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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
J Immunol. Author manuscript; available in PMC May 1, 2013.
Published in final edited form as:
PMCID: PMC3331930
NIHMSID: NIHMS361301

CD80 expression on B cells regulates murine T follicular helper development, germinal center B cell survival and plasma cell generation1

Abstract

Germinal center (GC) B cells and T follicular helper (TFH) cells interact in the production of high-affinity long-lived plasma cells (PCs) and memory B cells, though the mechanisms regulating the formation of these long-lived populations remain unclear. Because CD80 is one of the few markers shared by human and murine memory B cells, we investigated its role in the development of GCs, memory cells and PCs. In CD80-deficient mice, fewer long-lived PCs were generated upon immunization, compared to B6 controls. In concert, the absence of CD80 resulted in an increase in apoptotic GC B cells during the contraction phase of the GC. CD80−/− mice had fewer TFH compared to B6, and residual TFH cells failed to mature, with decreased ICOS and PD-1 expression and decreased synthesis of IL-21 mRNA. Mixed bone marrow chimeras demonstrated a B cell-intrinsic requirement for CD80 expression for normal TFH and PC development. Therefore, B cell expression of CD80 plays a critical role in regulating B-T interactions in both early and late GC responses. This, in turn, results in impaired ability to produce long-lived PCs. These data provide new insights into the development of GCs and AFCs and the functions of CD80 in humoral immunity.

Introduction

Memory of prior exposure, which results in long-term protection against repeat infection, is a fundamental aspect of the adaptive immune system. In humoral immunity, long-lived protection is mediated by memory B cells and PCs (1). These cells emerge from the large heterogeneous population generated during the GC reaction, from which only a small number of memory B cells and long-lived PCs will be selected to persist (25). Although short-lived PCs, and possibly some memory B cells, can form without a GC, the quality and quantity of long-lived memory B cells and PCs are largely determined within the GC (69).

B cell differentiation and selection within the GC are influenced by interactions with other cell types, including follicular dendritic cells and T cells. Notably, a population of Ag-specific T cells that migrates to the follicle post-activation, known as T follicular helper (TFH) cells, provides signals to B cells throughout the response (10, 11). The nature of these signals, and how they regulate B cell behavior, is dynamic; the details of such interactions are now actively being defined (7, 1214). Reciprocally, B cells also influence the behavior of TFH themselves; development of TFH is impaired in the absence of B cells (1315). It is through this evolving relationship that the size of memory B cell and PC populations, as well as their characteristics such as affinity and Ig isotype, are determined.

Notably, several of the molecules known to influence the GC response are members of the B7 and CD28 family of surface receptors, including ICOSL-ICOS (1618); BTLA (19); and CD80, CD86 and CD28 (20, 21). Generally, roles for these molecules have been defined during the formation of responses, with severe blocks in GC development in the absence of all except CD80 and BTLA. Absence of CD80 was reported to have little, if any, effect on GC formation, while the effect of BTLA on the GC is unclear. The roles of these molecules in long-term humoral immunity remain largely unexplored. We recently reported that another pathway in the B7:CD28 family—PD-1 and its ligands—regulated B cell and TFH responses during the contraction phase of the GC, with effects on GC size and cell survival, as well as the TFH compartment, and the size and quality of the long-lived PC pool (11). These findings led us to investigate whether other B7:CD28 molecules regulate GC cell behavior and differentiation decisions. Here we focus on CD80, because unlike CD86, which is required for GC responses, it does not have a known role in the induction phase of the GC response.

CD80 is a B7 family member that can bind CD28, CTLA-4 and PD-L1. In contrast to CD86, which binds the stimulatory molecule CD28 more avidly than the negative regulatory molecule CTLA-4, CD80 preferentially binds CTLA-4 (22). In keeping with a lack of known function in the early GC response, CD80 is upregulated on activated B cells much later than is CD86 (23). Emerging data suggest that CD80 could have a unique and non-redundant function in humoral responses after the stage of peak GC response. CD80 is upregulated on both human and murine memory B cells (2426). CD80 expression on human memory B cells has also been shown to co-stimulate CD4+ T cell proliferation in vitro, independently of CD86 (26). A recent report stated that CD80 does not have a role in GC B cell responses; however, it is notable that the CD80 and CD86 double-deficient mice had a more severe phenotype than did CD86-deficient mice, suggesting again that CD80 does have a role in forming long-lived populations (27).

