Antibody-Independent Function of Human B Cells Contributes to Antifungal T Cell Responses

Candida albicans is a commensal microorganism frequently found on the skin and in the gastrointestinal tract of healthy individuals (1). Overgrowth of C. albicans can cause candidiasis, which usually happens in the immunocompromised host whose B cells and T cells were usually both affected, such as in individuals with HIV/AIDS or those treated with immunosuppressant medications (2, 3). T cells, particularly Th1 and Th17 cells, play an important role in controlling and clearing fungal pathogens, including C. albicans (4–10). Patients with inborn deficiency of either Th1- or Th17-associated molecules exhibit increased incidence of fungal infections (11–18). Similarly, mice deficient in IL-12/IFN-γ or IL-23/IL-17 are susceptible to C. albicans infection (4, 5, 7, 19–21). Additionally, adoptive transfer of wild-type CD4⁺ T cells but not IL-17^−/− CD4⁺ T cells confer antifungal protective immune responses in recipient mice (5, 8). The role of B cells in combating C. albicans has been less clear. In experimental models, B cell–deficient mice exhibit increased susceptibility to experimental systemic candidiasis and, in humans, several case reports have described unusual fungal infections after B cell depletion (22–34). In a cohort of patients with diffuse large B cell lymphoma treated with rituximab, Lin et al. (26) observed that elderly patients tended to have a high rate of fungal infection. Consistent with this observation, Kamar et al. (35) compared the incidence of infections in 77 patients receiving B cell depletion therapy (BCDT) after kidney transplantation with 909 control patients without B cell depletion and found that the rate of fungal infections was significantly higher in the BCDT group, whereas the rate of bacterial infection was similar between two groups. Interestingly, the increase of fungal infection rate was strongly associated with T cell counts in the circulation after BCDT. Although B cells are best known for their role of producing pathogen-specific host-protective Abs during infection, patients with agammaglobulinemia surprisingly exhibit quite normal antifungal immunity (36), suggesting that human B cells do contribute to Candida control, although through some Ab-independent mechanism.

Recent studies have greatly advanced our understanding of Ab-independent roles of B cells in the regulation of cellular immune responses during both infectious and autoimmune conditions (37–40). These B cell Ab-independent functions may include Ag presentation to T cells (TCR/MHC signals) (41, 42), provision of costimulatory signals to T cells (costimulatory signals) (43, 44), and secretion of cytokines that can further shape T cell differentiation (cytokine signals) (37, 39, 42, 45–48). Expression of regulatory cytokines such as IL-10 and IL-35 from B cells has been shown to downregulate host-protective immune responses, thereby facilitating pathogen invasion (46). It remains unknown, however, whether B cells can promote host-protective (antipathogen) responses through Ab-independent mechanisms such as, for example, through expression of proinflammatory cytokines.

To examine the potential role for human B cells in Ab-independent antifungal responses, we first developed a novel pathogen-dependent, MHC-restricted human B cell/T cell coculture system. Using this system, we show that human B cells can present fungal Ag to Th cells in an HLA-DR–restricted manner, provide critical costimulatory signals through CD80 and CD86, and induce Th17 cell differentiation through an IL-6–dependent mechanism. We further demonstrate that depleting B cells both in vitro and in vivo in humans can result in reduced human T cell responses to C. albicans. Taken together, our findings indicate that Ab-independent functions of B cells can contribute to antifungal immunity through regulation of Ag-specific T cell activation and their proinflammatory differentiation.

All subjects provided written informed consent as approved by the Montreal Neurological Institute and Hospital Ethics Review Board. Healthy subjects were recruited from the Montreal Neurological Institute, McGill University. Rituximab-treated patients were recruited at the multiple sclerosis (MS) Clinic of the Montreal Neurological Institute and Hospital, and all had relapsing–remitting MS as defined by the McDonald criteria (49), were free of injectable immune-modulating treatment and steroid exposure for at least 3 mo prior to the baseline sampling, and had no prior exposure to oral MS therapies, mAb, i.v. Ig, plasmapheresis, or chemotherapy. We excluded patients with secondary or primary progressive MS, disease duration of >15 y, or a known history or presence of other neurologic or systemic autoimmune disorders. The age range of the patients included was 39–54 y at the time of B cell depletion, their female/male ratio was 3:1, and the average relapsing–remitting MS duration from diagnosis was 8 y.

