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CD4+ follicular regulatory T cells optimize the influenza virus–specific B cell response

Seasonal influenza virus infections cause severe illness and a substantial number of deaths worldwide every year. This study reveals that the CD4+ follicular regulatory T cells are necessary for optimal antigen-specific humoral immunity following influenza virus challenge.


Introduction
Antibodies form the first line of defense against invading pathogens and provide the basis for many successful vaccines through establishing a humoral memory response (Nutt et al., 2015). One of the hallmarks of humoral immunity is the progressive increase in the affinity of antibodies over time, a process occurring in germinal centers (GCs) within B cell follicles of secondary lymphoid organs (De Silva and Klein, 2015;Eisen, 2014). Upon encounter with pathogens, B cells mature into memory B cells and long-lived antibody-producing plasma cells (LLPCs) in GCs. Following reexposure to the pathogen, highaffinity protective antibodies secreted by LLPCs are a first line of protection; additionally, pathogen-specific memory B cells are activated and rapidly mature into plasmablasts to produce protective antibodies (De Silva and Klein, 2015). Within the GC, B cells undergo proliferation and somatic hypermutation of Ig genes, followed by a process in which the "fittest" B cells, those able to capture antigen via surface Ig and best present it on surface MHCII, are selected by follicular helper T (Tfh) cells . The strength of interaction between Tfh cells and GC B cells, which is proportional to the amount of antigen presented by GC B cells, drives the cyclic reentry and determines the cell cycle speed and number of divisions of GC B cell clones (Victora et al., 2010;Gitlin et al., 2014;Shulman et al., 2014).
Follicular regulatory CD4 + T (Tfr) cells, a population of regulatory T (Treg) cells, express the transcription factor Bcl6 and chemokine receptor CXCR5 and localize to the follicle and the GC following infection or immunization (Linterman et al., 2011;Chung et al., 2011;Wollenberg et al., 2011). Tfr cells suppress excessive immune responses, such as those occurring in autoreactivity, through a process mediated by the inhibitory coreceptor CTLA-4 (Wing et al., 2014;Sage et al., 2014). However, the role of Tfr cells in regulating antigen-specific B cell selection within the GC is currently unclear. The existing paradigm suggests that Tfr cells are inhibitory and restrain overall GC responses, including antigen-specific B cell responses (Wing et al., 2014;Clement et al., 2019;Sage et al., 2016). In contrast, other groups have reported no obvious effect on antigen-specific antibody responses in the absence of Tfr cells (Fu et al., 2018;Botta et al., 2017). Elucidating the regulatory function of Tfr cells in modulating antigen-specific B cell responses using physiologically relevant models has fundamental implications for our understanding of the selection process occurring within the GC.
Treg cells can suppress immune-mediated inflammation and self-reactive autoimmune diseases (Josefowicz et al., 2012). They also promote optimal immune responses by actively maintaining homeostasis. For example, Treg cells are necessary for the homeostatic priming of the CD8 + T cell response by increasing the avidity of the primary response (Pace et al., 2012). Treg cells also promote CD8 + T cell memory maturation by actively dampening inflammation during the resolution course of infection, aiding a return to homeostasis (Laidlaw et al., 2015(Laidlaw et al., , 2016Kalia et al., 2015). In line with these findings, we previously demonstrated that Tfr cell-derived IL-10 was necessary to maintain the GC B cell response following acute infection of lymphocytic choriomeningitis virus (Laidlaw et al., 2017).
Based on these observations, we hypothesized that Tfr cells might promote the selection of antigen-specific B cell responses, and accordingly, promote GC B cell output. We find that Tfr cells promote antigen-specific GC B cell responses during influenza viral challenge, with development of plasma cells and effective humoral memory. These findings indicate that Tfr cells optimize GC responses and regulate B cell recall responses, thus enhancing our understanding of the role of Tfr cells in vaccineinduced immunity, important in the optimization of vaccine development strategies.

Tfr cells promote the antigen-specific B cell response during the late course of influenza virus infection
To test the role of Tfr cells in regulating antigen-specific GC responses, we infected C57BL/6 mice with a mouse-adapted strain of influenza virus, A/Puerto Rico/8/1934 H1N1 (PR8), and isolated lymphocytes from the relevant draining mediastinal LNs (mLNs) at various time points following infection. Consistent with published studies (Botta et al., 2017), Tfr cell numbers peaked at day 30 following infection, at which time total GC B cells and antigen-specific GC B cells also reached maximum numbers (Fig. S1, A, C, and D); Tfh cell numbers peaked earlier, at day 12 post infection (p.i.; Fig. S1 B). We generated mice that specifically lack Tfr cells (Bcl6 f/f Foxp3-Cre) and confirmed their depletion at both early and late time points following infection, day 8 and day 40 p.i., respectively (Fig. S1 E). Bcl6 f/f Foxp3-Cre animals displayed no sign of overt disease at steady state and did not show differences in the basal number of GC B cells, Tfh cells, or Treg cells in the spleen compared with their littermate controls (Fig. S1 F).