To assess the roles of CD80 in TFH and GC B cell dynamics and subsequent production of memory B cells and long-lived plasma cells in vivo, we used genetic approaches. We report here that CD80 expressed by B cells does indeed play a substantial and non-redundant role in regulating both formative and contraction phases of the GC, affecting the maturation of TFH, the maintenance of late GC responses, and differentiation of memory B cells and long-lived PCs. These data add an additional dimension to a developing theme indicating that co-inhibitory pathways in the B7:CD28 family are important in controlling the extent and quality of the humoral immune response.

Materials and Methods

Mice and immunization

CD80−/− mice were generated as described (28) and fully backcrossed onto the B6 (29) or BALB/c backgrounds (30). B6 and BALB/c controls were from Jackson Laboratories. Igh-J−/− mice (31) were generated as described, backcrossed 10 times to BALB/c and maintained in our lab. For primary responses, mice were immunized intraperitoneally with 50 μg of alum precipitated NP-CGG, with a ratio of NP to CGG ranging from 25 to 31. Precipitated alum alone was also used as a control in various experiments. All mice were maintained under SPF conditions and immunized at 6–12 weeks of age. The Yale Institutional Animal Care and Use Committee approved all animal experiments.

Antibodies and detection reagents

The following staining reagents were used: (4-hydroxy-5-iodo-3-nitrophenyl)acetyl (NIP)-binding reagents (NIP-PE and NIP-APC) and mAbs against murine CD3e (2C-11), CD4 (GK1.5), CD90.2 (30H12), CD80 (1610A1) and Kappa (187.1) were prepared as described in (32). Abs against IgG1 (A85-1), CD19 (1D3), CD38 (90), CD162 (2PH1), CD45R (RA3-6B2), CD95 (Jo-2), CXCR5 (2G8), TCRβ (H57-597) and SA-PE-Cy7 were purchased from BD Biosciences. Abs against CCR7 (4B12), PD-1 (J43) and CD62L (Ly-22) were purchased from eBioscience. Streptavidin-Alexa 680 was purchased from Invitrogen, PNA from Vector Labs, and Abs against BrdU (PRB-1; Invitrogen), ICOS (C398.4A; Biolegend), PD-1 (29F.1A12; Biolegend) and TCRβ (H57-597; Biolegend) were also purchased.

Flow cytometry, BrdU detection, apoptosis measurement, and ELISpot and ELISA assays

These were performed as described (11). Occasional mice deemed to be non-responders to immunization were not used in the final analysis of data. Dead cell discrimination was performed using ethidium monoazide (EMA) according to the manufacturer's protocols (Molecular Probes).

Quantitative PCR

Total RNA was isolated from sort-purified populations. cDNA synthesis and qPCR was performed as previously described (33). Primer sequences are as follows: Il21 sense: 5′-TGAAAGCCTGTGGAAGTGCAAACC-3′, an t i-s ense : 5′-AGCAGATTCATCACAGGACACCCA-3′; and actin as stated in (33).

BM chimeras

BM cells from BALB/c or CD80−/− mice and Igh-J−/− mice were mixed in a 20:80 ratio. Cell suspensions were injected into lethally irradiated BALB/c recipients (recipients received total body irradiation, 2× doses of 450 cGy, separated by 3 h, from a 137Cs source). Mice were rested for at least 6 weeks before immunization, and were assessed for extent of chimerism at the time of harvest.

Immunofluorescence histology

Sections were stained with PNA and anti-CD4. Images were taken at a 20× magnification. ImageJ (NIH) and Adobe Photoshop (Adobe) were used for analysis.

Statistics

Statistical analyses was performed using either the Mann-Whitney nonparametric test, or a Student's T test (only where indicated), with two-tailed P-values indicated throughout as: *P < 0.05, **P < 0.01, ***P < 0.001. Two-way ANOVA analysis was also performed for Figure 2.