Human PBMC were separated by density centrifugation using Ficoll (GE Healthcare). CD19 beads were used to positively select B cells based on the manufacturer’s protocol. Typical purities routinely assessed by flow cytometry for B cells were >98% (Supplemental Fig. 1A). CD3⁺CD4⁺CD25⁻ responder T (Tresp) cells, as well as naive T cell subsets (CD3⁺CD4⁺CD45RA⁺CD45RO⁻), were all isolated using negative selection (Miltenyi Biotec, Auburn, CA) with confirmed purities of >98% (Supplemental Fig. 1B–D). All cells were cultured in serum-free X-VIVO medium (Life Technologies). B cells were then plated in U-bottom 96-well plates at 2 × 10⁵ cells per well in a total volume of 200 μl of medium. T cells were added to cocultures at 1 × 10⁵ cells per well.

Reagents used to activate B cells included soluble CD40L (1 μg/ml; Enzo Life Sciences), goat anti-human BCR F(ab′)₂ fragment Ab (10 μg/ml; Jackson ImmunoResearch Laboratories), CpG DNA (1 μM/ml, ODN2006; InvivoGen), Pam3CSK4 (1 μg/ml; InvivoGen), LPS (1 μg/ml, 0111:B4; Sigma-Aldrich), IL-4 (20 ng/ml; R&D Systems), IFN-γ (10 ng/ml; R&D Systems), IL-17 (1 ng/ml; R&D Systems), TGF-β (4 ng/ml; R&D Systems), IL-12 (20 ng/ml; R&D Systems), IL-21 (20 ng/ml; R&D Systems), and IL-13 (20 ng/ml; R&D Systems). Reagents used to simulate T cells included plate-bound anti-CD3 (UCHT1, 1 μg/ml), endotoxin-free protein extracts from C. albicans (M15, 1 μg/ml; Greer Laboratories), or Staphylococcus aureus (tlrl-hksa, 10⁸/ml; InvivoGen). CFSE was used, as per the manufacturer’s protocol, to quantify T cell proliferation. Briefly, isolated T cells were diluted in prewarmed X-VIVO medium at 1 × 10⁶ cells/ml with 1 μM CFSE slowly added to the cell suspension. Cells were then incubated at 37°C in a water bath for 20 min, washed twice, and then plated into 96-well U-bottom plates, as described above. Function-blocking Abs (1 μg/ml) targeting CD80 (37711), CD86 (199622), and IL-6R (17506) and matched control Abs were purchased from R&D Systems. HLA-DR (G46-6) function-blocking Ab and matched control Ab were purchased from BD Biosciences.

To generate B cell culture supernatant from activated B cells, B cells (at the concentration of 1 × 10⁶/ml) were first stimulated with CD40L plus anti-BCR plus IL-4 for 24 h, and then after two washes with fresh culture medium, B cells were replated in a U-bottom 96-well plate. B cell culture supernatants were collected after an additional 2 d.

Flow cytometry phenotyping of B cell and T cell subsets, as well as T cell proliferation and their intracellular cytokine staining (ICS), was accomplished with Abs targeting CD3 (SK7), CD4 (RPA-T4), CD8 (RPA-T8), CD20 (2H7), CD80 (BB1), CD86 (2331 [Fun-1]), IL-17A (N49-653), and IFN-γ (B27). All Abs were from BD Biosciences. To assess ICS, PMA (20 ng/ml; Sigma-Aldrich), ionomycin (500 ng/ml; Sigma-Aldrich), and GolgiStop (monensin; BD Biosciences) were added to cells 4–5 h before staining. Cells were then stained with Live/Dead marker (Life Technologies) at room temperature, after which cell-surface marker staining was performed. Cells were then fixed and permeabilized using Fixation/Permeabilization buffer (BD Biosciences). ICS Abs (noted above) were added and incubated for 30 min. Samples were then washed twice and analyzed by FACSCalibur or LSRFortessa (BD Biosciences).

