Successive annual influenza vaccination induces a recurrent oligoclonotypic memory response in circulating T follicular helper cells

Highly activated phenotype of ICOS+CD38+ cTfh at baseline

We first reasoned that recently activated Tfh in the circulation might be identified by the expression of proteins associated with cellular activation. We focused on cTfh, defined as non-naive CD4+CXCR5+PD-1+, that: expressed high levels of the costimulatory marker ICOS that is expressed by lymphoid Tfh (1), contributed to in vitro B cell help (3), and were induced in cTfh following influenza vaccination (5, 6). Of circulating cTfh at baseline, ~5% expressed ICOS and also CD38 (hereafter referred to as ICOS+CD38+ cTfh and compared to ICOS−CD38− cTfh). These ICOS+CD38+ cTfh expressed multiple other markers of cellular activation. Directly ex vivo, ICOS+CD38+ cTfh had higher expression of CD25, CD27, CD28, CTLA4, PD-1, Helios, and Ki67 and lower expression of CD127 than ICOS−CD38− cTfh (Figure 1A–C). Higher expression of CD3 and CD4 were also observed in ICOS+CD38+ cTfh compared to ICOS−CD38− cTfh, but CXCR5 expression was similar (Figure S1A). This pattern of activation markers was comparable in cryopreserved and freshly isolated PBMC (data not shown). High expression of CD25, CD27, CD39, CTLA4, and Helios was observed even in the absence of Foxp3 (Figure S1B). In sum, ICOS+CD38+ cTfh were a small fraction of all cTfh but expressed many proteins associated with T cell activation.

Activated cTfh express Bcl6 and can produce IL-10, IL-17, and IL-21

Bcl6 has a central role in the Tfh program (1) but protein expression was not detected in total peripheral blood cTfh in prior studies (2, 3, 5). To address whether ICOS+CD38+ cTfh may resemble lymphoid Tfh more than other cTfh, we evaluated expression of Bcl6 protein in the ICOS+CD38+ subset, as well as the expression of other key Th transcription factors and cytokine production. In peripheral blood, Bcl6 protein and mRNA transcript expression was highest in ICOS+CD38+ cTfh compared with ICOS−CD38− cTfh or CXCR5− memory CD4 T cells but was lower than in splenic germinal center Tfh (Figure 2A, Figure S2A–D), suggesting that the ICOS+CD38+ cTfh subset bear closer resemblance to lymphoid Tfh than ICOS−CD38− cTfh. Moreover, expression of Bcl6 was highest in the ICOS+CD38+ cTfh with the highest PD-1 expression (Figure 2B).

Prior studies have suggested Th1-, Th2-, and Th17-polarized subsets exist within the cTfh pool (2), but other studies have found low expression of master transcriptional regulators such as Tbet or RORγT in Tfh (22, 23). To address whether cTfh expressed these regulators, protein expression of these transcription factors was profiled in cTfh. The majority of activated cTfh lacked detectable expression of any of these lineage-associated transcription factors (Figure 2C, Figure S2E). Of these, Tbet was most commonly expressed in ICOS+CD38+ cTfh. T follicular regulatory cells (Tfr) were identified by expression of Foxp3 and CD25 and represented approximately 5% of the ICOS+CD38+ cTfh, consistent with the frequency of circulating Tfr from mouse studies (24). The transcription factor c-Maf has a critical role in Tfh biology (25–27) and displayed high expression in ICOS+CD38+ cTfh, as did total Foxo1. GATA3 is critical for development, maintenance, and function of lymphocytes (28) and was higher in ICOS+CD38+ cTfh compared to ICOS−CD38− cTfh (Figure 2C). Coexpression of Tbet and RORγT or GATA3 was observed in a minority of ICOS+CD38+ but not ICOS−CD38− cTfh (Figure 2D, Figure S2F). Thus, the ICOS+CD38+ cTfh subset expressed Bcl6 and c-Maf protein, but few cells expressed Tbet, RORγT, or Foxp3.