Figure 2
Memory B cell frequency is increased in the absence of CD80

Results

Decreased long-lived PCs in the absence of CD80

In previous work (20), it was established that circulating Ab concentrations on days 7 and 21 were comparable in CD80−/− and B6 mice. At these times, however, circulating Ab is substantially derived from short-lived plasmablasts rather than long-lived PCs. Therefore, we investigated whether CD80 may be involved in the formation of the long-lived PC compartment. CD80−/− and B6 controls were immunized with NP-CGG and AFCs in spleen and BM were assessed at multiple time-points post-immunization. Consistent with the previous report (20), AFC formation in the absence of CD80 was comparable to that in B6 at day 12 (Fig. 1A). In contrast, long-lived PCs (≥ day 84 post-immunization) in both spleen (Fig. 1B) and BM (Fig. 1C) were substantially decreased in mice lacking CD80. Therefore, CD80-deficient mice have a specific defect in formation and/or maintenance of long-lived PCs.

Figure 1
CD80 has a non-redundant role in PC formation

To determine when the decrease in AFCs was first detectable, we performed a time course during the primary humoral response. In the absence of CD80, both spleen (Fig. 1D) and BM (Fig. 1E) AFCs were decreased at day 15 post-immunization and BM AFCs were decreased compared to controls at all time-points assessed thereafter. Notably, the consistent decrease in BM AFC in CD80-deficient animals was approximately 3-5-fold, indicating that the substantial majority of the normal AFC response failed to form in the absence of CD80. The decrease in AFCs at day 15 correlated with an approximately 3-fold decrease in NP+IgG1+ circulating Ab at this time-point (Fig. 1F), and the decrease in long-lived NP-specific IgG1+ PCs at ≥ 12 weeks post-immunization was also reflected in a concurrent >3-fold decrease of circulating NP-specific IgG1+ Ab (Fig. 1G). Finally, IgM Ab titers were also slightly decreased at these time-points (Supplemental Fig. 1A–B), demonstrating that the decrease in NP-specific IgG1+ Ab and AFCs was not due to a block in isotype switching that would have reciprocally resulted in higher IgM concentrations. Therefore, we conclude that long-lived isotype-switched BM PCs were stably decreased in the absence of CD80 while short-lived responses were largely unaffected.

Enhanced formation of memory B cells in the absence of CD80

PC and memory formation can be considered alternative fates of GC B cells. Given this, we assessed memory B cell formation by flow cytometric analysis in B6 and CD80−/− mice post-immunization with NP-CGG. Memory B cells were identified among live B cells by the phenotype: NIP+IgG1+CD38+kappalo (11). In B6 mice, such cells can be identified as early as day 7 post-immunization (8), though most of these cells evidently do not survive to populate the long-term memory compartment. In B6 mice, the frequency of B cells of a memory phenotype at days 7 and 12 was stable. It then decreased during the contraction phase of the GC response to relatively low levels and appeared to stabilize at ~day 84 (Fig. 2A), as previously observed (11). In the absence of CD80, however, cells of a memory phenotype were increased ~2-fold compared to B6 mice at days 7 and 12 (Fig. 2A). Memory phenotype cells remained increased in CD80-deficient mice over the contraction phase of the GC response and this increase was still evident at least 12 weeks post-immunization (Fig. 2A). Splenocyte numbers were reduced in the absence of CD80; therefore, the increase in memory B cell numbers in CD80−/− mice was not as extreme as the increase in frequency (Fig. 2B). However, increased numbers of memory cells in CD80−/− mice were observed consistently at every time point past day 7; there was a significant difference between the curves between CD80−/− and B6 mice, for both the frequency and number of memory B cells (two-way ANOVA analysis). Lastly, considering that memory and PC differentiation may be alternative fates, we assessed the ratio of memory B cells to PCs, which at ≥week 12 post-immunization was approximately 4-fold greater in the absence of CD80, compared to B6 mice (Supplemental Fig. 2).

Altered GC B cell dynamics in the absence of CD80

The initial characterization of CD80-deficient mice demonstrated by histologic analysis that GC formation can occur in the absence of CD80 at day 7 (20). It has become evident in recent years that GC responses can persist for long periods, which can affect the overall humoral response and subsequent production of long-lived B cell populations (11, 24, 3437). Therefore, we re-assessed the influence of CD80 on both the formation and maintenance of the GC B cell population over time in response to NP-CGG.

