Research Article

IgA production requires B cell interaction with subepithelial dendritic cells in Peyer’s patches

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Science  13 May 2016:
Vol. 352, Issue 6287, aaf4822
DOI: 10.1126/science.aaf4822

A recipe for intestinal lgA

Our guts are teeming with microbes, some friendly and others not. Plasma cells in the gut secrete immunoglobulin A (IgA), which helps to keep the peace with resident commensal bacteria and fights pathogens. B cell isotype switching to IgA occurs in lymphoid tissues called Peyer's patches. Reboldi et al. studied the cellular processes that guide B cells toward making IgA in mice. B cells took an unexpected journey from Peyer's patches follicles to the intestinal mucosa to interact with specialized IgA-triggering dendritic cells. The B cells then migrated back to the follicles to become IgA-producing B cells.

Science, this issue p. 10.1126/science.aaf4822

Structured Abstract

INTRODUCTION

Secretory immunoglobulin A (IgA) is made by intestinal plasma cells and has roles both in protection from gut pathogens and in maintaining homeostasis of intestinal commensals. Peyer’s patches (PPs)—the major organized lymphoid tissues of the small intestine, numbering 100 to 200 in humans and 6 to 12 in mice—are the dominant source of IgA-producing cells. A number of molecular factors have been identified that promote B cell switching from IgM to IgA, the best defined being transforming growth factor–β (TGFβ). TGFβ is made in a latent form and must be activated before it can induce TGFβ receptor (TGFβR) signaling. In this study, we explore the requirements for B cell IgA switching in PPs, concentrating on the location where it takes place and the key cell types involved.

RATIONALE

Mice deficient in the chemokine receptor CCR6 had been reported to mount poor IgA responses, but the mechanism responsible was unclear. The CCR6 ligand, CCL20, is abundant in the subepithelial dome (SED) of the PP, and one thought was that CCR6 was required for positioning dendritic cells (DCs) in the SED. However, CCR6 was known to be expressed by B cells and to be up-regulated following B cell activation. In this study, we have pursued the hypothesis that CCR6 is required within B cells to promote migration events and cellular interactions in the SED necessary for PP IgA responses.

RESULTS

Using bone marrow (BM) chimera and cell transfer approaches, we find that CCR6 expression in PP B cells is necessary for their efficient switching to IgA and for production of intestinal IgA against cholera toxin and commensal bacteria. Loss- and gain-of-function approaches establish that intrinsic CCR6 expression is necessary and sufficient for B cells to access the SED. CCR6 is up-regulated on pre–germinal center (GC) B cells in a CD40-dependent manner, and a transfer model indicates a more prominent role for CCR6 in T cell–dependent than in T cell–independent IgA responses. PP pre-GC B cells are shown to express IgA germline transcripts and activation-induced cytidine deaminase (AID), consistent with IgA switching initiating in this compartment. Using intravital two-photon microscopy, we find that B cells within the SED undergo prolonged interactions with DCs. Using BM chimera experiments and blocking reagents, we establish that SED DCs are dependent on the cytokine lymphotoxin-α1β2 (LTα1β2). RORγt+ innate lymphoid cells (ILCs) are identified as a necessary source of this cytokine. Deficiency in LTβR-dependent DCs or RORγt-dependent ILCs results in reduced IgA+ B cell frequencies in PPs. Reciprocally, transgenic overexpression of LTα1β2 increases SED DCs and IgA switching. We then examined how the SED DCs augment IgA switching and found that they abundantly expressed αvβ8, an integrin that has an established role in converting TGFβ from its latent to its active state. Experiments with Itgb8f/f Cd11c-Cre mice and with an αvβ8 blocking antibody established that DC αvβ8 expression was necessary for PP IgA switching. In vitro experiments provided further evidence that DC αvβ8 could directly activate TGFβ during DC–B cell interactions and showed that LTβR and retinoic acid signaling promote αvβ8 expression on DCs.

CONCLUSION

Our study defines a role for the PP SED as a niche that supports events necessary for IgA switching, in particular the induction of TGFβ activation, and it provides an example of a DC–B cell interaction acting to guide B cell fate. By defining a network of interactions required for IgA switching, this study identifies approaches that could be used to augment IgA responses while also defining sites for defects that could underlie IgA deficiency, the most common immune deficiency syndrome in humans.

B cell and dendritic cell distribution in mouse Peyer’s patch.

Image is a cross-sectional view of a single PP dome and the neighboring villous epithelium of the small intestine. The 7-µm frozen section was stained to detect naïve and pre-GC B cells (IgD, blue) that occupy the follicle and SED; dendritic cells (CD11c, green) that are abundant in the SED, the interfollicular T zone, and the intestinal lamina propria; T cells (CD8, red) that are present in the interfollicular T zone, the lamina propria, and in the epithelium; and nuclei (DAPI, gray). Red staining also occurred nonspecifically in association with the epithelium, and this was most prominent for the follicle-associated epithelium that overlies the SED. The follicle-associated epithelium is the site of intestinal antigen delivery into the PP. The dark (IgD-negative) oval-shaped structure within the follicle is a GC.

Abstract

Immunoglobulin A (IgA) induction primarily occurs in intestinal Peyer’s patches (PPs). However, the cellular interactions necessary for IgA class switching are poorly defined. Here we show that in mice, activated B cells use the chemokine receptor CCR6 to access the subepithelial dome (SED) of PPs. There, B cells undergo prolonged interactions with SED dendritic cells (DCs). PP IgA class switching requires innate lymphoid cells, which promote lymphotoxin-β receptor (LTβR)–dependent maintenance of DCs. PP DCs augment IgA production by integrin αvβ8-mediated activation of transforming growth factor–β (TGFβ). In mice where B cells cannot access the SED, IgA responses against oral antigen and gut commensals are impaired. These studies establish the PP SED as a niche supporting DC–B cell interactions needed for TGFβ activation and induction of mucosal IgA responses.

Immunoglobulin A (IgA), the most abundantly produced antibody isotype in the body, has the dual roles of maintaining homeostasis with the microbiome and protecting from intestinal infection (1, 2). Plasma cells located in the lamina propria secrete IgA, but the early stages of IgA production take place mainly in Peyer’s patches (PPs) (3). PPs are lymphoid organs that are organized into B cell–rich follicles, T cell–rich interfollicular zones, and a subepithelial dome (SED) rich in CD11c+ dendritic cells (DCs) that separates the epithelium from the follicles (4) (Fig. 1A). Gut-derived antigens delivered across specialized epithelial cells continually stimulate PPs, and PP follicles harbor chronic T cell–dependent germinal centers (GCs) (1). PP GCs contain a high frequency of IgA+ cells, and these give rise to IgA plasma cells. Although a number of factors have been implicated in PP B cell switching to IgA, the strongest requirement established in vivo is for transforming growth factor–β receptor (TGFβR) signaling (57). However, the cellular interactions involved in promoting TGFβR signaling in PP B cells have been unclear.