To investigate the cytokine profile of cTfh, we next performed ex vivo stimulation and intracellular cytokine staining for cytokines associated with T helper subsets and important for germinal center reactions (1), including IL-17 (29) and IL-10 (30). Upon stimulation with PMA/ionomycin, ICOS+CD38+ cTfh produced a unique combination of IL-10, IL-17, and IL-21 compared to other CD4 subsets (Figure 2E, Figure S2G–I). Similar findings were observed when PBMC were stimulated with SEB (data not shown). Of note, ICOS+CD38+ cTfh were the strongest producers of IL-21 compared to other subsets. Indeed, the frequency of IL-21-producing cells in the ICOS+CD38+ cTfh subset was two-fold higher than the frequency in the ICOS−CD38− cTfh and three-fold higher than CXCR5− memory CD4. Overall, ~1/3 of ICOS+CD38+ cTfh expressed none of the assayed cytokines, ~1/3 expressed TNFα alone, and the remainder co-expressed multiple cytokines, primarily IL-10 and IFNγ (Figure 2F, Figure S2H). Most IL-17⁺ ICOS+CD38+ cTfh also produced IFNγ and TNFα. There was greater polyfunctionality in the ICOS+CD38+ cTfh than ICOS−CD38− cTfh, consistent with their activation state. While the majority of ICOS+CD38+ cTfh were capable of production of TNFα, this subset produced more IL-21, supporting the hypothesis that ICOS+CD38+ cTfh are more similar to lymphoid Tfh than ICOS−CD38− cTfh.

Influenza-specific cells can be identified within ICOS+CD38+ cTfh subset following influenza vaccination

We and others previously observed increased expression of ICOS among cTfh following influenza vaccination (5, 6). We next wanted to investigate whether protein expression of other markers of activation changed after immunization. Subjects in Cohort 1 (Table S1) received inactivated influenza vaccine during the Fall of 2014 (n=28). In an influenza challenge study, a subset of CD4 coexpressing CD38-Ki67 was observed one week after infection (13). In our cohort one week after vaccination we observed increased CD38 expression among nonnaive Ki67+ CD4 T cells (Figure 3A; Figure S3A). Prior to vaccination, cTfh represented ~25% of the Ki67+CD38+ CD4 T cells. Following vaccination, the proportion of cTfh in this Ki67+CD38+ CD4 T cell compartment increased between 4%–60% (Figure 3A–B). By instead gating directly on nonnaive CXCR5+PD-1+ CD4, we observed a vaccine-induced increase of 20%–60% in this subset of ICOS+CD38+ cTfh (Figure 3C, Figure S3B). Expression of PD-1 also increased in the ICOS+CD38+ cTfh following vaccination (Figure S3C–D), but no changes were detected with respect to expression of CD25, CD27, CD28, CD127, CTLA4, Helios, or Ki67 (data not shown). Of the nonnaive Ki67+CD38+ CD4 cells that were CXCR5−, there was no clear phenotypic change following influenza vaccination (data not shown). Thus, influenza vaccination was associated with increased frequency of the ICOS+CD38+ cTfh population.

We next hypothesized that the day 7 ICOS+CD38+ cTfh population would be enriched for influenza-specific cells. Prior studies have identified CD4 T cells specific for nucleoprotein (NP), matrix 1 (M), and hemagglutinin (HA) peptides following vaccination (13, 14, 31). However, these methods typically rely on cytokine production after stimulation. We observed production of TNFα, IFNγ, or IL-2 by only a minority of cTfh following stimulation with overlapping peptide pools for influenza proteins (HA-1, HA-3, NP, and M, data not shown), and IL-21 was produced by only a subset of cTfh following PMA/ionomycin stimulation (Figure 2E). Here, an independent approach using class II tetramers was employed (32). A second cohort of subjects was recruited (n=12, Table S1) and a similar ICOS+CD38+ cTfh response was observed one week after vaccination (data not shown). We examined HLA class II/peptide tetramer+ CD4 T cells at one week following vaccination in two HLA-DRB1*04:01 subjects. CD4 T cells specific for either HA residues 306–318 (HA_306–318) or HA residues 398–410 (HA_398–410) (12, 32, 33) were identified in these subjects, including clear HA-specific cTfh (Figure 3D). Of these HA tetramer+ cTfh, 50–91% co-expressed ICOS and CD38. Following influenza vaccination the majority of these tetramer-positive cTfh also expressed CXCR3 (Figure S3E), consistent with prior reports (6). Thus, the identification of influenza HA-specific cells in the cTfh subset one week after inactivated influenza vaccine suggests that antigen-specific cTfh were indeed induced by vaccination and could be found in the ICOS+CD38+ cTfh subset.