By flow cytometric analysis there was no significant difference in either the frequency (Fig. 3A) or number (Fig. 3B) of CD95hiCD38lo GC phenotype B cells in B6 or CD80−/− mice. Thus, the alterations in numbers of memory cells and PCs may be due to alterations in GC B cell differentiation or selection rather than overall homeostasis. To assess whether GC B cell survival may be affecting differentiation into memory B cells and PCs, we investigated apoptosis and proliferation of GC B cells (11, 38). In the absence of CD80, GC B cells underwent increased apoptosis during the height and early contraction phases of the reaction (Fig. 3C). The significant increase in apoptotic GC B cells was also observed when GC B cells were gated for Ag-specificity (data not shown). The increase in apoptotic GC B cells in CD80−/− mice was greater at the later time points, correlating with the observed decrease in AFCs (Fig. 1). The reduction in AFCs, or increase in memory-phenotype cells, may also be affected by differences in proliferation. However, at these same time-points, the fraction of CD80−/− GC B cells in S-phase, as determined after a short pulse with BrdU, was comparable to B6 (Fig. 3D). Together these studies indicate that in the absence of CD80 signaling there was a decrease in survival of GC B cells, particularly late in the response, as well as a decrease in numbers of PCs, while inversely there was an increase in differentiation to memory B cells.

Figure 3
Increased cell death and proliferation in the GCs of CD80−/− mice

B cell-intrinsic role for CD80

CD80 is expressed on many myeloid cells in addition to B cells, and DC expression of CD80 has an important role in T cell activation (39). To investigate whether the role of CD80 in primary humoral responses was due to B–T interactions, or due to defects in T cell interactions with other CD80-expressing APCs, we restricted CD80-deficiency to B cells using mixed BM chimeras. BM from Igh-J−/− mice, which lack B cells, was mixed with BM from BALB/c or CD80−/− mice in an 80:20 ratio, and used to reconstitute lethally irradiated BALB/c mice. The former group of chimeras lacked CD80 in all B cells but could express it in 80% of all other hematopoietically-derived cell types while the latter group served as controls.

When CD80-deficiency was restricted to B cells, a decrease in AFCs in both the spleen (Fig. 4A) and BM (Fig. 4B) was observed at day 25. Chimeras with B cells lacking CD80 were also impaired in BM AFC development at day 15 (Fig. 4D), although there was no significant decrease in splenic AFCs at this early time point (Fig. 4C), As with mice lacking CD80 on all cells (Fig. 3), the frequency of GC B cells was not altered in the absence of CD80 on B cells (Fig. 4E). Interestingly, memory B cell frequency was also comparable to controls in the BM chimeras, suggesting that the absence of CD80 specifically on B cells does not impair memory B cell development (Fig. 4F). In summary, the decrease in long-lived AFCs observed in the absence of CD80 was reproduced when only B cells lacked CD80 expression.

Figure 4
B cell intrinsic role for regulation of B cell differentiation

TFH cells are decreased and functionally altered in the absence of CD80

Interactions between B cells and TFH cells are critical for the GC, particularly for sustaining the reaction (7, 11, 36, 37). We assessed TFH frequency and number to investigate whether CD80 is required for formation of this population. In the absence of CD80, there was a decrease in the frequency of TFH-phenotype cells at day 12 post-immunization (Fig. 5A). To investigate whether there was a deficiency in generation of TFH cells, we determined the numbers of TFH cells at day 7 post-immunization, as assessed by gating on PD-1hi CD4+CD44hi cells that also either increased CXCR5 expression or decreased CD162 downregulation (40). In the absence of CD80, there was a significant decrease in the numbers of TFH cells compared to B6 mice (Fig. 5B). Both types of analysis were used in the time course, which showed that TFH cell frequency was decreased in CD80−/− relative to B6 mice at all time-points assessed, including early, peak and contraction phases of the GC response (Fig. 5C). TFH cells can be identified by flow cytometry, but not all of them are evidently in the GC (41). Thus, to extend the flow cytometric analysis, we also investigated the presence of T cells within immunohistologically defined GCs of both B6 and CD80-deficient mice. (Fig. 5D, E; Supplemental Fig. 3). At day 15 post-immunization, there were fewer CD4+ T cells within PNA+ GCs in CD80−/−, compared to B6 mice. This difference was also seen at days 12 and 19 (Fig. 5E). Therefore, in the absence of CD80, there was impaired development of TFH cells both as assessed by expression of typical cell surface markers as well as by identification directly in GCs.