B cell–intrinsic CCR6 requirement for IgA switching in PPs

Previous studies have shown that CC-chemokine receptor–6 (CCR6)–deficient mice have altered PP organization and reduced antigen-specific IgA levels (8, 9). The CCR6 ligand, CCL20, is made abundantly by PP follicle-associated epithelium, and DC distribution in the SED was affected by CCR6 deficiency (8, 9), though this was not seen in another study (10), leaving the mechanism by which CCR6 augments IgA production unclear. An analysis of B cell distribution in wild-type PPs showed that in addition to their dense presence in follicles, IgD+ B cells were detectable more sparsely within the SED, overlapping with the network of CD11c+ Zbtb46+ DCs in this region (Fig. 1A) (11). Although CCR6 is widely expressed by B cells (12), the dynamics of PP B cell CCR6 expression have not been determined. A fraction of PP IgD+ and IgD B cells had high CCR6 surface staining (Fig. 1B), and further phenotypic analysis based on Fas (CD95), CD11c, and IgM expression showed that these B cells were enriched in pre-GC and memory B cells, respectively (fig. S1A). To confirm that PP IgD+CCR6+Fas+CD11c+ cells correspond to pre-GC cells (13, 14), we transferred wild-type follicular B cells to monoclonal MD4 Ig-transgenic mice that have little endogenous PP GC activity. A large fraction of the transferred polyclonal B cells, likely stimulated by intestinal antigen in PPs, acquired an IgDCCR6CD38GL7+ GC phenotype after 1 week (Fig. 1C). Tracking cell differentiation and division at 3 and 4 days after transfer established that CCR6 was up-regulated before the appearance of IgD GC B cells (Fig. 1D). Fas and CD11c were up-regulated with a similar time course (fig. S1B). Some cells that had undergone four or more divisions were CCR6hiIgDlo/– (Fig. 1D and fig. S1B), indicating that the CCR6+IgD gate (Fig. 1B and fig. S1B) may contain some pre-GC cells as well as memory B cells.

Fig. 1 B cell access to the PP subepithelial dome (SED) is CCR6-dependent.

(A) Representative images of Peyer’s patch (PP) dome stained with anti-CD11c (blue) and anti-IgD (brown) (left) or with anti-GFP (green) and anti-IgD (blue) (right). Dashed white line demarcates the follicle-SED boundary. Scale bars, 20 μm. (B) Representative flow cytometric analysis of CD19+ B cells in PPs for IgD and CCR6 expression. (C and D) Representative staining for fluorescence-activated cell sorting (FACS) of transferred CellTrace Violet–labeled polyclonal B cells (red) in MD4 hosts (endogenous B cells, black) for IgD and GL7 at day 7 (C) and IgD and CCR6 at days 3 and 4 after transfer. (E) Migration of PP follicular (left) and pre-GC B cells (right) from Ccr6+/+ and Ccr6–/– mice to the indicated chemokines. (F) Representative CCR6 expression on transferred wild-type (WT) and Cd40–/– B cells in MD4 hosts (upper panels) or wild-type B cells in MD4 hosts treated with either isotype control or anti-CD40L (lower panels) after 7 days. (G) Representative images of distribution of transferred B cells (Thy1.1, brown) in sections of PP from mice receiving control vector or CCR6-transduced B cells. Slides were counterstained with hematoxylin. (H) Representative images of distribution of B cells in PPs of chimeras reconstituted with 50% Ighb Ccr6+/+ or Ccr6–/– and 50% Igha wild-type BM. Sections were stained to detect Ccr6+/+ or Ccr6–/– B cells (IgDb, blue) and control B cells (IgDa, brown). (I) Distribution of polyclonal Ccr6+/+ and Ccr6–/– B cells in PPs of mixed transfer MD4 recipients [75% CD45.2 Ccr6+/+ or Ccr6–/– and 25% carboxyfluorescein succinimidyl ester (CFSE) CD45.1 wild-type] 4 days after transfer. Sections were stained to detect Ccr6+/+ or Ccr6–/– B cells (CD45.2, blue) and control B cells (CFSE, brown). Each symbol in (E) represents an individual mouse, and data are pooled from at least three independent experiments. Data in (A), (B), (C), (D), (F), (G), (H), and (I) are representative of at least three independent experiments. ***P < 0.005 (unpaired Student’s t test).

In accord with this CCR6 expression pattern, pre-GC and memory B cells, but not follicular or GC B cells, efficiently migrated toward CCL20 in a CCR6-dependent manner (Fig. 1E and fig. S1C). By contrast, PP DCs showed little migration to CCL20 while responding well to CCL21 and CXCL12 (fig. S1D). CCR6 levels and function were up-regulated in follicular B cells shortly after B cell receptor (BCR) engagement in vitro with anti-IgM (fig. S1E), though not after incubation with anti-CD40, consistent with in vitro findings for CCR6 function in activated human B cells (15). However, tracking polyclonal B cell activation in PPs with the adoptive transfer system revealed that B cells required CD40 and CD40L for CCR6 up-regulation in vivo (Fig. 1F and fig. S1F). Together, these data provide evidence that CCR6 induction in naïve B cells responding to endogenous PP-associated antigens involves CD40-dependent interactions with T helper (TH) cells. Pre-GC cells also had slightly higher amounts of CXCR4, CXCR5, and CCR7 than naïve B cells, though their response to the corresponding chemokines was not increased compared to naïve B cells (Fig. 1E and fig. S1G).

To determine whether CCR6 up-regulation could be sufficient to control B cell localization to the SED within PPs, we transferred B cells from bone marrow (BM) chimeras transduced with CCR6-encoding retrovirus to wild-type recipients. Three days later, the CCR6-overexpressing B cells, identified by expression of a Thy1.1 reporter, were situated preferentially in the SED (Fig. 1G and fig. S2A). By contrast, B cells transduced with the control retrovirus were distributed uniformly within the follicle and SED (Fig. 1G and fig. S2A). To test whether CCR6 was necessary for B cell localization in the SED, we examined B cell distribution in 50:50 mixed BM chimeras that contained CCR6-deficient or littermate control (Ighb) cells mixed with wild-type (Igha) cells. Notably, CCR6-deficient and wild-type B cells were equally represented in the follicle, but CCR6-deficient B cells were unable to migrate into the SED (Fig. 1H and fig. S2B). Using the procedure of adoptive cell transfer into MD4 hosts, we found that B cells accessed the SED in a CCR6-dependent manner within 4 days of activation by endogenous antigen (Fig. 1I and fig. S2C).