Although tetramer staining clearly demonstrated that influenza-specific CD4 T cells are contained in the ICOS+CD38+ cTfh population following vaccination, a tetramer-based approach is likely to underestimate the total influenza vaccine-induced population as few epitopes were interrogated. Expression of activation-induced markers (AIM) has been used to identify antigen-specific CD4 T cells following antigen stimulation (34, 35). This approach has the advantage of not being restricted to a single epitope specificity and offers a broader view of antigen-specific cTfh populations. Thus, to complement the tetramer approach, PBMC pre and post vaccination were stimulated with overlapping influenza peptide pools for NP, M, H1, and H3 for 18 hours. After stimulation of PBMC from day 7 post vaccination, we observed a clear population of antigen-specific ICOS+CD38+ cTfh identified by upregulation of CD69 and CD200 (Figure 3E–F). This population was not present in unstimulated controls (Figure S3F) and was 30- to 150-fold higher at day 7 post vaccination compared to day 0 in the ICOS+CD38+ population or compared to the ICOS−CD38− cTfh subset (Figure 3E–F, Figure S3F). We did not observe an increase in the overall frequency of ICOS+CD38+ cTfh following 18-hour stimulation (data not shown), suggesting that acquisition of ICOS and CD38 either requires a longer period of stimulation, stronger stimulus, or perhaps other signals. Together, these data demonstrate the presence of robust influenza-specific responses within the ICOS+CD38+ cTfh population at day 7 after influenza vaccination.

Increased clonality of ICOS+CD38+ cTfh following influenza vaccination

Compared to other CD4 T cell populations, the ICOS+CD38+ cTfh subset was enriched for influenza-specific cells based on tetramer and response to influenza antigen stimulation in vitro (AIM analysis). To examine the clonotypes in the vaccine response more broadly, T cell receptor sequencing (TCRseq) from multiplex PCR amplification of genomic DNA (20) was employed on sorted cTfh populations. In most subjects, clonality of the ICOS+CD38+ cTfh population increased following influenza vaccination (Figure 4A–B, Figure S4A–B). The ICOS−CD38− cTfh had reduced clonality following vaccination but this did not correlate with the increase in clonality in ICOS+CD38+ cTfh (Pearson r=−0.16) (Figure S4A). The day 7 response in ICOS+CD38+ cTfh was typified by the small but clear increase of many clonotypes, rather than dominance of a single expanded clonotype. Another measure of clonal dominance, the Gini Index, correlated strongly with clonality and corroborated these results (Figure S4C–D). Correlations were observed between the change in frequency of the ICOS+CD38+ cTfh population and clonality, both when examining day 7 alone as well as when the fold-change at day 7 vs day 0 was examined (Figure 4C and Figure S4E). We did not detect a clear relationship with the hemagglutination-inhibition antibody titer and ICOS+CD38+ cTfh clonality (Figure S4F–G), perhaps reflecting the fact that hemagglutination-inhibition antibody is only a small fraction of the total antibody induced by vaccination. However, the change in clonality of ICOS+CD38+ cTfh did correlate with the change in frequency of circulating plasmablasts (Figure 4D–E), suggesting a relationship between this cTfh subpopulation and the humoral immune response. These data demonstrate a rapid, oligoclonal ICOS+CD38+ cTfh response to vaccination that strongly correlated with the magnitude of the response. Influenza vaccination resulted in antigen-specific expansion of cTfh populations that could be observed in the PBMC and had a connection to the overall B cell response.

Repeated influenza vaccinations elicit a recurring oligoclonal response

Nearly all humans become seropositive for influenza virus specific antibodies by the end of childhood (36). Thus, we assumed that, prior to entry into the study, all adult subjects had either been immunized against, or been infected with, influenza A virus at least once in the past. However, it was unknown whether a recall response with successive vaccinations would elicit the same oligoclonal response, given the passage of time, the annual reformulation of the influenza vaccine, and the possibility of influenza infection prior to re-vaccination. For six of the subjects in Cohort 2, cryopreserved PBMC were available following influenza vaccination over multiple years. Of these, four had PBMC available for the 2014–2015 and 2015–2016 vaccination years and two had PBMC available for the 2013–2014, 2014–2015 and 2015–2016 vaccination years. Repeated induction of ICOS+CD38+ cTfh was observed at day 7 following each successive vaccination (Figure 5A and Figure S5A).