To investigate whether the GC response was affected in the absence of Tfr cells, we isolated lymphocytes from mLNs of Tfr cell-deficient (Bcl6 f/f Foxp3-Cre) and control (Bcl6 f/f ) animals at day 38 p.i. This time point was chosen with the rationale that the GC response might be affected the most in the absence of Tfr cells following their numerical peak at day 30 p.i. The percentage and number of B cells with a GC phenotype (GL7 + CD38 − of B220 + IgD low population) were comparable between the two groups of mice ( Fig. 1 A). The GC sizes were also similar as determined by confocal imaging of the mLN sections ( Fig. 1 B). We then assessed the GC B cells that have specificity for the influenza virus surface glycoprotein hemagglutinin (HA). There was a marked decrease in percentage as well as number of HAspecific GC B cells as measured by recombinant HA probes, which have a mutation in the sialic acid binding site (Tyr98Phe) to prevent nonspecific binding to sialic acid on the cell surface ( Fig. 1 C; Angeletti et al., 2017;Whittle et al., 2014). The transcriptomes of HA-specific GC B cells were similar between the Tfr cell-deficient and -sufficient groups (Fig. 1 D). The viral loads in Tfr cell-deficient (Bcl6 f/f Foxp3-Cre) and control (Bcl6 f/f ) animals were also comparable at both day 5 and day 9 p.i., suggesting that the numerical difference in antigen-specific GC B cells between the two groups was not due to differences in viral load or viral clearance (Fig. 1 E).
As GC B cells give rise to high-affinity, antibody-producing plasma cells, we next sought to determine whether the antigenspecific plasma cell compartment was impaired in the absence of Tfr cells following influenza virus infection. We quantified HAspecific antibody-secreting cells (ASCs) using ELISPOT assays. ELISPOT analysis of bone marrow cells showed that there was a statistically significant reduction in the number of HA-specific ASCs in the Tfr cell-deficient animals compared with that in the controls, despite comparable numbers of total bone marrow cells between the two groups ( Fig. 2 A). The intensity of the spots was also reduced in the absence of Tfr cells, as shown by the representative images of ELISPOT assays. This suggests the possibility that the antigen-specific antibodies produced per ASC are reduced in the absence of Tfr cells or that their affinity is lower in Tfr cell-deficient animals.
While serum antibodies produced early in an immune response are mainly derived from short-lived plasmablasts generated as part of an extrafollicular response, those from later time points are mostly produced by ASCs developed via GCs (Nutt et al., 2015). We next evaluated antigen-specific antibody responses by assessing titers of antibodies to the influenza virus HA using quantitative ELISA. We found that while the induction of HA-specific IgG antibodies was similar between Bcl6 f/f Foxp3-Cre and Bcl6 f/f controls at day 15 p.i., IgG antibodies targeting HA were significantly reduced in mice lacking Tfr cells relative to controls at day 36 p.i. (Fig. 2 B). Consistently, Tfr cell-deficient mice had significantly lower titers of HA-specific Ig isotypes IgG1 and IgG2a (IgG2c in C57BL/6 animals) than those in their wild-type counterparts (Fig. 2 C). We next asked whether this phenomenon was unique to the antigen HA. We measured antibody responses to neuraminidase (NA), the second most abundant surface glycoprotein of influenza virus, which also contributes to protection from influenza virus infection . Likewise, NA-specific IgG antibody titers were reduced in Tfr cell-deficient mice relative to the controls at day 36 p.i. (Fig. 2 D). Thus, mice that lacked Tfr cells had decreased antigen-specific antibody titers, which was consistent with the reduction in antigen-specific GC B cells and plasma cells. Tfr cell-deficient mice also had reduced affinity of anti-HA IgGs compared with wild-type controls (Fig. 2 E). We observed the same comparative reductions in antigen-specific GC B cells and Volcano plot of -log 10 P value vs. log 2 fold change of the most differentially expressed 5,000 genes. The dashed line plotted indicates the level of Bonferroni-corrected P value <0.05. No gene is found to show significance change with adjusted P value <0.05. (E) Plaque assay measuring viral load in Bcl6 f/f and Bcl6 f/f Foxp3-Cre mice at day 5 and day 9 p.i. Statistical analyses were performed using the unpaired two-tailed Student's t test (*, P < 0.05; **, P < 0.01). Data for A and C are from one experiment representative of four experiments with five to seven mice per group. Data for B and E are from one experiment representative of two experiments with three to five mice per group. Data for D are from two independent experiments with three to four mice per group pooled for each sample. n.s., not significant. antibody responses in Bcl6 f/f Foxp3-Cre animals when we used Foxp3-Cre littermates as controls (Fig. S2, A and B). The total serum IgG concentrations were higher in the Tfr cell-deficient animals compared with the controls, though the differences did not reach statistical significance (Fig. 2 F). We did not detect serum anti-nuclear antibody (ANA) responses in the absence of Tfr cells following influenza virus infection or in steady state with a serum dilution of 1:20 (Fig. S3, A and B). However, with a serum dilution of 1:5, we observed weak positive staining using sera from Tfr cell-deficient animals, but not those from control animals following infection or in steady state from either group (Fig. S3, C and D). Blinded scoring of ANA staining revealed that the Tfr cell-deficient group had higher positivity scores compared with those of the control group (Fig. S3 E;Peng and Craft, 2020).