Because CCR6 up-regulation on follicular B cells is associated with the transitional stage between naive and GC B cell phenotypes, we sought to directly test the importance of CCR6 in PP B cell fate. We used mixed wild-type (Igha): CCR6-deficient (Ighb) BM chimeras to determine the intrinsic role of CCR6 in B cells and ensure that other CCR6-dependent properties of PPs were intact (810, 16). CCR6-deficient GC B cells in these chimeras suffered reduced switching to IgA compared to wild-type GC B cells in the same animals and showed instead an increased propensity for switching to IgG1 (Fig. 2A). In accord with most PP IgA+ cells being GC B cells (Fig. 2B), the frequency of IgA+ cells was decreased among total Ccr6–/– B cells (Fig. 2C and fig. S3A). Analysis of mesenteric LNs (MLNs) in the same animals showed only a low frequency of IgA+ cells and no impact of CCR6 deficiency (fig. S3B). CCR6 deficiency did not significantly affect the fraction of PP B cells with pre-GC, GC, or memory phenotypes (fig. S3C). Analysis of wild-type and CCR6-deficient cells cotransferred to MD4 hosts showed that the early appearance of IgA+ GC cells was CCR6-dependent (Fig. 2D) and, again, CCR6 deficiency did not affect the induction of pre-GC, GC, or memory cells (fig. S3D). IgA class switching in vitro was not affected by CCL20 (fig. S3G), consistent with the CCR6 requirement being to support B cell positioning within the PP. Interestingly, the mixed chimeras also had reduced frequencies of Ccr6–/– TH17 cells in PPs (fig. S3F). However, because the defective IgA response was specific to the allotype marked CCR6-deficient B cells, actions of the receptor in other cell types cannot account for the CCR6 requirement in B cells. The inability to undergo productive IgA class switching in PPs had a notable impact on mucosal IgA. In mixed chimeras, free IgA derived from CCR6-deficient B cells was underrepresented in fecal pellets (Fig. 2E), and CCR6-deficient B cells made a diminished contribution to the IgA coating intestinal bacteria (Fig. 2F). The role of CCR6 in controlling B cell class switching to IgA was not restricted to the homeostatic situation because after oral immunization of mixed BM chimeras with cholera toxin (CT), a potent lethal toxin that causes severe diarrhea, the IgA response was dominated by antibody derived from the wild-type B cells (Fig. 2G).

Fig. 2 B cell isotype switching to IgA is CCR6-dependent.

(A) Frequency of WT and Ccr6+/+ or Ccr6–/– GC B cells expressing IgA or IgG1 in PPs from mixed BM chimeras, as determined by FACS. (B) GC and memory contribution to IgA+ B cell pool in PPs. (C) Representative FACS of frequency of WT and Ccr6+/+ or Ccr6–/– B cells expressing IgA or IgG1 in PPs from mixed BM chimeras. (D) Representative FACS staining (left) and frequency (right) of IgA+ B cells among transferred B cells in MD4 recipient PPs after 7 days. (E) Fecal IgA from WT (Igha) and Ccr6+/+ or Ccr6–/– (Ighb) mixed BM chimeras as measured by allotypic ELISA. (F) Percentage of fecal bacteria coated with IgAa or IgAb measured by FACS from mixed BM chimeras as in (A). (G) Fecal CT-specific IgA from WT (Igha) and Ccr6+/+ or Ccr6–/– (Ighb) mixed BM chimeras orally treated with CT, measured by allotypic ELISA. (H) Fecal IgA from WT (Igha) and Ccr6+/+ or Ccr6–/– (Ighb) B cells cotransferred into either Rag1–/– or μMT, or from mixed BM chimeras as in (A), measured by allotypic ELISA. Each symbol in (A), (B), (E), (F), (G), and (H) represents an individual mouse, and data are pooled from at least three independent experiments. Data in (C) and (D) are representative of at least three independent experiments. **P < 0.01, ***P <0.005 (unpaired Student’s t test).

The IgA response against intestinal commensals is thought to involve both T-independent and T-dependent antibody production (17, 18). Because the great majority of IgA+ B cells in wild-type PPs are GC phenotype cells, we anticipated that the role of CCR6 in promoting IgA may be most prominent during T-dependent responses. To test this hypothesis, we transferred mixtures of wild-type Igha and Ccr6+/+ or Ccr6–/– Ighb B cells to mice lacking endogenous B cells and that were either T cell–deficient (Rag1–/–) or T cell–replete (μMT). Allotype-specific analysis of fecal IgA 1 month later revealed that CCR6 was not required for B cells to mount a T-independent IgA response in the Rag1–/– hosts (Fig. 2H), whereas the response in the T cell–replete μMT hosts showed a CCR6 dependence similar to the responses in mixed BM chimeras (Fig. 2H). In accord with the CCR6-dependent secretory IgA responses occurring in PPs, when mixed BM chimeras were generated with lymphotoxin-β-receptor (LTβR)–deficient hosts that are unable to form PPs (19), B cell CCR6 expression did not influence total or commensal-bound fecal IgA (fig. S3G).

IgA class switching is initiated in the subepithelial dome

To determine whether IgA class switch recombination (CSR) was initiating at the pre-GC stage, we examined IgA germline transcript (αGT) expression by semiquantitative and quantitative polymerase chain reaction (PCR) in naïve (IgD+CCR6), pre-GC (IgD+CCR6+), GC (IgDCCR6), and memory (IgDCCR6+) B cells from wild-type PPs. Pre-GC cells showed a significant increase in αGTs compared to naïve and GC B cells (Fig. 3, A and B). IgA GTs were also abundant in IgDCCR6+ B cells, perhaps reflecting the presence of both late-stage pre-GC cells and memory-cell–derived pre-GC cells in this gate. Although IgA is the major memory B cell isotype, a fraction of the cells in this gate are unswitched (figs. S1A and S3H). We also detected mature, rearranged, Iμ-Cα transcripts and switch circle transcripts (Iα-Cμ) in the pre-GC cells, though in this case, the levels were higher in IgDCCR6 GC B cells, as expected from the high fraction of IgA+ B cells in the GC (Fig. 3A). Consistent with switching initiating in the pre-GC compartment, activation-induced cytidine deaminase (AID) transcripts were elevated in pre-GC compared to naïve B cells (Fig. 3B), and the frequency of AID-GFP+ cells was higher (Fig. 3C). Although AID expression was lower in pre-GC than in GC cells (Fig. 3B), the amounts of AID required for CSR are less than those required for somatic hypermutation of V regions (20).

Fig. 3 Pre-GC B cells in PPs initiate IgA class switching.

(A and B) Representative semiquantitative (A) or quantitative (B) RT-PCR on B cells from PPs sorted according to IgD and CCR6 expression, for the indicated transcripts. (C) Representative FACS staining (left) and frequency (right) of AID-GFP+ PP B cells from reporter mice, according to IgD and CCR6 expression. Each symbol in (B) and (C) represents an individual mouse, and data are pooled from three independent experiments. Data in (A) are representative of three independent experiments. A.U., arbitrary units.