Figure 5
Reduced frequency of cells of a TFH phenotype in the absence of CD80

Interestingly, we noted that, in the absence of CD80, there was also decreased surface expression of the commonly used markers of TFH, PD-1 (Fig. 6A–B) and ICOS (Fig. 6A, C) on TFH cells. The reduced expression of these CD28 family members, which normally occurs early in the response, suggested that the TFH cells in CD80−/− mice had not completed maturation. To further investigate whether the perturbations observed in TFH development extended beyond phenotypic alterations, we also assessed proliferation (Fig. 6D) and death (Fig. 6E) of TFH cells. Interestingly, a higher fraction of TFH cells was proliferating in the absence of CD80 (Fig. 6D). However, the rates of proliferation were quite small, with only ~1% of TFH cells in S phase (as determined after a 2hr pulse of BrdU) in B6 and ~2–3% in CD80−/− mice. Conversely, a higher percentage of TFH cells were undergoing cell death in the absence of CD80 (Fig. 6E). Thus, although TFH cells in CD80-deficient mice had a higher proliferative rate than those in B6 mice, they also underwent more cell death. Given the net reduction in TFH cell numbers at steady state, presumably the higher apoptotic rate overcomes the increase in proliferation, resulting in an overall decrease in TFH frequency. These studies overall demonstrate that CD80 normally influences both the quality and homeostasis of TFH.

Figure 6
Alterations in TFH phenotype, proliferation and survival in the absence of CD80

CD80 expression on B cells regulates TFH frequency and phenotype

To assess whether B-T interactions were the cause of the defective maturation and turnover of TFH cells, we assessed the frequency and number of TFH cells in mixed BM chimeras lacking CD80 on B cells. When CD80 deficiency was restricted to B cells, TFH frequency was decreased, compared to BALB/c (Fig. 7A), and the MFI of PD-1 was also significantly decreased at day 15 (Fig. 7B). Therefore, direct B-T interaction via CD80 on B cells is responsible for maintenance of TFH cell number and quality, as evidenced by effects on expression of PD-1, a key TFH surface protein that has been shown to influence TFH function (11).

Figure 7
CD80 regulates T cell phenotype, frequency and function

CD80 regulates TFH function and subsequent Bcl-6 expression by GC B cells

TFH cells sustain GC B cell development at least in part through the production of cytokines, in particular IL-21 (35, 37). To assess this function, we used FACS to purify T cells from CD80−/− mice and their B6 controls and performed qPCR analysis to compare amounts of Il21 mRNA. Il21 mRNA expression was decreased in CD80−/− TFH cells compared to B6 (Supplemental Fig. 4). Although the reduction in transcript quantities was not as great as observed in PD-1−/− mice (11), the lower number of TFH cells in CD80−/− mice would amplify the difference in overall availability of IL-21 between B6 and CD80−/− mice. IL-21 has recently been demonstrated to maintain Bcl-6 expression in GC B cells (35, 37). Indeed, lower Il21 transcript amounts correlate with lower Bcl-6 expression in GC B cells from CD80-deficient BM chimeras at day 15 post-immunization, a time point when GC B cells are known to be IL-21 sensitive (27, 37) (Fig. 7C). Therefore, B cell expression of CD80 is important for maintenance of both TFH and GC B cell phenotypes and functions.

Discussion

Taken together, the data presented here indicate that CD80 has non-redundant roles in TFH development, GC B cell survival and long-lived PC development during primary humoral responses. As many types of APC can express CD80, a critical finding was that chimeric mice lacking CD80 on all B cells, but with normal expression patterns on non-B cells, showed the key phenotypes of the mice lacking CD80 in all cells. This allowed us to deduce B cell-intrinsic role for CD80, suggesting that it regulates survival and differentiation of both B and T cells within the GC through ongoing B-T interactions. These effects were seen at distinct times post-immunization: alteration in TFH numbers and surface marker expression levels were observed by day 7, and heightened GC apoptosis and decreased PC formation were observed immediately after the peak of the GC response and beyond. Furthermore, defects in humoral responses in the absence of CD80 were observed in both B6 and BALB/c mice, and at multiple time-points. Thus, in concert with results from studies on PD-1 signaling in the humoral response (11, 42) a biological theme is emerging that certain B7 family members regulate the later phases of GC function and thus control the size and quality of the long-lived humoral immune compartment.