Adoptive transfer of Treg cells rescues impairment in the antigen-specific GC B cell response These data demonstrated that Tfr cells promote antigen-specific B cell responses during GC development. To further validate that Tfr cells are sufficient for the optimal influenza virus-specific GC B cell response, we adoptively transferred sorted CD4 + Foxp3 + Treg cells from congenically marked naive Treg cell reporter animals (Foxp3 EGFP CD45.1 + ) into the Bcl6 f/f Foxp3-Cre group 1 d before influenza virus infection with the idea that Treg cells differentiate into Tfr cells upon infection (Fig. 3 A). We could not adoptively transfer Tfr cells as we were limited by their number obtained during influenza virus infections. We observed that the adoptive Treg cell transfer restored HA-specific GC B cells in Bcl6 f/f Foxp3-Cre mice, both in percentage and number, to levels comparable to those in Bcl6 f/f control mice at day 40 p.i. (Fig. 3 B). Additionally, we did not observe the transferred Treg cells transdifferentiate into Tfh cells (Fig. 3 C). We confirmed that the transferred Treg cells differentiated into Tfr cells in the Bcl6 f/f Foxp3-Cre animals ( Fig. 3 D). These results demonstrate that Treg cells and most likely Tfr cells are important for the optimal antigen-specific GC B cell responses.
Polyclonal and antigen-specific Tfh cell responses are not affected in the absence of Tfr cells In addition to affecting GC B cells, Tfr cells have the potential to influence the Tfh cell population. We assessed the Tfh cell response in mLNs at day 8 and day 38 following influenza virus infection. Tfh cells were gated as CD4 + CD44 hi Foxp3 − Ly6C lo Psgl-1 lo PD-1 hi CXCR5 hi (Marshall et al., 2011;Crotty, 2011). At day 8 p.i., although the percentage of Tfh cells in mice that lacked Tfr cells was reduced compared with control mice, their absolute numbers were comparable between the two groups ( Fig. S4 A). The expressions of the lineage-defining transcription factor Bcl6 and proliferation marker Ki67 were similar between the two groups ( Fig. S4 B). RNA sequencing (RNA-seq) analysis revealed top differentially expressed genes not typically expressed by T cells, and thus we concluded there was no apparent difference in gene expression profiles of Tfh cells in Bcl6 f/f Foxp3-Cre and Bcl6 f/f control mice at this time point (Fig. S4 C and Table S1). HA-specific and nucleoprotein (NP)-specific Tfh cell percentages and numbers were also similar between the Tfr cell-sufficient and Tfr cell-deficient groups ( Fig. S4 D). During the late course of infection, at day 38 p.i., both the percentages and absolute numbers of Tfh cells were similar between the Tfr cell-sufficient and Tfr cell-deficient groups (Fig. S4  E). These data indicate that the Tfh cell response is not affected by the lack of Tfr cells during influenza virus infection. Additionally, both the percentages and absolute numbers of Treg cells were similar between the Tfr cell-sufficient and Tfr cell-deficient groups at day 9 and day 38 p.i. (Fig. S4, F and G), suggesting that total Treg cells are not affected by the lack of Tfr cells following infection.
Tfr cell-deficient mice use distinct V H (variable region of the heavy chain) genes compared with their wild-type counterparts following influenza virus infection To explore whether the B cell receptor (BCR) repertoire was altered in the absence of Tfr cells, we performed high-throughput BCR repertoire profiling. We sorted GC B cells from mLNs of Bcl6 f/f Foxp3-Cre and Bcl6 f/f control animals at day 15 p.i., as well as naive follicular B cells from mLNs of uninfected animals, and performed high-throughput sequencing of the BCR heavy chain mRNA. This approach minimizes PCR bias and consequence of errors introduced by sequencing as previously described (Vander Heiden et al., 2014). We analyzed ∼5,468, 5,399, and 561 reconstructed unique and error-corrected Igh sequences that passed quality control from Bcl6 f/f Foxp3-Cre, Bcl6 f/f , and naive, uninfected animals, respectively (Table S2). Principal component analysis revealed that the V H gene usage in BCR sequences of follicular B cells from naive animals clustered separately from those of GC B cells from infected animals ( Fig. 4 A). Moreover, Tfr cell-deficient and Tfr cell-sufficient mice showed distinct V H gene usage signatures following infection ( We next investigated the significance of differential V H gene usage in the absence of Tfr cells by reanalyzing Sanger-based sequencing of B cells from recombinant HA-immunized mice in a published study (Kuraoka et al., 2016). Notably, the V H genes that were differentially expressed between Bcl6 f/f Foxp3-Cre and Bcl6 f/f control mice following infection were positively correlated with the V H genes enriched following HA immunization as assessed by Pearson correlation analysis (Fig. 4 D). No statistically significant correlation with the expanded V H genes in response to anthrax protective antigen (PA), an irrelevant antigen control, was detected ( Fig. 4 D). Other features of the repertoire analysis appeared to be similar between the two groups following influenza virus infection, including clonality, selection pressure, isotype usage, and somatic hypermutation (Fig. S5, B-F). We therefore reasoned that Tfr cells were able to shape the B cell repertoire following infection by selecting for B cells that use specific BCR sequences known to be involved in antigen-specific responses.