Lymphotoxin-dependent PP dendritic cells are required for IgA switching

On the basis of the above kinetic and anatomical findings, we reasoned that PP B cells might travel to the SED to receive a stimulus that dictated IgA class switching. The SED is a CD11c+ Zbtb46+ DC–rich area (Fig. 1A), containing mainly CD11b+ and CD11b-CD8-double negative (DN) DCs, with CD8+ DCs localized in the interfollicular region (IFR) (4, 9, 21, 22). CD11c+ DCs were minimally detected in PP GCs (fig. S4A). In vitro studies with human and mouse cells have shown that B cell–DC coculturing can augment IgA switching, though a role for such interactions in vivo has not been established (2329). To test whether SED DCs were important for B cell IgA switching, we sought to identify perturbations affecting these DCs. LTβR signaling contributes to CD11b+ DC homeostasis in the spleen (30), but whether it has a role in maintaining DCs in PPs is unknown. Analysis of Ltbr transcripts in sorted DC subsets from PPs showed that CD11b+ and DN DCs had high expression (Fig. 4A), and these cells were positive for surface LTβR by flow cytometry (Fig. 4A and fig. S4B). When wild-type mice were reconstituted with Ltbr–/– BM, they had a deficiency in CD11b+ and DN DCs, whereas CD8+ DCs were less affected (Fig. 4B). Notably, in these same chimeras, B cell switching to IgA was reduced, and switching to IgG1 was increased (Fig. 4C). A similar defect in the balance between IgA and IgG1 class switching was observed in Ltbr–/: Itgax-diphtheria toxin receptor (CD11c-DTR) mixed BM chimeras that had been treated with diphtheria toxin (DT) such that most DCs remaining in the animals were LTβR-deficient, whereas all hematopoietic CD11c cell types were 50% wild-type (Fig. 4D). In a reciprocal experiment, we tested whether increased LTα1β2-LTβR signaling was sufficient to promote SED DC accumulation and IgA class switching by examining PPs from transgenic mice overproducing LTα1β2 (30). In these animals, CD11b+ DCs were increased in number, and a greater fraction of GC B cells had switched to IgA compared to littermate controls (Fig. 4, E and F). The transcription factor BATF3 controls the development of CD8a+ DCs (31), and mice reconstituted with Batf3–/– BM showed a near-absence of CD8a+ DCs in PPs (fig. S4C). In these mice, B cell switching to IgA was normal (fig. S4D), indicating that CD8a+ DCs are dispensable for PP IgA class switching.

Fig. 4 LTβR-dependent PP DC support IgA class switching.

(A) Quantitative PCR analysis of Ltbr transcript abundance in the indicated DC subsets from PPs (left) and median fluorescence intensity (MFI) ratio for LTβR on the DC subsets from PPs of BM chimeras reconstituted with WT or Ltbr–/– BM (right). (B) Representative FACS staining (left) and absolute cell number (right) of the indicated DC subset from PPs of BM chimeras reconstituted with WT or Ltbr–/– BM. (C) Frequency of IgA+ and IgG1+ GC B cells in PPs from BM chimeras as in (B), determined by FACS. (D) Frequency of IgA+ and IgG1+ GC B cells in PP from mixed chimeras reconstituted with CD11c-DTR and either Ltbr+/+ or Ltbr–/– BM, treated with DT for 3 weeks. (E) Absolute number of the indicated DC subset from PPs of WT or LT-transgenic (tg) mice. (F) Frequency of IgA+ and IgG1+ GC B cells in PPs from WT or LT-tg mice, as determined by FACS. (G) Absolute number of the indicated DC subset from PPs of mice treated with LTβR-Fc or hIgG-Fc for 7 days. (H) Representative FACS staining of polyclonal B cells in PPs 7 days after transfer to MD4 recipients that were treated with LTβR-Fc or hIgG-Fc (left) and frequency of IgA+ B cells and GC B cells in PPs among transferred B cells (right). Each symbol in (A), (B), (C), (D), (E), (F), (G), and (H) represents an individual mouse, and data are pooled from at least three independent experiments. In (B), (C), (D), (E), (F), (G), and (H), *P < 0.05, **P < 0.01, ***P < 0.005 (unpaired Student’s t test); in (A), ***P < 0.005 (one-way ANOVA with Bonferroni’s post-hoc test).

A concern with the above studies was that chronic LTβR deficiency in DCs might lead to distant alterations such as changes in the microbiome that have indirect effects on IgA class switching. To address this concern, we used the adoptive transfer approach introduced above (Fig. 1C). Treatment with LTβR-Fc, an LTα1β2 antagonist, during the short period of the transfer decreased the number of CD11b+ DCs (Fig. 4G) and reduced the ability of the transferred B cells to undergo IgA class switching (Fig. 4H), without affecting their participation in the GC reaction (Fig. 4H). Taken together, these findings provide strong evidence that LTβR signaling in DCs is directly required for promoting B cell IgA class switching.

Innate lymphoid cells maintain PP dendritic cells required for IgA switching

Innate lymphoid cells type 3 (ILC3, also known as lymphoid tissue inducer cells) are an important source of LTα1β2 for LN and PP organogenesis and during mucosal immune responses (1, 29, 32, 33). ILC3s in PPs expressed high levels of surface LTα1β2 (Fig. 5A). Although a previous study showed that ILC3-derived LT augmented lamina propria IgA responses, the mice analyzed in that study were PP-deficient, preventing any assessment of the ILC3 role in PP IgA responses (29). To test the importance of ILC3s in controlling IgA class switching in PPs, we reconstituted irradiated mice using BM cells deficient for RORγt, a transcription factor essential for ILC3 development (34). B cells in chimeras lacking ILC3s showed an impaired ability to undergo IgA class switching and an increased propensity to switch to IgG1 compared to B cells from wild-type chimeras (Fig. 5B). These findings suggested that RORγt+ cells were critical in promoting IgA class switching. However, RORγt (encoded by Rorc) is expressed not only in ILC3s, but also in various T cell types (35). To further define the ILC contribution to B cell class switching, we reconstituted irradiated mice with wild-type BM, Rorc–/– BM, or a mixture of Rorc–/– and Rag1–/– BM. In the latter mice, the BM mixture could give rise to ILC3s (from the Rag1–/– BM) but none of the RORγt-dependent T cell populations. Notable, whereas animals lacking RORγt in all hematopoietic cell types showed decreased IgA and increased IgG1 class switching, the animals reconstituted with a mix of Rorc–/– and Rag1–/– BM showed IgA and IgG1 class switching similar to that of animals reconstituted with wild-type BM (Fig. 5C). Consistent with ILC3s promoting IgA switching via effects on DCs, the number of CD11b+ and DN DCs was reduced in the chimeras lacking ILC3s (Rorc–/– chimeras) but not in those selectively lacking RORγt-dependent T cells (Rorc–/–: Rag1–/– chimeras, fig. S5A). These results provide strong evidence that ILC3s are the only RORγt-dependent population critical in controlling B cell class switching to IgA in PPs.

Fig. 5 LT- and ILC3-dependence of PP IgA response.

