster d, that was significantly lower in older adults compared to youngsters (Figure 6C). Based on subsequent heatmap visualization of marker expression in the clusters (Figure 6D), this cluster was positive for markers including CD25, CTLA-4 and TIGIT and showed somewhat higher Helios expression compared to other clusters. Additionally, we applied the gating approach derived from cluster analysis of proliferated CD4⁺ T cells after pneumococcal antigen stimulation using a similar phenotypical marker panel (He et al., unpublished) and found significant higher proportion of CD4⁺ T cells that were CD127^-FoxP3^-Helios⁺ (Figure 6E). Helios is considered a marker associated with (CD4⁺) T-cell activation (Akimova et al., 2011; Bengsch et al., 2018). In summary, unbiased viSNE cluster analysis of Bp-specific CD4⁺ T cells that have proliferated did not reveal phenotypical differences between youngsters and older adults based on the expression of co-inhibitory receptors CTLA-4, PD-1, and TIGIT or of activation marker CD25. The significantly higher frequency of CD127^-FoxP3^-Helios⁺ proliferated CD4⁺ T cells in youngsters may however suggest a reduced activation status at older adult age. Altogether, these data indicate that older adult Bp cases show features of CD4⁺ T-cell Bp-specific immunosenescence following clinical infection, highlighted by significantly lower cytokine and proliferative responses than younger adolescent counterparts. Moreover, absence of age-related differences in fold-expansion and co-inhibitory marker expression of Bp-specific proliferated CD4⁺ T cells, with a significant reduction of an activated CD4⁺ T-cell subset at older age, suggest that the immunosenescence can largely be attributed to a reduced size and thereby responsiveness of the maintained Bp-specific memory CD4⁺ T-cell pool.