Sequential vaccination with partially conserved antigens reveals that Tfr cells promote effective memory formation Immunological memory is crucial for B cell responses to influenza virus in humans (Andrews et al., 2015). Since the specificity of the primary response affects secondary responses, impaired antigen-specific B cell primary responses in the absence of Tfr cells could lead to defective memory responses. To investigate this possibility, we used a sequential vaccination model with repeated exposure of antigenically distinct strains of influenza viruses which shared the conserved immunosubdominant stalk domain of the glycoprotein HA to induce stalk-specific antibody responses Nachbagauer and Palese, 2020;Choi et al, 2019). We immunized Tfr celldeficient and -sufficient mice with H1N1 intramuscularly, followed by a boost with a laboratory reassortant strain, H5N1, at day 42 after priming and then a second boost with another Representative plots of Tfr cells derived from the transferred Treg cells gated as CD45.1 + Foxp3 + in Bcl6 f/f , Bcl6 f/f Foxp3-Cre and Bcl6 f/f Foxp3-Cre + Treg cell animals. Tfr cells are defined as CD4 + CD44 hi Ly6C − PSGL1 lo CXCR5 hi PD1 hi Foxp3 + . Statistical analyses were performed using the unpaired two-tailed Student's t test (*, P < 0.05; **, P < 0.01). Data are from one experiment representative of two experiments with four or five mice per group performed 38 d following influenza virus infection. reassortant strain, H6N3, at day 84 (Fig. 5 A). These three viral strains (all influenza A group 1 viruses) have divergent HA head domains but share a similar conserved stalk domain. Thus, following each boost with a heterologous strain, the animals mount primary responses toward the head domains (to which they are naive), while reexposure to the conserved stalk domain can boost memory response toward this domain (Nachbagauer and Palese, 2020).
We measured memory responses by detecting stalk-reactive antibodies in Bcl6 f/f Foxp3-Cre and Bcl6 f/f control animals. As previously described, we performed stalk-specific ELISAs using a chimeric HA cH11/1 comprising an H11 head domain (to which mice were not exposed) and an H1 stalk domain as substrate (Nachbagauer et al., 2014). The reactivity measured by ELISA toward this substrate is predominantly specific to the H1 stalk  Kuraoka et al. (2016) associated with HA specificity (compared with non-HA specifics; left) or with PA specificity (compared with non-PA specific sequences; right) at day 16 after immunization. Correlations were computed as Pearson correlation coefficients, and a P value was assigned to the significance of the correlation with a significance threshold of P < 0.05. For the BCR repertoire analysis, data are from one experiment with four mice per group performed 15 d following PR8 influenza virus infection. Cor., Pearson correlation coefficient. and Bcl6 f/f mice measured by influenza hemagglutination inhibition assay following sequential vaccination. Statistical analyses were performed using the unpaired two-tailed Student's t test (*, P < 0.05). Data for A-C are from one experiment representative of two experiments with five to seven mice per group performed at various time points following sequential vaccination with diverse group 1 influenza viruses. Data for D are pooled from two independent experiments with five to seven mice per group following sequential vaccination. AUC, area under the curve; n.s., not significant. domain. At day 40 after the first boost, Tfr cell-deficient mice showed a trend of reduced HA stalk-specific IgG titers compared with their wild-type counterparts (Fig. 5 B). This difference between the two groups of animals became statistically significant following the second boost (Fig. 5 B). The increase of anti-stalk antibody response following the second boost was not dramatic and could represent a "ceiling effect" that has been observed previously in chimeric HA-based vaccine studies, which used sequential immunization with vaccine strains expressing the conserved HA stalk domain and exotic HA heads (Nachbagauer et al., 2016). Consistent with reduced HA stalkspecific serum IgG titers, Tfr cell-deficient mice also had significantly impaired HA stalk-specific plasma cells compared with their wild-type counterparts as determined by HA stalkspecific ELISPOT assay, while the numbers of total bone marrow cells were similar between the two groups (Fig. 5 C). Sera from both Tfr cell-deficient and -sufficient animals showed comparable weak ANA staining at 40 d after the second boost (data not shown), possibly due to advanced age of the animals at the end of the experiments. HA head-specific antibody titers, measured by hemagglutination inhibition assay, which assesses the antibodies against the HA head domain , were also reduced in Tfr cell-deficient animals compared with their wild-type littermates (Fig. 5 D). This is consistent with the observation that animals had impaired primary responses in the absence of Tfr cells. These data indicate that Tfr cells promote productive antigen-specific humoral memory responses in response to heterologous influenza infections.

Discussion
We found that Tfr cells support influenza virus-specific B cell responses to infection and vaccination. In the absence of Tfr cells, influenza virus-specific GC B cells, LLPCs, and antibody titers are significantly reduced. We discovered an altered BCR repertoire following infection in mice lacking Tfr cells, which may account for a reduction in the expanded influenza virusspecific B cells. The impaired primary response then further leads to impaired humoral memory responses following heterologous influenza virus vaccinations. Previous studies have explored the role of Tfr cells in modulating antigen-specific responses in the setting of influenza virus infection, with results showing that anti-influenza virus antibodies and plasma cells did not appear to be different between Tfr cell-deficient and Tfr cell-sufficient animals (Botta et al., 2017;Fu et al., 2018). These two studies assessed antigenspecific B cell responses at day 9 and day 30 following infection, respectively. At day 9 p.i., the difference between the two groups was still not detectable in our experiments. At day 30 p.i., there was a trend of reduced influenza virus-specific plasma cells in Bcl6 f/f Foxp3-Cre animals compared with controls in the published work (Botta et al., 2017), which is consistent with our data supporting a role for Tfr cells at late time points following infection (36 d p.i. and beyond). The kinetics also correspond well with the number of Tfr cells peaking at 30 d following influenza virus infection.