(A) Representative FACS staining showing gating strategy and LTα1β2 on ILC3s in PP of WT mouse using LTβR-Fc. (B) Frequency of IgA+ and IgG1+ GC B cells in PPs from chimeras reconstituted with WT or Rorc–/– BM. (C) Frequency of IgA+ and IgG1+ GC B cells in PPs as determined by FACS in BM chimeras reconstituted with WT, Rorc–/–, or a mix of Rorc–/– and Rag1–/– BM. (D) Frequency of IgA+ and IgG1+ GC B cells in PPs as determined by FACS in BM chimeras reconstituted with a mix of Lta–/– BM and either WT or Rorc–/– BM. (E) Representative immunofluorescence of PP SED from Rorc(γt)-EGFP mouse stained for the indicated markers. Yellow arrowheads indicate ILC3s proximal to DCs. Scale bar, 20 μm. Each symbol in (B), (C), and (D) represents an individual mouse, and data are pooled from at least three independent experiments. Data in (A) and (E) are representative of at least three independent experiments. In (B) and (D),***P < 0.005 (unpaired Student’s t test); in (C), ***P < 0.005 (one-way ANOVA with Bonferroni’s post-hoc test). ns, not significant.

To evaluate the effect of LT on ILC3s in controlling B cell class switching, we made Rorc–/–: Lta–/– mixed BM chimeras. In such animals, all the RORγt+ cells are LTα-deficient, whereas 50% of the RORγt cells are wild-type for LTα. When ILC3s lacked LTα, GC B cells class switched preferentially to IgG1 over IgA (Fig. 5D). These mice also showed a deficiency in CD11b+ and DN DCs, in accord with the dependence of these DCs on LTβR (fig. S5A). The ILC3-deficient mice also had a reduction in CD8+ DCs, suggesting additional influences of these cells on DC maturation. Although B cells are an established source of LTα1β2 within follicles (19), they express considerably less of this cytokine than ILC3s (fig. S5B), and in chimeric animals selectively lacking LTα1β2 from B cells, PP GC IgA+ cell frequencies and CD11b+ DC frequencies were normal (fig. S5C). Finally, using RORγt-eGFP reporter mice to track RORγt+ cell distribution in PPs, we found RORγt+CD3 ILC3s in the SED making contact with CD11c+ DCs (Fig. 5E and fig. S5D). These results indicate that in PPs, B cell class switching is controlled by the LT-LTβR axis, likely through direct interaction between ILC3s expressing LTα1β2 and SED DCs expressing LTβR.

B cells undergo prolonged interactions with PP subepithelial dome DCs

To determine whether B cells in the SED were interacting with DCs, we performed intravital imaging using two-photon laser-scanning microscopy (TPLSM). CD11c-YFP (yellow fluorescent protein) mice were injected with CFP+ B cells, and 10 to 14 days later, individual PPs were surgically exposed and stabilized for imaging by attachment to a platform placed over the mouse abdomen (36). Subepithelial dome B cells were identified as being situated in the YFP+ cell-rich area just beneath the epithelial layer. Contours were drawn immediately internal to the YFP+ cells in each z-plane and used to generate a three-dimensional surface to separate the SED and follicle (Fig. 6A and movie S1). B cells within the SED or follicle moved with similar velocities (Fig. 6B), but B cells within the SED showed smaller displacement, indicating a greater amount of confinement (Fig. 6C). On average, two-thirds of B cells in the SED engaged in short (Scan) or long (Pause) interactions with DCs during 30-min imaging sessions (Fig. 6, D and E, and movies S2 and S3). To examine B cell migration between follicle and SED, we manually annotated the tracks of cells that crossed between zones. As well as observing B cells migrating from the follicle into the SED, we observed B cells moving in the reverse direction, from the SED into the follicle, in some cases following long, seemingly directed tracks (Fig. 6F and movie S4). When B cells were incubated in vitro with CCL20, CCR6 became down-regulated (fig. S5E), and we suggest that ligand-mediated receptor desensitization over time allows follicular chemoattractant cues to dominate over CCL20 and attract cells away from the SED.

Fig. 6 PP B cell migration dynamics and interaction with SED DCs.

(A) Representative TPLSM of PP for transferred CFP+ B cells (green) in CD11c-YFP mice. YFP+ cells (red) are shown by volume rendering. White dotted line indicates location of the SED. (B) Median velocity and (C) displacement versus square root of time of B cells in follicle (black) and in the SED (blue). (D) Percentage of B cells in the SED pausing (Pause), scanning (Scan), or not contacting (No contact) CD11c-YFP+ cells. (E) Representative time-lapse images of CFP+ B cell interaction with CD11c-YFP+ cell in the SED. Arrowheads highlight a single B cell–DC interaction. (Yellow color is due to bleed-through between channels.) Time is shown in minutes:seconds. (F) Representative z-projection view of PP follicle and SED of the type in (A), showing only the CFP+ B cells. White dotted line, SED–follicle border; yellow lines, tracks of B cells in the follicle; blue lines, tracks of B cells in the SED; pink lines, tracks of B cells moving from the SED to the follicle; white lines, tracks of B cells moving from the follicle to the SED. Each symbol in (B) and (D) represents an individual mouse, and data are pooled from at least three independent experiments. Data in (A), (C), (E), and (F) are representative of at least three independent experiments.

DC integrin αvβ8 is required for TGFβ activation and induction of IgA switching

Because our findings indicated that interaction between B cells and DCs in the SED was required to achieve a successful IgA class switch, we investigated the factors involved in this interaction. Several molecules have been described as promoting IgA class switching in vivo (26, 28, 37); however, the most profound phenotype has been reported in mice where TGF-βRII was specifically deleted in B cells (6). Activated B cells abundantly express TGFβ transcripts (38), and B cell deficiency in this cytokine leads to a reduction in fecal IgA (7). TGFβ activation can be mediated by αvβ6 or αvβ8 integrins binding to latency-associated peptide (LAP) and exerting forces that liberate the active cytokine (39). CD11b+ and DN DCs in PPs showed abundant transcripts for the Itgb8 (β8) and Itgav (αv) integrin chains (Fig. 7A), and a subset of these DCs had surface expression of the integrin (Fig. 7B). We therefore tested whether DC expression of integrin β8 was required for IgA switching. Irradiated hosts reconstituted with BM from Itgb8flox/flox Cd11c-Cre mice showed a defect in IgA class switching in PP GCs and a propensity toward increased IgG1 class switching (Fig. 7C). By contrast, β8 deficiency did not affect the frequency of IgA+ or IgG1+ B cells in MLNs (fig. S5F). Short-term treatment with anti-β8 in vivo reduced the ability of transferred naïve B cells to undergo IgA class switching in PPs (Fig. 7D), making it unlikely that the switching defect was due to indirect effects of DC β8 deficiency. Moreover, when DCs deficient for β8 were sorted from PPs and cocultured in vitro with stimulated B cells, they failed to support IgA class switching, whereas control DCs supported robust IgA switching (Fig. 7E). Blocking TGFβ signaling in these B cell–DC cocultures with anti-TGFβ, anti-LAP, or anti-β8 reduced the ability of B cells to undergo IgA class switching (Fig. 7F). DC subset analysis revealed that CD11b+ DCs could induce IgA class switching in vitro, in accord with their integrin β8 and LTβR expression (Fig. 7G). Finally, using BM-derived DCs, we found that LTβR engagement with an agonistic antibody led to a weak but reproducible induction of β8 integrin expression (Fig. 7H). Retinoic acid (RA) has an established role in augmenting IgA production, possibly through actions on DCs (40). RA treatment of the DC cultures led to a slightly greater β8 integrin induction than LTβR agonism, and when the two stimuli were combined, they acted in an additive manner (Fig. 7H).