Recently, two papers have addressed the roles of CD80, CD86 and CD28 in the humoral immune response. Salek-Ardakani and colleagues (27) used vaccinia virus and came to conclusions very similar to several of the original study on CD80 and CD86 deficiency (20): that CD86 and CD28 are required for GC and Ab formation, whereas CD80 was not. As noted above, the phenotype we observed pertains to the late stage of the GC and the long-lived AFCs that are generated there. Neither the much earlier (20) nor this more recent (27) study systematically assessed these time points and compartments. A second study reported the intriguing finding that CD28 expression in the B cell lineage optimized, but was not absolutely required for, plasma cell generation (43). In agreement with our findings, they observed decreases in PCs in the absence of CD80, as well as CD86 or CD28. However, in contrast to our findings, they attributed this to CD28 expressed on PCs which they argued sustains PC survival in the BM via interactions with CD80 or CD86 on non-B cells. Hence, this mechanism differs from the one we have documented in several ways: a) our data show a non-redundant role for CD80, whereas both CD80 and CD86 play similar roles in the system of Rozanski et al. (43); b) we demonstrate a B cell intrinsic role for CD80 whereas Rozanski et al. implicate a B cell-extrinsic role for either CD80 or CD86; and c) we trace our phenotype to the late GC, during which TFH numbers and development are affected, whereas in Rozanski et al. the effects are thought to affect post-GC long-lived PCs only.

Diverse role of negative regulators in GC, memory cell, and AFC development

Since CD80 is dispensable for promoting GC development but CD86 and CD28 are required (20, 21, 27), it is logical to propose that the phenotypes observed here in CD80-deficient mice may relate to interactions of CD80 with one or both of its other binding partners, CTLA-4 and PD-L1. Both of these interactions are known to produce “negative” signaling outcomes (44). Negative regulators act later in the GC response than their co-stimulatory counterparts. Mice deficient in these “co-inhibitory” pathways share some phenotypes, probably due to similar mechanisms of action of receptors that transduce negative signals. In mice deficient in PD-1 signaling (11) or CD80, similar findings included: substantial reductions in long-lived AFCs, particularly in the BM, a higher fraction of apoptotic GC cells, and decreased IL-21 mRNA transcript by TFH cells.

Although the shared phenotypes suggest connections between the pathways and/or shared modes of signaling (e.g. between PD-1 and CTLA-4 or PD-L1), the different phenotypes observed in the PD-L2−/− (11), PD-L1−/− (42), and CD80−/− mice indicate that there are specific and unique roles for each pathway. In the absence of PD-1 signals, there was an increase in T cells with a TFH phenotype; in contrast, in the absence of CD80, there was a decrease in T cells with a TFH phenotype. There was also an increase in frequency of memory B cells in both CD80−/− and PD-L2−/− mice that was not observed in the other gene-deficient mice. While memory cell and AFC generation were inversely affected with the loss of CD80, PD-L2, or IL-21 signals (35, 37), PD-1−/− mice had reduced numbers of both cell types (11). Thus, the generation of memory B cells and AFCs are sometimes but not always inversely linked and these pathways are differentially controlled. Unfortunately, very little is known overall about mechanisms that control memory B cell formation; therefore much more research is required to determine how PD-L2 and CD80 influence memory B cell production.

The possible explanations for both similarities and differences in the phenotypes of mice deficient in B7:CD28 family negative regulators are complex, given the many different potential interactions that these receptors can have. One common factor downstream of all these pathways is IL-21 secreted by TFH. Recent studies have shown that IL-21R signaling in GC B cells is critical in the late phase of the GC reaction and in its absence GCs do not maintain their size and the AFC compartment size is reduced (35, 37). Optimal production of IL-21 and possibly other cytokines (45, 46) by TFH could therefore be one mechanism by which the B7 family co-inhibitory pathways have similar effects on the GC and AFC compartments, although without the ability to delete specific ligands (i.e. CD80) and specific receptors (i.e. CD28 and CTLA4) in GC B and T cells at appropriate time points, we must point out that the TFH and B cell phenotypes are correlative in our study. Although we favor the explanation that the reduced numbers and qualitative TFH defects we demonstrated are actually causative of the loss of AFCs in CD80-deficient mice, it remains possible that reverse signaling in B cells is instead responsible for failed B cell differentiation into AFCs. If so, then CD80-deficient B cells would still fail to form AFCs even in a 50:50 mixed BM chimera, in which recipients are reconstituted with both wild-type and CD80-deficient BM. In this system, TFH would get signals from wild-type B cells and would likely be intact but reverse signaling would be absent in CD80-deficient B cells nonetheless. Interestingly, in a previous study by Lumsden and colleagues (47), the authors performed the above experiment with CD80 and CD86 double-deficient BM and demonstrated that there was no cell-intrinsic requirement for CD80 and CD86 provided that other B cells express it in trans. This argues against B cell-intrinsic reverse signaling and instead is consistent with the interpretation that reduction of TFH in CD80-deficient mice affect the ability of these mice to produce long-lived PCs.