Our finding contrasts with previous studies, in which Tfr cells were found to suppress antigen-specific B cell responses (Sage et al., 2016;Clement et al., 2019). In the latter examples, both the in vitro and in vivo experiments were done following immunization with the antigen NP-OVA (protein-hapten conjugate), with the insight gained from these studies serving as an important foundation upon which we build. It is likely that the function of Tfr cells depends on the complexity of the antigen, and the timing. We reason that in live viral infection, such as with influenza virus, a complex antigen with distinct epitopes and antigen persistence due to replication, the maintenance of diverse B cell clones, both antigen-specific and non-antigenspecific, is distinct from those in an immunization model with simpler antigens (Tas et al., 2016).
The Cre/LoxP system can have infidelity issues, as has been observed (Schmidt-Supprian and Rajewsky, 2007). To confirm that the phenotype was not due to leakiness of Cre recombinase, we conducted adoptive transfer experiments in which Treg cells rescued the impairment of antigen-specific GC B cells in the Tfr cell-deficient animals, suggesting that the lack of Tfr cells is responsible for the phenotype. Furthermore, the analyses of Tfh cells at day 8 p.i. and day 38 p.i. did not reveal differences in numbers of Tfh cells, confirming that they were not affected in this Cre system. Overall, we exclude the possibility of Cre leakiness causing impaired antigen-specific GC B cell responses.
An impaired antigen-specific humoral response was also not due to differences in viral load or viral clearance, as viral titers were similar between the Tfr cell-deficient and -sufficient groups at various time points. Additionally, in the sequential vaccination experiments, the route of immunization was intramuscular. The virus is not able to complete more than one replication cycle due to the lack of specific host proteases in the muscle tissue, which are required for cleavage of the influenza virus surface HA (Böttcher-Friebertshäuser et al., 2010). Therefore, we argue that virus clearance does not play a role in the measured antibody response in this setting as well.
It seems intuitive and evolutionarily sensible to have regulatory cells participating in GC responses to optimize the output, which is antigen-specific high-affinity responses. Yet how do Tfr cells achieve that goal? One possibility is that Tfr cells preferentially suppress non-antigen-specific GC B cells while sparing the antigen-specific ones. This would be reminiscent of the role played by Treg cells in optimizing CD8 + T cell responses: Treg cells preferentially suppress responses by T cells that have weak, lower-affinity interactions with their cognate antigen (also the reason that Treg cells control self-reactive T cells due to their low-affinity) while sparing high-affinity interactions (Pace et al., 2012). One can imagine a similar scenario for the GC selection process of antigen-specific B cells regulated by Tfr cells. Specifically, as the positive selection signal is directly proportional to the strength of interaction between Tfh cells and GC B cells, Tfr cells can selectively inhibit non-antigen-specific GC B cells based on their low-strength interactions with Tfh cells, whereas antigen-specific GC B cells that engage in high-strength interactions with Tfh cells can overcome this suppression. The expression of positive selection signals, such as mTORC and cMyc signaling, which are up-regulated in GC B cells having recently received Tfh cell help, is likely to serve as a proxy for the strength of interaction that can be regulated by Tfr cells (Dominguez-Sola et al., 2012;Ersching et al., 2017). Furthermore, overproliferation and outgrowth of non-antigen specific B cell clones in the absence of Tfr cells can compete for limited Tfh cell help within a given GC, therefore resulting in impaired antigenspecific B cell responses.
It will be of interest to identify Tfr cell-derived molecules (e.g., cytokines, surface molecules) that might mediate the aforementioned process of optimizing antigen-specific B cell responses within the GC. We explored one Tfr cell-derived molecule, IL-10, which does not affect the level of influenza virus-specific B cell response or polyclonal Tfh cell response (data not shown). It will be important for future work to focus on specific Tfr cell-derived molecules and elucidate their functions in modulating different aspects of productive GC responses. In summary, our study reveals a novel function of Tfr cells in promoting antigen-specific B cell responses following influenza virus infection. Targeting Tfr cells or Tfr cell-derived molecules may provide more effective vaccine development strategies in the future.

Design
The objective of this study was to determine the role of Tfr cells in regulating antigen-specific B cell responses following influenza virus infection. Tfr cell-deficient and Tfr cell-sufficient mice were subjected to influenza virus infections or immunizations. The antigen-specific B cell responses were assessed by flow cytometry, ELISPOT, ELISA, and BCR sequencing. Detailed methods are described in the sections below. Control and experimental groups were sex and age matched. Littermates were used for all experiments where possible. The investigators were not blinded. The sample sizes and experimental replications are indicated in the figure legends.