We first investigated the stability of the cTfh repertoire over time, independent of vaccination. The overlap score (Materials and Methods) provides an estimate of similarity in repertoire of a population of T cells across different time points. To assess this, similarity of the TCRB repertoire of each subset was compared to itself one year post vaccination (i.e. just prior to re-vaccination). The ICOS+CD38+ cTfh subset had low overlap score over 1 year in the same individual suggesting high turnover of this subset of cTfh in the absence of vaccination (Figure 5B). In contrast, the ICOS−CD38− cTfh and CXCR5− memory CD4 subsets had more stable TCR repertoires over time. Among these populations, the ICOS−CD38− cTfh subset had more overlap across one year than the CXCR5− memory subset. These observations indicate more dynamic turnover in the clones contained in the ICOS+CD38+ subset compared to the ICOS−CD38− subset of cTfh in the steady state.

We next considered the effect of vaccination on the TCRB repertoire of these CD4 T cell populations. There was essentially no overlap when comparing the ICOS+CD38+ repertoire at Year 1 Day 7 post vaccination to Year 2 Day 0, indicating that, within the limit of detection of these assays, the clones present in the ICOS+CD38+ cTfh subset following influenza vaccination were no longer detectable in this subset a year later (Figure 5C). However, on day 7 post vaccination in Year 2, clones from the Year 1 post vaccination ICOS+CD38+ cTfh subset re-emerged in the ICOS+CD38+ cTfh subset. Neither the ICOS−CD38− cTfh nor CXCR5- memory CD4 subsets showed such a response (Figure 5D and Figure S5B). The timing of the vaccination in the prior years did not have an apparent effect on the recall responses in ICOS+CD38+ cTfh (Figure S5B). The median number of clonotypes in the recurrent oligoclonal response for all subjects was 66 but varied widely from subject to subject and did not appear dependent on number of years of observation (Figure 5E). These data strongly suggested a recurrent oligoclonal response in the ICOS+CD38+ cTfh subset to repeated influenza vaccination in all subjects.

To investigate these clonality relationships in more detail, we focused on the two subjects for whom three years of data were available. There was a robust increase in overlapping clonotypes between Year 1 Day 7 and subsequent Day 7 time points, but we observed only weak overlap with subsequent Day 0 time points in the ICOS+CD38+ cTfh subset (Figure 5F and Figure S5B–C). Moreover, clones shared at every Day 7 time point in ICOS+CD38+ cTfh were absent at every Day 0 time point, whereas clones shared at every Day 7 time point in ICOS−CD38− cTfh or CXCR5− memory were sometimes present at Day 0 time points in these subsets as well (Figure 5G and Figure S5D) suggesting more stability in the clonality of the latter subsets. Rare clones in the ICOS−CD38− cTfh and CXCR5− subsets demonstrated the same pattern as the ICOS+CD38+ cTfh subset and increased in frequency after vaccination, suggesting possible influenza vaccine responses in these subsets. For clones shared at every Day 7 time point in ICOS+CD38+ cTfh, there was no quantitative change in the median frequency at each Day 7 time point, arguing against successive quantitative boosting with each vaccination (Figure S5E). Finally, a cumulative frequency was obtained by adding together all of the frequencies for the Year 1 Day 7 clonotypes in the Year 2 Day 0 time point. The cumulative frequency of the recurrent oligoclonal response clones ranged from 6–15% of the total ICOS+CD38+ cTfh subset (Figure 5H). Together, these data demonstrate considerable clonal dynamics in the ICOS+CD38+ subset of cTfh, suggesting that this subset contains transient clones responding to recent antigenic stimulation. Moreover, these observations support the idea that influenza vaccination induces a repeated recall of conserved clonotypes in the ICOS+CD38+ cTfh following each yearly vaccination.