Cytokine levels in culture supernatants were measured by an OptEIA ELISA kit (IFN-γ and IL-6; BD Biosciences) or an ELISA Ready-SET-Go! kit (IL-17A; eBioscience), following the manufacturers’ protocols. Briefly, ELISA plates were coated with capture Ab at least 12 h in advance. After another hour of blocking with blocking buffer (10% FCS, PBS), samples were added to the plate and incubated for 2 h at room temperature. Then, detection Abs were added and incubated for 1 h at room temperature. The plate was carefully washed with ELISA washing buffer (0.05% Tween 20, PBS) between each step. The color of the plate was eventually developed by tetramethylbenzidine (BD Biosciences) and the reaction was stopped by 0.01 N H₂SO₄. Finally, the plates were read by a Bio-Rad Laboratories microplate reader (model 550).

All values are expressed as mean ± SEM, and p values were assessed as appropriate by either a Student t test or one-way ANOVA, with a Tukey post hoc test using GraphPad Prism version 6. A p value <0.05 was considered significant.

To assess the capacity of human B cells to induce antipathogen T cell responses, we first established a novel coculture system in which human CD4⁺CD25⁻ Tresp cells are incubated with autologous freshly purified CD19⁺ B cells that were pulsed with either vehicle control, protein extracts from heat-killed C. albicans, or heat-killed S. aureus as another form of control. Because the Tresp population would be expected to include both naive and memory T cells, we also sorted naive (CD45RO⁻CD45RA⁺CD4⁺CD3⁺) T cells to directly assess de novo T cell responses in our system. We found that pathogen-pulsed B cells (whether with C. albicans or S. aureus) significantly enhanced proliferative responses of the CD4⁺ Tresp cells (Fig. 1A, 1B) as well as naive CD4⁺ T cells (Fig. 1C, 1D). C. albicans appeared to induce less naive T cell proliferation than did S. aureus, and, unlike S. aureus (which can function as a superantigen), T cells exposed to C. albicans alone did not proliferate (data not shown). Memory T cells were more responsive to C. albicans in the presence of B cells as compared with naive T cells (Supplemental Fig. 2). The naive CD4⁺ T cell responses to C. albicans–pulsed B cells were MHC restricted, as functional blockade of HLA-DR essentially abrogated their pathogen-induced proliferative responses (Fig. 1E, 1F).

Next, we tested the phenotype of B cells in the B cell/T cell coculture system in the presence of C. albicans. Human purified B cells were labeled with CFSE and cocultured with CD4⁺CD25⁻ T cells. At day 3, expression of costimulatory molecules (CD80 and CD86) and cytokines (TNF-α, IL-6, and GM-CSF) was assessed to characterize the phenotype of activated (proliferating) B cells. As compared with nonproliferating cells, proliferating B cells exhibited significantly higher CD80, CD86, IL-6, and GM-CSF expression (Fig. 2A–E). IL-10 expression by B cells was not different between proliferating and nonproliferating B cells (data not shown). This suggested that B cells are also activated in the coculture system and that the responding B cells (proliferating B cells) acquire the ability to upregulate immune responses. At day 10, proliferating B cells became plasmablasts after the coculture with T cells in presence of C. albicans (Fig. 2F).

Next, we wanted to test whether activated B cells would exhibit a higher capacity to induce C. albicans–specific T cell responses. We either left B cells resting or preactivated them with CD40L plus anti-BCR plus IL-4 for 24 h and then irradiated both the resting B cells and activated B cells (to prevent the confounder of B cell proliferation), before coculturing the B cells with T cells in the presence of C. albicans. Compared to resting B cells, activated B cells strongly promoted C. albicans–specific T cell proliferation (Fig. 3A, 3B) and cytokine production (IFN-γ, Fig. 3C–E; IL-17, Fig. 3F–H). These effects of activated B cells were highly dependent on the costimulatory molecules CD80 and CD86 expression, because neutralizing both significantly decreased the C. albicans–specific T cell responses (Fig. 3).