Mice C57BL/6 mice were purchased from the National Cancer Institute or The Jackson Laboratory. C.Cg-Foxp3 tm2Tch /J (Foxp3 EGFP ), B6.129(FVB)-Bcl6 tm1.1Dent /J (Bcl6 f/f ), B6.129 (Cg)-Foxp3 tm4(YFP/cre) Ayr/J (Foxp3-Cre), and B6.129P2(C)-Cd19 tm1(cre)Cgn /J mice were purchased from The Jackson Laboratory. Il10 f/f mice (Roers et al., 2004) have been described. Il10ra f/f mice were generated by the Flavell laboratory as previously described (Laidlaw et al., 2015). All animal experiments were done with approval of the Yale Institutional Animal Care and Use Committee. Reassortant viruses were rescued as described previously (Martínez-Sobrido and García-Sastre, 2010). All viruses were propagated in 10-d-old specific pathogen-free eggs (Charles River) after inoculation with 100 PFUs of virus. Infected eggs were incubated at 37°C for 48 h before harvesting and clearance of allantoic fluid. Virus-containing fluid was aliquoted and stored at −80°C before infection. For purified viral stocks, allantoic fluid was spun at 125,000 g for 2 h over a 5% sucrose cushion. Pelleted virion particles were resuspended in PBS and stored at −80°C before vaccination. Viral loads were measured by plaque assay on Madin-Darby canine kidney cells. Briefly, Madin-Darby canine kidney cells were plated in 6-well dishes at 8 × 10 5 cells/well and allowed to incubate overnight at 37°C and 5% CO 2 . Samples were diluted in PBS and added to cells to allow viral absorption for 1 h at 37°C and 5% CO 2 with periodic shaking. Cells were then rinsed with PBS, and an agar overlay was added to allow the virus to infect in the presence of tosyl phenylalanyl chloromethyl ketone-treated trypsin. Plaques were visualized with crystal violet staining and counted.

Viruses, proteins, and cells
All the recombinant HA and NA proteins used in ELISA and ELISPOT assays were kind gifts from F. Krammer (Icahn School of Medicine at Mount Sinai, New York, NY) and were generated and purified as previously described (Margine et al., 2013).

Infection and treatment
Mice were given i.n. administration of 8 PFUs of influenza virus A/PR8/34 (PR8) in 30 µl of PBS. Prior to i.n. infections, the mice were anesthetized by ketamine hydrochloride and xylazine (Phoenix Scientific) in 200 µl of PBS via i.p. injections. For the sequential vaccination strategy, mice were primed with 2.5 µg per mouse of the A/Michigan/45/2015 (H1N1) through intramuscular vaccination, followed 6 wk later by H5N1 vaccination intramuscularly. All the animals received a second boost on day 84 with H6N3. The influenza virus strains used in the sequential vaccination experiments were approved for usage by the Icahn School of Medicine at Mount Sinai Biosafety Committee.
Flow cytometry, cell sorting, and antibodies for surface and intracellular staining Staining of transcription factors was performed after permeabilization with the FoxP3 Fixation and Permeabilization Kit (eBioscience). For intracellular cytokine staining, cells were stimulated with PMA (50 ng/ml) and ionomycin (1 µg/ml) in 96well U-bottom plates in complete media for 4-5 h, and brefeldin A (BD Biosciences) was added for the last 3-4 h. Intracellular staining was performed using the BD ICS kit as per the manufacturer's instructions with overnight incubation (4°C) of permeabilized cells with transcription factor-specific antibodies. Antibodies are listed in Table S3. Recombinant HA probe for B cells was generated as previously described (Angeletti et al., 2017). The HA I-A(b) tetramer (PR8 HA 91-107 RSWSYI-VETPNSENGIC; Miller et al., 2015) and the NP I-A(b) tetramer (NP 311-325 QVYSLIRPNENPAHK; Botta et al., 2017) were produced by the National Institutes of Health Tetramer Core Facility. Flow cytometry data were acquired on a BD LSRII with FACSDiva software and were analyzed with FlowJo software (Tree Star).
Isolation of lymphocytes from tissues mLNs were homogenized with a cell strainer. Lymphocytes were then washed and counted.
ELISA ELISAs were performed as described previously . Specifically, 96-well high-binding, flat-bottomed plates were coated with 50 µl/well of recombinant HA protein at a concentration of 2 µg/ml in coating buffer (50 mM sodium carbonate and 50 mM sodium hydrogen carbonate, pH 9.4) and were incubated overnight at 4°C. The coating buffer was removed, and plates were incubated for 1 h at room temperature with 220 µl of blocking solution (3% goat serum, 0.5% Tween-20, and 3% milk powder in PBS). Serum was serially diluted threefold in blocking solution with a starting dilution of 1:50, and plates were incubated for 2 h at room temperature. Plates were then washed three times with PBS with 0.5% Tween-20 (PBS-T) followed by adding 50 µl of isotype-specific secondary antibody diluted in blocking solution to each well (IgG [ab97265], IgG1 [ab97240], and IgG2c [ab97255]; Abcam). After 1 h of incubation at room temperature, plates were washed four times with PBS-T. Plates were developed with 100 µl of Sig-maFast OPD (Sigma-Aldrich), and the development was stopped after 10 min by adding 50 µl of 3 M HCl. Plates were then read at a wavelength of 490 nm with a Synergy H1 hybrid multimode microplate reader (BioTek). Background was determined as the average of blank wells plus three times the standard deviation between those wells. Area under the curve was measured after fitting a regression curve using the formula [Agonist] vs. response − variable slope (four parameters) and using the previously determined background value or 0.07, whichever was higher. To measure affinity by ELISA, a 10-min, 7 M urea wash was introduced after incubating serum samples with the HA substrate. Percentage of IgG binding after urea wash was determined by dividing the IgG bound to HA treated by 7 M urea by untreated IgG bound.