Influenza-specific clonotypes are contained within the recurring oligoclonal response

The increase in circulating frequency of ICOS+CD38+ cTfh after influenza vaccination was observed consistently in nearly every subject. We hypothesized that clonotypes identified by tetramer or AIM strategies should thus be enriched in the ICOS+CD38+ cTfh subset at one week after vaccination. To test this idea, we sorted HA_308–316 or HA_398–410-positive CD4 T cells at 6 months post vaccination. At this time point, none of the tetramer+ CD4 T cells were ICOS+CD38+ (Figure 6A–B, Figure S6A). We then sequenced TCRB changes of these cells and compared the TCRB sequences to those previously obtained from the total ICOS+CD38+ cTfh on day 7 post vaccination. Despite only obtaining only a small number of TCRB sequences (n=10–35) due to the low frequency of tetramer+ CD4 T cells, there was clear overlap of the tetramer-specific clonotypes with the bulk ICOS+CD38+ cTfh subset at the day 7 time point each year for both tetramers for one of the two subjects examined (Figure 6A, Figure S6B). Even when only a single HA_398–410 clonotype in Subject 999 was identified within the recurring ICOS+CD38+ cTfh oligoclonal response, this finding was highly unlikely by chance alone (p=0.0002 by Fisher’s Exact test). The tetramer clonotypes together comprised 0.37–0.70% of the ICOS+CD38+ cTfh population at day 7 after vaccination each year for Subject 999 (Figure 6B). The inability to detect shared clones between tetramer+ CD4 T cells and the ICOS+CD38+ cTfh subset in Subject 108 (Figure S6C) was likely due to the low number of tetramer events that could be captured 6 months after vaccination in this subject, suggesting either involvement of other specificities in the cTfh response, or that capturing only 10–30 TCRB sequences for a single epitope specificity may be at the limit of detection for this type of analysis.

To further explore these questions and test whether the ICOS+CD38+ cTfh population contained influenza-specific cells in a manner that did not depend on HLA-class II tetramers, we next employed the AIM approach described above. As above, PBMC were stimulated with pools of influenza peptides and CD69+CD200+ICOS+CD38+ cTfh were sort purified on day 7 after influenza vaccination. Using this approach we identified 33–216 clonotypes by TCRseq, depending on the subject (Figure S6D). Many TCRB sequences were identified in the influenza-specific AIM approach that were shared with TCR sequences identified in the total ICOS+CD38+ cTfh subset at day 7 post vaccination of each year (Figure 6C, Figure S6E). Again similar to the recurring oligoclonal response, the AIM clonotypes showed similar pattern of increased frequency in the ICOS+CD38+ cTfh subset at each day 7 time point in three of the six subjects (Figure 6D). The overlap between the recurrent oligoclonal response and the AIM clonotypes was statistically significant in four of six subjects (Figure 6E, Figure S6F). Of note, AIM clones and recurring oligoclonal response clones were not prominently found in other subsets, suggesting strong enrichment for influenza specific clones in the cTfh compartment (Figure S6G–H), though because the non-cTfh compartments are substantially larger, undersampling cannot be excluded. Nevertheless, these data confirm the presence of influenza-specific clonotypes within the ICOS+CD38+ cTfh subset one week after each influenza vaccination, as well as within the yearly recurring oligoclonal response.

ICOS−CD38− cTfh may serve as a long-term reservoir of cTfh memory

The TCRB sequence has been used as a “fingerprint” to track clones through different compartments in other studies (37–39), although a particular clone could sometimes be found in multiple distinct CD4 subsets (40). Here, to track clones across subsets over time, clones present for each subset at Year 1 Day 7 were compared against all the other subsets at Year 2 (Figure 7A–C and Figure S7A). A cumulative frequency was obtained by adding together all of the frequencies for the Year 1 Day 7 clonotypes in the Year 2 Day 0 time point. As expected, clones that were originally present in the CXCR5− memory or ICOS−CD38− cTfh subsets in Year 1 Day 7 could be found in the CXCR5− memory (Figure 7A) or ICOS−CD38− cTfh (Figure 7B) subsets at Year 2 Day 0. Clones present in the ICOS−CD38− cTfh in Year 1 Day 7 were most frequently found in the ICOS−CD38− cTfh subset at Year 2 Day 0 (Figure 7B), consistent with the higher overlap scores for ICOS−CD38− cTfh at one year in Figure 5B.