To assess whether C. albicans–dependent B cell–mediated activation of human T cells induces de novo differentiation of Th1 or Th17 from naive CD4⁺ T cells, we measured IFN-γ (Th1) and IL-17 (Th17) secretion in culture supernatants (ELISA; Fig. 4A, 4B), as well as within the cells in coculture (ICS and flow cytometry; Fig. 4C–E). Whereas B cell–mediated Ag-dependent activation of naive T cells whether with C. albicans or S. aureus induced both Th1 and Th17 cytokine responses, we observed an Ag-specific reciprocal induction of IFN-γ and IL-17 responses. Whereas S. aureus induced a more limited 2-fold increase in IL-17 but a considerably more substantial 30-fold increase in IFN-γ, C. albicans induced similar fold increases of IFN-γ and IL-17A (Fig. 4A, 4B). ICS and gating on the naive T cells in coculture confirmed that T cells (Fig. 4C–E), but not B cells (data not shown), were the source of the reciprocally regulated cytokines.

To test the signaling pathway that governs the de novo induction of Th1 and Th17 responses, we tested the expression of pSTAT1/T-bet and pSTAT3/ RORγt in naive T cells. We found that B cells significantly upregulated pSTAT1/T-bet (Supplemental Fig. 3) and pSTAT3/RORγt (Fig. 4F, 4G) pathways in naive T cells. Blockade of IL-12 and IL-27 (known as Th1-promoting cytokines), either alone or in combination, did not alter T cell IFN-γ expression induced by the B cells (Supplemental Fig. 4). Because naive T cell differentiation into Th17 cells is known to depend on IL-6–mediated activation of the STAT3/RORγt signaling pathway (50), and B cell IL-6–specific knockout mice exhibit a deficiency in Th17 responses (42, 45), we considered whether IL-6 from the C. albicans–pulsed human B cells could contribute to the observed induction of human Th17 responses. Indeed, blocking IL-6 signaling with a function-blocking Ab against the IL-6R significantly decreased the capacity of activated C. albicans–pulsed B cells to induce expression of T cell pSTAT3 (Fig. 4F), as well as RORγt (Fig. 4G) and IL-17 (Fig. 4H). In experiments further exploring the induction of IL-6 by human B cells, we noted that both adaptive and innate stimuli can induce IL-6 (Fig. 5A, 5B) and that the IL-6 secretion could be modulated (either increased or decreased) in the presence of different exogenous cytokines (Fig. 5C). The addition of IL-4 induced particularly high levels of IL-6 production (Fig. 5C), an effect mainly mediated through IL-4R (but not CD132) (Fig. 5D).

The data above demonstrated that CD80/CD86 and IL-6 expressed by B cells support C. albicans–associated T cell responses. Next, we tested whether a particular B cell subset is implicated in mediating the C. albicans–associated T cell activation. Cytokine-defined B cell responses are recognized to play important roles in autoimmunity and infectious diseases (48). We previously demonstrated that human IL-6⁺ B cells and GM-CSF⁺ B cells can upregulate proinflammatory responses of T cells and myeloid cells, respectively. In our present B cell/T cell coculture system, we were able to identify three B cell subpopulations based on the expression of IL-6 and GM-CSF (i.e., IL-6⁻GM-CSF⁻, IL-6⁺GM-CSF⁻, and IL-6⁺GM-CSF⁺; Fig. 6A). We noted that >50% of proliferating B cells in our B cell/T cell Candida coculture system were double cytokine–expressing (IL-6⁺GM-CSF⁺) B cells (Fig. 6B). Of note, CD80, CD86, and TNF-α were selectively induced in the IL-6⁺GM-CSF⁺ B cells (Fig. 6C–E). Taken together, these data suggest that IL-6⁺GM-CSF⁺ B cells are a major B cell subset responding to C. albicans, and the expression of CD80 and CD86 by this subset provides important costimulatory signals to the responding T cells.