Total IgG were measured using Mouse IgG ELISA kit (Innovative Research) following manufacturer's instructions.
ELISPOT for HA-specific Igs MultiScreen HTS plates (Millipore) were coated with 50 µl/well of recombinant HA protein at a concentration of 2 µg/ml in PBS overnight at 4°C. The coating solution was removed, and plates were washed six times with PBS-T followed by incubation with 200 µl of complete medium (10% fetal bovine serum and 1% L-glutamine, penicillin, and streptomycin in RPMI [Life Technologies]) for 2 h at 37°C. Subsequently, bone marrow cells from mice infected with the PR8 influenza virus were isolated and plated (3 × 10 6 cells/well) in duplicate in complete medium for 4-5 h at 37°C. Plates were then washed six times with PBS-T followed by addition of secondary antibody (50 µl) in blocking solution (0.5% BSA in PBS; anti-IgG-alkaline phosphatase; Southern Biotech [1030-04]) to each well, then incubated overnight at 4°C. The plates were washed three times with PBS-T and three times with PBS. Spots were developed with Vector Blue (Vector Laboratories) and quantified using an ImmunoSpot analyzer (Cellular Technology Limited).

ANA detection
Serum dilutions of 1:20 and 1:5 in PBS were used. ANA staining was performed using commercial HEp-2 slides (Antibodies Inc., 15-123) following the manufacturer's instructions, with degree of staining graded in a blinded manner (Peng and Craft, 2020).

BCR repertoire library preparation and data analysis
Lymphocytes from mLNs were isolated as described above and enriched for B cells by positive selection using magnetic beads (StemCell). GC B cells from infected animals were sorted as B220 + , IgD low , GL7 + , CD95 + cells, and naive follicular B cells from uninfected animals were sorted as B220 + , IgM + , CD23 + CD21 − cells. Following FACS sorting, cells of both types were pelleted, and RNA was isolated with the use of the RNeasy Plus Micro Kit (Qiagen). High-quality RNA with an RNA integrity number >9 was subsequently used to prepare BCR libraries with the NEBNext Immune Sequencing Kit (mouse; New England Biolabs) following the manufacturer's instructions. Specifically, ∼50-100 ng of RNA was reverse transcribed into cDNA with the addition of a unique molecular identifier (UMI) and an adaptor sequence followed by purification of the nascent cDNA using streptavidin magnetic beads. A PCR reaction was conducted to amplify the VDJ region of the heavy chain and incorporate the index barcode followed by a subsequent cleanup of the PCR products with AMPure XP beads (Beckman Coulter). Quantitative PCR analysis was performed to determine the optimal number of cycles for the second PCR amplification of each sample. This second PCR was then performed with the addition of a P5 adaptor to the Ig constant region and a sample-indexed P7 adaptor to the universal adaptor end. Final PCR products were purified again using AMPure XP beads and quantified through automated electrophoresis (Tapestation; Agilent). Equal amounts of each sample (6 µg) were pooled and followed by high-throughput 325 + 275 base pair paired-end sequencing with 30% PhiX spike-in on the Illumina MiSeq platform according to the manufacturer's guidelines.
Processing of BCR repertoires was performed using tools from the Immcantation framework (http://immcantation.org). The following preprocessing steps were performed using pRESTO v0.5.10 tools (Vander Heiden et al., 2014). Reads with a mean Phred score <20 were removed. Constant region (>0.8 identity) and template switch primers (>0.5 identity) were identified, while those sequences without a match were excluded. Primer sequences for pRESTO were TS-Shift0 TACGGG, TS-Shift1 ATACGGG, TS-Shift2 TCTACGGG, TS-shift3 CGATACGGG, and TS-shift4 GATCTACGGG. UMI sequences were identified from the first 17 nucleotides preceding the template switch site. A consensus sequence was constructed for forward and reverse reads for each UMI group and was aligned with a minimum overlap of eight nucleotides and with a mismatch identity <0.3 to assemble VDJ sequences. To reconstruct full-length VDJ sequences, sequence pairs failing alignment were aligned against the ImMunoGeneTics (IMGT) mouse immunoglogbulin V H genes reference database (IMGT/GENE-DB v3.1.19; retrieved June 21, 2018; mismatch identity <0.5; E-value threshold <1 × 10-5.6). Duplicate VDJ sequences were removed. IgBLAST v1.7.0 was used for germline assignments (Ye et al., 2013). Nonfunctional VDJ sequences were removed. Clonal clusters were assigned using Change-O v0.3.4, an approach described previously, and a junction threshold of 0.084 was used (Gupta et al., 2015). Germline sequences were then reconstructed for each clonal cluster (V H ) with D segment and N/P regions masked (replaced with Ns) using the CreateGermlines.py function within Change-O v0.3.4. Mutations and the mutabilities of potential motifs were quantified using ShazaM v0.1.8 in R v3.4.2 relative to germline sequences. Diversity analysis was performed using tools from Alakazam v0.2.11 and assessed using the generalized Hill index involving downsampling to the number of VDJ sequences from the sample with the fewest sequences and 2,000 bootstrap replicates (Hill, 1973). Selection strengths were quantified using the BASELINe implementation for complementarity determining region and framework region (Yaari et al., 2012). Aggregate posterior distribution functions for each genotype were determined via convolution of posterior distribution functions from each sample of a given genotype.