Fig. 7 DC integrin-β8 promotes TGFβ-dependent IgA switching.

(A) Quantitative PCR analysis of Itgb8 (left) and Itgav (right) transcript abundance in the indicated DC subsets from PPs. (B) Median fluorescence intensity (MFI) ratio for β8 on the indicated DC subsets from PPs of either CD11c-Cre (WT) or CD11c-Cre+ Itgb8fl/fl mice. (C) Frequency of IgA+ and IgG1+ GC B cells in PPs of either CD11c-Cre or CD11c-Cre+ Itgb8fl/fl mice as determined by FACS. (D) Frequency of IgA+ and IgG1+ GC B cells in PPs 7 days after adoptive transfer in MD4 mice and treatment with either isotype control or anti-β8. (E) Frequency of IgA+ splenic B cells upon 5 days of culture with DCs sorted from PPs or mLNs of CD11c-Cre or CD11c-Cre+ Itgb8fl/fl mice. (F) Frequency of IgA+ splenic B cells upon 5 days of culture with DCs sorted from PPs and incubated with the indicated antibodies. (G) Frequency of IgA+ splenic B cells upon 5 days of culture with the indicated DC subset sorted from PPs. (H) Median fluorescence intensity (MFI) for β8 on the BMDCs generated from CD11c-Cre or CD11c-Cre+ Itgb8fl/fl BM upon 7 days of culture and incubation with the indicated reagents. Each symbol in (B), (C), (D), (E), (F), (G), and (H) represents an individual mouse, and data are pooled from at least three independent experiments. Data in (A) are pooled from three independent experiments. In (B), (F), and (G), *P < 0.05, **P < 0.01, ***P < 0.005 (one-way ANOVA with Bonferroni’s post-hoc test); in (C), (D), and (E), **P < 0.01, ***P < 0.005 (unpaired Student’s t test).

The increased switching in vivo to IgG1 under conditions of reduced IgA switching may be a consequence of the ready availability of IL4 in PPs because it is highly expressed by PP Tfh cells (fig. S5G). Consistent with this interpretation, in irradiated hosts reconstituted with BM from IL4R-deficient mice, there was an almost complete absence of IgG1+ cells in PPs (fig. S5H). Under normal conditions, TGFβ-mediated IgA switching may dominate, eliminating the intervening Ig constant regions and thereby limiting switching to IgG1.

Discussion

These studies identify a network of cellular and molecular interactions underpinning the induction of IgA responses in PPs (fig. S6). After activation by foreign or commensal-derived antigen and receipt of CD40-dependent helper signals, PP B cells up-regulate CCR6 and are attracted by CCL20 into the SED, where they undergo extensive interactions with CD11b+ DCs. This DC population is maintained by LTα1β2 provided locally by ILC3s. CD11b+ DCs express integrin αvβ8 and promote TGFβ activation during interactions with B cells. After receipt of TGFβ and likely additional SED-derived signals, B cells return to the follicle by directed migration and participate in the GC response.

Our studies indicate that sustained CCR6 up-regulation in PP B cells occurs in a CD40-, and thus most likely T cell–dependent, manner, and CCR6 deficiency strongly affected T-dependent IgA responses. How CD40 signaling promotes sustained CCR6 expression is not yet clear. Because BCR engagement is sufficient to promote CCR6 up-regulation in vitro, it remains possible that CCR6 augments certain T-independent IgA responses, with expression perhaps being sustained by other inputs such as from Toll-like receptor ligands. It is notable that memory B cells in PPs have high amounts of CCR6, and we speculate that they have privileged access to the SED, perhaps facilitating more rapid exposure to newly arriving antigens.

A key source of TGFβ1 for intestinal IgA production is the B cells themselves (7). However, B cell TGFβ1 deficiency does not cause a complete block in IgA production. Given the widespread expression of TGFβ family members (Immgen.org), we consider it likely that more than one cell type contributes latent TGFβ for DC-mediated activation and triggering of IgA switching. A number of other signals have been implicated in promoting IgA production, including RA, inducible nitric oxide synthase (iNOS), APRIL, and interleukin-6 (IL-6) (24, 28, 37, 40), and our studies do not exclude a role for these mediators in directly or indirectly supporting IgA class switching in PPs. In particular, we suggest that RA helps establish an environment where αvβ8+ DCs can develop or be maintained. Although β8-integrin–deficient mice were not reported to have reduced serum IgA (41), these mice suffer from inflammatory disease due to regulatory T cell (Treg) deficiency, and this likely allows other factors to induce IgA switching or to generate active TGFβ. Consistent with our findings, a recent study of lung DCs noted a correlation between DC Itgb8 transcript expression and induction of TGFβ-dependent IgA switching (42). We speculate that during B cell–DC interactions in PP SEDs, synaptic contacts form where DC αvβ8 exerts force on TGFβ-LAP tethered on the B cell, leading to TGFβ activation (39) and engagement of B cell TGFβR to induce isotype switching. By defining a network of interactions required for IgA switching, this study identifies approaches that could be used to augment IgA responses while also defining sites for defects that could underlie IgA deficiency, the most common immune deficiency syndrome in humans (43).

Methods

Mice

Wild-type and Ly5.2 (CD45.1) congenic C57BL/6 (B6) mice, 6 to 12 weeks old, were from the National Cancer Institute. Lta−/−, Ltbr−/−, Igha, MD4-Ig tg, and Lt-tg [line Ltb10 (44)] mice were from an internal colony. Itgax (Cd11c)-cre Itgb8fl/fl mice (41) were backcrossed to C57BL/6J for 10 generations. Itgax-DTR, Ccr6−/−, Rorc−/−, Rag1−/−, Batf3−/−, Cd40−/−, and AID-GFP mice were from Jackson laboratories. IL4-hCD2 (KN2) and Il4ra−/− mice were kindly provided by the Locksley lab. Animals were housed in a specific pathogen-free environment in the Laboratory Animal Research Center at the University of California San Francisco (UCSF), and all experiments conformed to the ethical principles and guidelines approved by the UCSF Institutional and Animal Care and Use Committee.

Flow cytometry

Spleen, PPs, and mLN cell suspensions were generated by mashing the organs through 70-μm cell strainers. For DC isolation, PPs and mLN were digested with 1.6 mg of type II collagenase (Worthington Biochemical) and deoxyribonuclease I for 10 min at 37°C. Digested PPs were mashed into a single cell suspension through a 70-μm cell strainer in phosphate-buffered saline (PBS) buffers containing 2% fetal calf serum and 2 mM EDTA. Cells were stained with antibodies to CD4 (GK1.5), B220 (RA3-6B2), CD19 (1D3), IgD (11-26c.2a), CD95 (Jo2), GL7, CD38 (90), CCR6 (140706), CD8 (53), MHCII (AF6-120.1), IgA (1040-09), IgG1 (RMA1-1), CD11c (N418), CD11b (M1/70), CD45.1 (A20), CD45.1 (104) (from Biolegend, BD Biosciences, rnBiotech, or eBioscience). Biotin conjugates were detected with streptavidin Qdot605 (Invitrogen). To detect intracellular IgA, cells were stained with fixable viability dye (eFluor780; eBioscience) to exclude dead cells then stained for surface antigens, treated with BD Cytofix Buffer and Perm/Wash reagent (BD Biosciences), and stained with anti-IgA.