Our findings suggested that B cells can contribute to C. albicans–mediated T cell responses. To test whether the presence of B cells is necessary for optimal host T cell responses to C. albicans, we first depleted B cells from PBMC in vitro (using CD19 microbeads) and compared T cell responses to C. albicans between B cell–depleted PBMC and whole PBMC. In vitro B cell depletion resulted in a substantial decrease of T cell proliferation as well as cytokine production (Fig. 7A–D). To examine whether this could also happen in vivo, we next took advantage of accessing PBMC from patients who underwent B cell depletion therapy as part of an anti-CD20 (rituximab) treatment trial for MS. Cryopreserved PBMC isolated from the same MS patients prior to and following B cell depletion were thawed and stimulated, as above. We found that T cell proliferative responses, as well as both IFN-γ and IL-17 production in response to C. albicans, were significantly decreased after in vivo B cell depletion (Fig. 7E–I). Interestingly, adding back B cell supernatants (generated from the same patients prior to the B cell depletion) reconstituted the T cell production of IL-17 in an IL-6–dependent manner (Fig. 7G–I). Taken together, these data corroborate the concept that the IL-6 from B cells is important for B cell–mediated generation of fungi-specific Th17 responses and is relevant to the Ab-independent role of B cells during fungal infections.

In this study, we first established a novel Ag-specific human B and T cell coculture system. Using this system, we could demonstrate that human B cells are able to mediate fungal-associated Ag-specific activation of T cells in an MHC class II–restricted and costimulatory molecule (CD80 and CD86)–dependent manner. We further demonstrated that IL-6 from B cells could induce differentiation of Th17 antifungal responses from naive T cells. In vivo, depletion of B cells in humans resulted in reduced C. albicans–associated T cell responses.

Studies of human B cell–T cell interactions have historically been challenging. One common approach has been to coculture human B cells with T cells that are polyclonally activated (such as with anti-CD3, or anti-CD3 together with anti-CD28). However, during infection or autoimmune disease, T cells are generally thought to be activated in an Ag-specific, MHC-restricted manner that is driven by a relatively limited number of Ags (51). Assessing more restricted Ag-specific human B cell–T cell interactions has been limited by the low precursor frequencies of both B cells and T cells to the particular Ag. To some extent, this can be overcome in the context of vaccinations such as with tetanus toxoid, where precursor frequencies of both B cells and T cells reactive to tetanus toxoid are induced in vivo, making subsequent in vitro coculture studies of circulating Ag-specific T cells and B cells more practical (52). This approach, however, is limited to the few pathogens routinely used in human vaccination. Compared to these existing systems, our novel approach does not require prevaccination and uses microorganisms, which results in very robust Ag-specific proliferative and cytokine responses.

Both mice and human studies have revealed that Th1 and Th17 cells are actively involved in antifungal immunity (4–7, 9, 17, 53, 54). Optimal Ag-specific T cell responses require help from APC (55–57). Dendritic cells and myeloid cells are traditionally viewed as classic APC during infection (55, 57), although recent studies have expanded the definition of APC (56, 58). B cells can also act as potent APC, especially when the concentration of Ags is low (41, 42, 52, 58–60). However, this function of B cells has never been tested in the context of antifungal responses. Prior work done by van de Veerdonk et al. (54) suggested that B cells may contribute to the antifungal host defense. The authors showed that in vitro addition of rituximab (versus control Ig) to PBMC reduced anti-Candida responses. Our study first reinforces these findings by demonstrating that in vitro B cell depletion results in diminished T cell responses to Candida, and then further demonstrates that in vivo treatment with rituximab in patients results in reduced ex vivo Ag-specific responses to Candida, suggesting that human B cells can play a nonredundant Ag-presenting role during C. albicans infection.