A clear difference between the PD-1 pathways and CD80 is that CD80, unlike PD-L1 or PD-L2, can also engage the stimulatory receptor CD28. If stimulatory receptor interactions promote TFH expansion, then failure of CD80 to contribute to CD28 engagement might account for decreased numbers of TFH in the absence of CD80. In contrast, the absence of PD-1, which does not engage any known stimulatory signals, leads to an increase in TFH numbers. In addition, CD80 can interact with PD-L1, as well as CD28 and CTLA-4; therefore, there could be crosstalk between the CD80 and PD-1 mediated pathways. For example, CD80-PD-L1 binding could sequester each other thereby reducing PD-L1-PD-1 interactions. Finally, in addition to cross-talk mediated by direct binding interactions with PD-L1, CD80 affects the expression of PD-1 (Figures 6 and and7),7), and hence indirectly controls the PD-1 pathway.

Despite these interactions of CD80 with PD-1 signaling, CD80 cannot interfere directly with PDL2. Yet, PD-L2−/− mice also have a clear phenotype that recapitulates many (but not all) aspects of the phenotype of PD-1−/− mice, indicating that PD-1 signals themselves must be important independent of CD80. Thus PD-1 and CD80 pathways must have at least some non-redundant functions.

Why are negative regulators required for fully functional humoral responses?

It is interesting to consider how pathways that, in other contexts are thought to negatively regulate responses, are required in the context of humoral immunity to sustain the GC response and development of long-lived AFCs. We suggest that these pathways could be viewed as turning off the proliferative phase of the early primary response and favoring differentiation instead. Notably, PC development requires signals that interrupt proliferation and cell cycling, as demonstrated by impaired PC differentiation in B cells lacking the CDK inhibitor p18 (48). Signals that interrupt proliferation could occur by reverse signaling of PD-L1 or PD-L2 in GC B cells or indirectly due to lack of proliferative and survival signals coming from TFH. Present and prior data show that negative regulators promote TFH maturation and in particular IL-21 mRNA expression; IL-21 is well documented to promote PC differentiation (49, 50).

The complexity of the network that has evolved to regulate the late GC response emphasizes that the quality and quantity of the long-lived phase of humoral immunity must be carefully regulated to promote optimal long-term immunity. Niches for both memory cells and PCs are likely to be limited (51). A far greater number of precursors capable of entering these pools is generated than actually survives long-term. Further, stringent affinity-based selection governs the quality of these cellular compartments. Together with other publications, the current work substantially expands the concept of B7-family based regulation of the output and quality of the GC and the long-lived AFC pool, revealing a unique function for CD80 and uncovering effects of CD80 on generation and function of TFH, GC cells and AFCs. These studies thus enhance our understanding of the development and function of long-lived humoral immunity.

Supplementary Material

ACKNOWLEDGMENTS

We thank S. Michalek for CD80−/− mice and samples; the Yale Cell Sorter Facility for cell sorting; the Yale Animal Resource Center for animal care; and Dr. Steven Kerfoot for critical reading of the manuscript.

ABBREVIATIONS

(AFC)
Antibody-forming cell
(EMA)
ethidium monoazide
(GC)
germinal center
(PC)
plasma cell
(TFH)
T follicular helper

Footnotes

1Supported by National Institutes of Health (AI43603 to M.J.S.) and AI56299 (to A.H.S), the National Health and Medical Research Council (K.L.G.-J.) and Arthritis Australia (K.L.G.-J.).

AUTHOR CONTRIBUTIONS K.L.G.-J., S.M.A and M.J.S. designed research; K.L.G.-J., S.M.A. and E.S. did research; A.S. generated and contributed knockout mice and advice; and K.L.G.-J. and M.J.S. analyzed data and wrote the manuscript.

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