Immunohistochemistry and immunofluorescence microscopy

For immunohistochemistry, cryosections of 7 μm were acetone fixed and stained as described (45) with combinations of the following antibodies: anti-IgD (11-26c.2a, BD Biosciences), anti-IgDa (AMS9.1, BD Biosciences), anti-IgDb (217-170, BD Biosciences), and anti-IgMa (DS-1, BD Biosciences). In some case, the slides were counterstained with hematoxylin. For immunofluorescence, tissues were fixed in 4% paraformaldehyde in PBS for 2 hours at 4°C, washed three times for 10 min in PBS, then moved to 30% sucrose in PBS overnight. Tissues were flash frozen in Tissue-Tek Cryomold (VWR) the next day, and 7-μm sections were cut and then dried for 1 hour before staining. Sections were rehydrated in PBS with 1% bovine serum albumin (BSA) for 10 min and then stained in primary antibody overnight at 4°C and stained for subsequent steps for 2 hours at room temperature, all in PBS with 1% BSA, 2% mouse serum, and 2% rat serum. Sections were stained with primary antibodies: Rabbit anti-GFP (polyclonal, Life Technologies), goat anti-mouse IgD (goat polyclonal GAM/IGD(FC)/7S, Cedarlane Labs), Alexa647-conjugated anti-CD11c (N418, Biolegend), and PE-conjugated anti-CD3 (17A2, Biolegend). Sections were then stained with the following secondary antibodies: Alexa488-conjugated donkey anti-rabbit (A-21206, Life Technologies) and aminomethylcoumarin (AMCA) conjugated donkey anti-goat (705-156-147, Jackson Immunoresearch).

Cell transfer, immunization, and transwell assays

For MD4 B cell positioning analysis, (1 to 2) × 107 MD4 wild-type (WT) or MD4 Ccr6−/− B cells were transferred in C57BL/6 recipients for 3 days before immunizing with 5 mg of hen egg lysozyme (HEL) intravenously (i.v.), and PPs were harvested at different time points. For polyclonal B cell transfer, MD4 WT recipients were adoptively transferred with (1 to 2) × 107 splenic B cells from congenic C57BL/6 for the indicated time. In some cases, B cells were stained with CellTrace Violet (Life Technologies) according to the manufacturer’s protocol. For LTβR-Fc treatment, mice were treated with LTβR-Fc protein (provided by J. Browning) by i.v. injection of 100 μg of protein every 3.5 days for 7 days. For anti-Itgβ8 treatment, mice were treated with neutralizing antibody by i.v. injection of 10 mg of antibody per kilogram of body weight every 3.5 days for 7 days. For anti-CD40L treatment, mice were treated with neutralizing antibody by i.v. injection of 1 mg of anti-mouse CD40L (clone MR1) every 3.5 days for 7 days. For cholera toxin immunization, mice were injected three times orally with 10 μg of cholera toxin (CT) (EMD Bioscience), oral immunizations were performed 7 days apart, and mice were analyzed 7 days after the final immunization. Transwell migration assays were done with 5-μm transwells using 106 digested PP cells and enumeration of transmigrated cells by flow cytometry as previously described (46). Chemokines were obtained from PeproTech or R&D Systems.

Sorting

For DC sorting, PPs and mLNs were digested and stained as described above and sorted on a FACSAira III with a 70-μm nozzle. In same cases, DCs were isolated from PPs of 8- to 10-week-old mice that had been injected subcutaneously in the flank with 5 × 106 B16 murine Flt3L-secreting tumor cells 7 to 10 days earlier. This treatment led to a ~10-fold expansion in total PP DC numbers.

Bone marrow chimeras, retroviral transduction, and DT treatment

Ly5.2 congenic B6 mice were lethally irradiated with either 1100 or 1300 rad in split doses and reconstituted with (1 to 3) × 106 BM cells from the indicated donors. Mice were analyzed 10 to 14 weeks later. For retroviral transduction, PlatE cells were transfected with murine stem cell virus (MSCV) retroviral constructs encoding full-length mouse Ccr6 with Lipofectamine 2000 (Invitrogen) following the manufacturer’s protocol. For transduction of BM-derived cells, BM cells were harvested 4 days after 5-flurouracil (Sigma) injection and cultured in the presence of recombinant IL-3, IL-6, and mouse stem cell factor (SCF) (100 ng/ml, Peprotech). BM cells were spin-infected twice with a retroviral construct expressing Ccr6 and an internal ribosomal entry site (IRES)–Thy1.1 cassette as a reporter. One day after the last spin infection, the cells were injected into lethally irradiated C57BL/6 recipients. Eight to 12 weeks later, splenic B cells were isolated and then injected in C57BL/6 recipients, and their positioning in PP was assessed 3 days later. For DT treatments, BM chimeras received 4 ng of DT (EMD Bioscience) per gram of body weight every 72 hours for 3 weeks.

Rag1−/− and μMT cells transfer

Splenic B cells were sorted on a FACSAira III with a 70-μm nozzle as CD3, CD43, CD4, CD8, CD11c, Ly6C, Ly6G, CD90 neg, and 106 cells from each genotype were transferred i.v. Recipients were analyzed 4 weeks later.

Enzyme-linked immunosorbent assay (ELISA)

Ninety-six–well plates (Thermo Fisher Scientific) were coated with purified anti-IgA (RMA-1, BD) or 0.5 nmol/ml monosialotetrahexosylganglioside (GM1) (Sigma-Aldrich) followed by 0.5 μg/ml CT overnight at 4°C (47). The plates were washed and clocked with PBS–5% BSA before diluted fecal samples were added and twofold serial dilution was made. Samples were incubated overnight at 4°C, followed by biotinylated anti-mouse antibodies: anti-IgA (C10-1, BD), anti-IgAa, and anti-IgAb (Hy16 and HISM2, UCSF Hybridoma Core) at 1 μg/ml in PBS–0.1% BSA. Detection antibodies were labeled by streptavidin-conjugated horseradish peroxidase and visualized by the addition of Substrate Reagent Pack (R&D). Color development was stopped with 3 M H2SO4. Purified mouse IgA (Southern Biotech) served as standard. Absorbances at 450 nm were measured on a tunable microplate reader (VersaMax, Molecular Devices). Antibody titers were calculated by extrapolating absorbance values from standard curves where known concentrations were plotted against absorbance using SoftMax Pro 5 software.