Given the relative stability and greater circulating frequency of the ICOS−CD38− cTfh subset compared to ICOS+CD38+ cTfh, we hypothesized that ICOS−CD38− cTfh represented a long-term “reservoir” for cTfh from which specific clones may be recalled into the ICOS+CD38+ cTfh pool. To test this idea, we asked if clones present at Year 1 Day 7 in the ICOS+CD38+ cTfh could be found in other subsets at Year 2. Indeed, clones present at Year 1 Day 7 in ICOS+CD38+ cTfh were most frequently found in the ICOS−CD38− cTfh subset at Year 2 Day 0 (Figure 7C). Similarly, ICOS+CD38+ cTfh clones present at Year 2 Day 7 were most frequently found in ICOS−CD38− cTfh at Year 3 Day 0 (Figure 7D, Figure S7B). As noted earlier, influenza vaccination induced an increase in the frequencies of the Year 1 Day 7-matched clonotypes in the ICOS+CD38+ cTfh (Figure S7A), but this effect was not seen in CXCR5− memory or ICOS−CD38− cTfh. Some clones from Year 1 Day 7 ICOS−CD38− cTfh were present in Year 2 Day 0 ICOS+CD38+ cTfh, suggesting some low-level conversion of ICOS−CD38− cTfh to ICOS+CD38+ cTfh.

Together, these data indicate a strong clonal connection between ICOS+CD38+ and ICOS−CD38− cTfh. Clones expanded following influenza vaccination may, therefore, preferentially convert into the ICOS−CD38− cTfh subset. These results support a model where cTfh cycle back and forth between activated and inactive states following repeated antigen exposure, with the ICOS−CD38− cTfh subset representing the reservoir for memory cTfh.

Human subjects

Subjects were eligible if they were community-dwelling and had not received influenza vaccine in the prior 6 months; they were excluded if they had contraindications to influenza vaccine, active substance abuse, HIV/AIDS, clinically active malignancy, immunomodulatory medication need (i.e. chemotherapy, corticosteroids), or active illness (i.e. active respiratory tract infections). Seasonal inactivated influenza vaccine (Fluvarix, GlaxoSmithKline) was administered and peripheral venous blood was drawn on days 0, 7, and 28 after vaccination. For Cohort 1, study subjects were recruited and consented in the Fall of 2014 at the Clinical Research Unit at Duke University Medical Center (Durham, NC, USA), in accordance with the Institutional Review Boards of both Duke University and the University of Pennsylvania (Philadelphia, PA, USA). Blood was collected into heparinized tubes and shipped overnight to Philadelphia, PA. For Cohort 2, study subjects were recruited and consented in between 2013 and 2015 at the University of Pennsylvania. Samples from 2013–2014 were cryopreserved whereas 2015 samples were used immediately after collection. Human spleen samples from relatively healthy adults were obtained as de-identified excess medical tissue via the Cooperative Human Tissues Network (CHTN), typically after trauma or incidental splenectomy. Splenocytes were obtained by mechanical dissociation and cryopreserved until needed. All human subjects research was performed in accordance with the relevant Institutional Review Boards.

Flow cytometry

Fresh PBMC and plasma were isolated using Ficoll-Paque PLUS (GE Healthcare) or SepMate isolation (Stem Cell Technologies) and stained for surface and intracellular markers. Permeabilization was performed using the Foxp3 Fixation/Permeabilization Concentrate and Diluent kit (eBioscience). Antibodies and clones are described in Table S2. Cells were resuspended in 1% para-formaldehyde until acquisition on a BD Biosciences LSR II cytometer and analyzed using FlowJo (Tree Star) and ViSNE (Cytobank). Fluorescence-minus-one controls were performed in pilot studies. The Bcl6 protein expression analysis was performed on both a BD LSR II (18-color instrument) and also repeated using a BD Symphony A5 (28-color) cytometer. The advantage of the latter experiment was that the Bcl6 antibody was the only reagent used on the violet laser greatly reducing fluorescence spillover from other fluorochromes and substantially improving the signal-to-noise ratio for detection of low protein amounts.