The BCR sequencing data reported in this paper are tabulated in the online supplemental material and archived at NCBI Bio-Project (accession no. PRJNA673160).

RNA-seq library preparation and data analysis
Total RNA was purified with the use of a QIAzol and RNeasy Plus Micro kit (Qiagen). Purified total RNA was submitted to the Yale Center for Genomic Analysis, where it was subjected to isolation of mRNA and library preparation. Libraries were pooled, six samples per lane, and were sequenced on an Illumina HiSeq 2500 (75 base pair paired-end reads), followed by alignment with STAR software (Anders et al., 2013). A "count-based" differential expression protocol was adapted for this analysis (Anders et al., 2013). Mappable reads were counted with the feature Count and imported into R software for analysis of differential expression with DESeq2 software. A multi-factor design was used to analyze the two conditions (Bcl6 f/f Foxp3-Cre and Bcl6 f/f ) and account for pairwise groupings of the six samples. For the generation of the volcano plot, genes with average counts >1 were chosen for visualization. The RNA sequencing data are archived in the NCBI GEO database (accession no. GSE161788).

Statistical analysis
Results represent the mean ± SEM unless indicated otherwise. Statistical significance was determined by the paired or unpaired Student's t test. Statistical analyses were performed using Prism GraphPad software v7.0 (*, P < 0.05; **, P < 0.01; ***, P < 0.001) except in Fig. 4 C (a two-tailed Wilcoxon test was used with a significance threshold of *, P < 0.1; **, P < 0.05). Principal component analysis was performed using the pca function in R. Pearson correlation coefficients were computed using the corr.test function in base R.
Online supplemental material Fig. S1 shows the kinetics of CD4 + Tfr cells, Tfh cells, GC B cells, and HA-specific GC B cells following influenza virus infection and characterization of Tfr cell-deficient animals. Fig. S2 illustrates that Bcl6 f/f Foxp3-Cre animals have reduced HA-specific GC B cells and antibody response as compared with Foxp3-Cre littermate controls. Fig. S3 reveals that Tfr cell-deficient animals show weakly positive ANA staining following influenza virus infection. Fig. S4 shows that mice lacking CD4 + Tfr cells mount similar Tfh cell responses as compared with control mice at 8 and 38 d following influenza virus infection. Fig. S5 demonstrates that many mLN Igh repertoire features are consistent between wild-type and mice lacking CD4 + Tfr, including clonality, selection pressure, isotype usage, and somatic hypermutation. Table S1 lists the differentially expressed genes in Tfh cells between Bcl6 f/f Foxp3-Cre and Bcl6 f/f mice with adjusted P value <0.05. Table S2 is a summary of sequencing reads and reconstructed Igh VDJ sequences from naive and infected Bcl6 f/f and Bcl6 f/f Foxp3-Cre mice. Table S3 shows antibodies used for flow cytometry and microscopy.
S.H. Kleinstein, R. Nachbagauer, and J. Craft analyzed and interpreted the data; Y. Lu and J. Craft wrote the manuscript.
Disclosures: K.C. O'Connor reported personal fees from Alexion and grants from Ra Pharma outside the submitted work; is the recipient of a sponsored research subaward from the University of Pennsylvania, the primary financial sponsor of which is Cabaletta Bio; and is a consultant and equity shareholder of Cabaletta Bio. No other disclosures were reported.  Statistical analyses were performed using the unpaired two-tailed Student's t test (**, P < 0.01). Data for A, B, and E-G are from one experiment representative of three experiments with five to eight mice per group. Data for C are from three independent experiments with three or four mice per group pooled for each sample. Data for D are from one experiment representative of two experiments with four to six mice per group. n.s., not significant. (D) Mutability, the frequency at which 5-mer sequences with hot (WRC/GYW and WA/TW) or cold (SYC/GRS) spots are observed to be mutated, is plotted for each sample. Horizontal bars show the mean mutability frequency. (E) Clonal diversity at Hill diversity indices of Q = 0 (richness) and Q = 2 are plotted. Each point represents an estimated diversity score based on bootstrap realizations of the clonal abundance curve from each sample. Horizontal bars correspond to the mean diversity index across samples of a given genotype. (F) Baseline selection pressure analysis is shown as a density on the y axis and selection strength (å) on the x axis for both the complementarity determining region (CDR) and the framework region (FWR). For BCR repertoire analysis, data are from one experiment with four mice per group performed 15 d following PR8 influenza virus infection.