IL-6 is known to induce Th17 cell differentiation from naive T cells both in mice and in humans (50). IL-6^−/− mice are more susceptible to C. albicans (61). Several groups including ours have shown that murine B cells can provide an important source of IL-6 for in vivo Th17 cell differentiation (42, 45). Our present study extends these observations by demonstrating that human B cell–derived IL-6 can contribute to antifungal host protection by inducing Th17 responses. Previous work from our laboratory identified a subset of GM-CSF⁺ B cells that are high producers of IL-6 and TNF-α (48, 62). The in vitro activation condition that promotes B cell GM-CSF production (CD40L plus anti-BCR plus IL-4) also strongly induces the production of IL-6 and TNF-α from human B cells (62). This is of interest, as all three cytokines have been shown to play important roles during Th17-mediated host responses to C. albicans infection (61, 63, 64). Although IL-6 is important in inducing de novo Th17 cell differentiation, both GM-CSF and TNF-α have been shown to prime myeloid cells that enhanced Th17 fungal-specific T cell responses (63, 64), in part through IL-23–mediated maintenance of Th17 cells (63). In turn, Th17 responses (and GM-CSF itself) are known to activate and recruit neutrophils that play important roles in clearance of the fungi (65). The human GM-CSF–expressing B cells were also found to express high levels of the costimulatory molecules CD80 and CD86 (62). Collectively, these observations suggest that GM-CSF⁺TNF-α^highIL-6⁺ B cells, as Th17 cells do in T cells, may constitute a B cell subset with particularly important roles in antifungal immunity.

To date, the increased incidence of fungal infections such as Pneumocystis jirovecii following BCDT has largely been limited to elderly lymphoma patients, or those who are immunocompromised in the context of BCDT preceding kidney transplantation (22–35). In contrast, fungal infections have not emerged as a common complication of BCDT during the anti-CD20 clinical trial programs in autoimmune diseases, such as MS. One possible reason is that these patients tend to be younger when treated with BCDT. Indeed, older age is an established risk factor for fungal infection with or without BCDT (26). It is also possible that the abnormally increased proinflammatory T cell (including Th1 and Th17) responses implicated in MS and other autoimmune diseases may in fact protect from emergence of fungal infections even when the contribution of B cells to antifungal immunity is diminished. Importantly, unlike the situation in lymphoma treatment and transplantation (in which BCDT is typically a time-limited intervention), the emerging prospect of long-term BCDT in patients with chronic autoimmune diseases raises concern that fungal infections will become a considerably more common treatment-emergent complication in such populations, especially as they age. In the context of MS (for which approval of BCDT is imminent), an added concern is that many patients who will be offered BCDT will have been previously treated with immune-modulating therapies, which may have lingering effects on T cell responses. In such patients, the combination of BCDT with even partial deficits in Th1 and Th17 responses may manifest with insufficient antifungal responses.

Taken together, our findings both underscore the growing appreciation of Ab-independent roles of B cells, including their implication in normal host-protective responses, and alert us to the possibility that the incidence of BCDT-related infectious complications may increase as chronically treated patient cohorts age, and as more patients are exposed to serial combination therapy.

Principal investigators: Amit Bar-Or,^1,2,3 Alexandre Prat,⁴ and Jennifer Gommerman⁵

Members: Ayman Rezk,^1,2 Boli Fan,³ Craig Moore,¹ Dennis Lee,⁵ Farah Jalili,¹ Georgina Galicia Rosas,⁵ Gillian Muirhead,^1,2 Hanane Touil,^1,2 Jen Yam,⁵ Laila Al Alwan,^1,2 Laura Michel,⁴ Luke Healy,^1,2 Malwina Mencel,^1,2 Natalia Pikor,⁵ Olga Rojas,⁵ and Rui Li^1,2

¹Neuroimmunology Unit, Montreal Neurological Institute, McGill University, Montreal, Quebec H3A 2B4, Canada; ²Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA 19104; ³Experimental Therapeutics Program, Montreal Neurological Institute, McGill University, Montreal, Quebec H3A 2B4, Canada; ⁴Centre de Recherche du Centre Hospitalier de l’Université de Montréal–Hôpital Notre-Dame, Montreal, Quebec H2L 4M1, Canada; and ⁵Department of Immunology, University of Toronto, Toronto, Ontario M5S 1A8, Canada.

We thank all members of the Canadian B Cells in MS Team. We also appreciate the important input of the experimental therapeutic program at the Montreal Neurological Institute for judicious handling of patient and control samples.

The authors have no financial conflicts of interest.