HLA Class II tetramers

HLA Class II tetramers were prepared as previously described (32). Staining by tetramers against hemagglutinin residues 306–318 (PKYVKQNTLKLAT) or hemagglutinin residues 398–410 (SVIEKMNTQFTAV) in two HLA-DRB1*04:01 subjects was performed for 1 hour at room temperature, followed by magnetic bead enrichment at 4°C and fixation. Fixed samples were acquired on a BD Biosciences LSR II cytometer. For sorting experiments, tetramer staining was performed in a similar fashion but samples were maintained unfixed at 4°C until they could be sorted on a BD FACS Aria cell sorter.

Activation-induced marker stimulations

PBMC were thawed and rested overnight at 37°C in RPMI with 10% fetal calf serum and 1% L-glutamine. Overlapping peptide pools for hemagglutinin 1 (A/California/7/2009, catalog #NR-19244), hemagglutinin 3 (A/Perth/16/2009, catalog #NR-19266), nucleoprotein (A/California/7/2009, catalog #NR-18976), and matrix 1 (A/California/7/2009, catalog #NR-18977) were obtained from BEI Bioresources and resuspended in DMSO. Stimulation was performed in flat-bottom plates with 0.5 μg/mL of each peptide pool for 18 hours, followed by surface staining for 20 minutes at room temperature. Acquisition and sorting were performed on a BD FACS Aria cell sorter.

Hemagglutination-inhibition assays

Sera were treated with receptor destroying enzyme and then heated for 30 minutes at 55°C. Sera were serially diluted in 96-well round bottom plates. Four agglutinating doses of A/California/7/2009 or A/Switzerland/9715293/2013 were added to the sera in a total volume of 100 μL. Turkey red blood cells were added (12.5 μL of a 2% (vol/vol) solution)), and agglutination was read out 60 minutes later. Titers are expressed as the inverse sera dilution that inhibited viral agglutination.

RT-qPCR

PBMC and splenocytes were thawed and sorted for subsets on a BD FACS Aria cell sorter followed by RNA extraction by Qiagen Micro Plus kit. Reverse transcription using the High-Capacity Reverse Transcription kit was performed as per manufacturer instructions (Applied Biosystems). Real-time PCR was performed using pre-designed primers and hydrolysis probes for B2M (Hs.PT.58v.18759587, IDT) and BCL6 (Hs.PT.56a.19673829.g, IDT) using the PrimeTime Gene Expression Master Mix (IDT) on a ViiA7 Real-Time PCR system (Applied Biosystems).

T cell receptor sequencing

PBMC were sorted on a BD FACS Aria followed by DNA extraction by Qiagen QIAamp DNA Micro kit. Amplification, library preparation, sequencing, and preliminary bioinformatics analysis was performed by Adaptive Biotechnologies (Seattle, WA). TCRB was sequenced at Survey level resolution.

TCR sequencing analysis

Calculations were performed in R for Shannon’s entropy and normalized Shannon’s Entropy (46, 47). Gini index was calculated using the R package “ineq” (48). Clonality was calculated as 1 - normalized Shannon’s Entropy. The overlap score was calculated as the sum of the counts for all shared sequences observed both in sample A and sample B, divided by the total number of counts for all sequences in samples A and B (shared and unshared) in the two samples. Some figures were generated with packages “ggplot2” and “gplots” (49, 50). All R code used for these analyses are available at https://github.com/Sedmic/cTfh_TCRB_2015-2016FluSpec.

Cytokine production

PBMC were stimulated for five hours in the presence of PMA/ionomycin or left unstimulated as a control. Monensin was added for the final four hours of stimulation. Staining was performed at 37°C in the stimulation medium followed by fixation with 1% para-formaldehyde for 5 minutes at room temperature. Cells were then permeabilized with the Foxp3 Fixation/Permeabilization Concentrate and Diluent kit and intracellular staining was performed for one hour at room temperature with antibodies against IL-2, IL-10, IL-13, IL-17, IL-21, TNFα, and IFNγ. Pestle and SPICE (51) were used to analyze polyfunctionality.

Statistics

Statistical analyses were performed with Prism 5 (GraphPad) or with R. Data was compared using Student’s t test, paired t test, or one-way Analysis of Variance (ANOVA) with Tukey post-hoc analysis, as indicated. Correlation analyses were performed as the Pearson correlation. Fisher’s Exact test was used to compare overlap in TCR sequences, based on 10⁷ theoretical TCRB sequences.