Flow cytometric analysis of IgA-bound bacteria

Flow cytometric analysis of gut bacteria in feces was done as described (48). Briefly, fecal pellets were suspended in filtered PBS (100 μl to 10 mg of feces), homogenized well, and centrifuged at 400g for 5 min to remove larger particles from the fecal suspension. Supernatant containing bacteria was centrifuged at 8000g for 10 min. The bacterial pellet was blocked on ice in 1 ml of BSA-PBS (1% w/v) for 15 min. Samples were spun at 8000g for 10 min. Bacteria were stained with anti-IgAa and anti-IgAb on ice for 20 min and washed with PBS. Finally, bacterial pellets were resuspended in SYBR green I [1/10000 (v/v) dilution Life Technologies] and analyzed with an LSRII flow cytometer.

RNA isolation and real-time reverse transcription–polymerase chain reaction (RT-PCR)

Total RNA was isolated from sorted DCs and B cells from PPs with the TRIzol reagent (Life Technologies) following the manufacturer’s protocol. Real-time PCR was performed with SYBR Green PCR Mix (Roche) and an ABI prism 7300 sequence detection system (Applied Biosystems, Foster City, CA).

Hprt forward: AGGTTGCAAGCTTGCTGGT; reverse: TGAAGTACTCATTATAGTCAAGGGCA

Ltbr forward: CCAGATGTGAGATCCAGGGC; reverse: GACCAGCGACAGCAGGATG

Itgb8 forward: CTGAAGAAATACCCCGTGGA; reverse: ATGGGGAGGCATACAGTCT

Itgav forward: CGCCTATCTTCGGGATGAATC; reverse: CCAACCGATACTCCATGAAAA

aGT forward: CCAGGCTAGACAGAGGCAAG; reverse: CGGAAGGGAAGTAATCGTGA

Aicda forward: GCCAAGGGACGGCATGAG; reverse: GATGTAGCGTAGGAACAACAA

Semiquantitative RT-PCR on sorted B cells for AID, alpha germline transcripts (αGT), Iμ-Cα, and Iα-Cμ were amplified with primers and conditions described before (26).

In vitro culture

For CCR6 up-regulation, splenic B cells were stimulated with 10 μg/ml anti-IgM [F(ab′)2 goat anti-mouse IgM, Jackson Immunoresearch] for the indicated time. For IgA class switch B-DC coculture experiments, magnetic cell sorter (MACS)–isolated splenic B cells (typically at 50,000 cells per well) were stimulated with 10 μg/ml anti-IgM and 20 μg/ml anti-CD40 (clone FGK4.5, UCSF Hybridoma Core) in the presence or absence of sorted DCs at a ratio of 1:1 for 5 days. The sorted DCs were from PPs of untreated mice in all cases except for the experiment involving sorted DC subsets, where they were from B16-Flt3L–treated mice. For IgA class switching in the absence of DCs, MACS-isolated splenic B cells were stimulated with 10 μg/ml anti-IgM and 20 μg/ml anti-CD40 (clone FGK4.5, UCSF Hybridoma Core) in the presence of TGFβ (2 ng/ml) and RA (100 nM) for 5 days.

For BMDCs, 5 × 106 BM cells were cultured in 10-cm tissue culture dishes in 10 ml of medium supplemented with supernatants from 3T3 cells transfected with the gene-encoding murine GM-CSF (granulocyte-macrophage colony-stimulating factor) for 7 days. Cells were treated with 1 μg/ml LTβR agonistic antibody (clone 3C8) and 100 nM RA every 3.5 days for 7 days.

Intravital two-photon laser-scanning microscopy (TPLSM) of PPs

Mice were anesthetized by intraperitoneal injection of 10 ml kg−1 saline containing xylazine (1 mg ml−1) and ketamine (5 mg ml−1). Maintenance doses of intramuscular injections of 4 ml kg–1 of xylazine (1 mg ml–1) and ketamine (5 mg ml–1) were given approximately every 30 min. An incision was made in the abdominal wall, and the small intestine was gently stretched and scanned by eye to identify PP structures. Only small areas (1 to 2 cm long) were exposed at any time. Once a PP was located, the area was embedded in warm saline and stabilized by placing a spring-loaded platform over the mouse and screwed down until the cover glass made contact with the PP. The tissue was placed with the interface between the intestinal lumen and PP facing upwards in an orientation that allowed maximal viewing of the SED. The mouse was placed on a Biotherm stage warmer at 37°C (Biogenics) for the duration of the imaging. Images were acquired with ZEN2009 (Carl Zeiss) with a 7MP two-photon microscope (Carl Zeiss) equipped with a Chameleon laser (Coherent). For video acquisition, a series of planes of 2- or 3-μm z-spacing spanning a depth of 30 to 69 μm were collected every 15 to 20 s. Excitation wavelengths were 850 to 890 nm. Because most of the transferred CFP+ B cells occupied the follicular compartment, we used automated tracks (generated by Imaris 7.4.2 ×64, Bitplane) to highlight the follicular region, as previously described (36). The SED was identified by the presence of CD11c-YFP DCs and by its typical shape and location above the follicles. The interfollicular regions, which are also rich with CD11c-YFP DCs, were identified on the basis of their distinct positioning and were excluded from analysis. Videos were made and analyzed with Imaris 7.4.2 ×64 (Bitplane). To track cells, surface seed points were created and tracked over time. Tracks were manually examined and verified. Data from cells that could be tracked for at least 15 min were used for analysis. Data presented in Fig. 6 were collected from four independent movies, some of which were split into 2 by 30 min segments and analyzed with Imaris (Bitplane AG), MATLAB (MathWorks), and MetaMorph software. In Fig. 6D, the contact time between B cells and DCs in the SED was measured manually (n = 150 B cells, derived from four independent movies). The behavior of a B cell engaged in prolonged interactions was defined as “Pause” (5 to 25 min of contact time), “Scan” (2 to 5 min of contact time), or “No contact” when spending less than 2 min in association with a DC. Statistical analysis was performed with Prism software (GraphPad Software).

Supplementary Materials

References and Notes

Acknowledgments: We thank M. Matloubian for help with the B16-Flt3L, J. Bluestone for RORγt-GFP mice, M. Krummel for Zbtb46-GFP mice, M. Rosenblum for Baft3−/− mice, R. Locksley for KN2 mice and Il4ra−/− mice, F. Kroese for HISM2 hybridoma, M. Miller for advice regarding intravital microscopy, Y. Xu and J. An for expert technical assistance, and O. Bannard for comments on the manuscript. The data presented in this manuscript are tabulated in the main paper and in the supplementary materials. UCSF has filed a patent (U.S. patent application 14/778,997) describing the αvβ8 blocking antibody used in this manuscript. D.S. and A.A. are listed as inventors. A.R. was a recipient of the Irvington postdoctoral fellowship from the Cancer Research Institute. J.G.C. is an investigator of the Howard Hughes Medical Institute. The work was supported in part by National Institute of Allergy and Infectious Diseases U19 grant AI077439 (to D.S.) and RO1 grants AI045073 and AI074847 (to J.G.C).
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