Research Article

Mucus Enhances Gut Homeostasis and Oral Tolerance by Delivering Immunoregulatory Signals

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Science  25 Oct 2013:
Vol. 342, Issue 6157, pp. 447-453
DOI: 10.1126/science.1237910

Guardian of the Gut

The intestine is able to tolerate continual exposure to large amounts of commensal bacteria and foreign food antigens without triggering an inappropriate inflammatory immune response. In the large intestine, this immunological tolerance is thought to occur via a physical separation between environment and host imposed by a continuous mucous layer built up from the secreted mucin protein, MUC2. However, in the small intestine, this mucous layer is porous, necessitating an additional layer of immune control. Shan et al. (p. 447, published online 26 September; see the Perspective by Belkaid and Grainger) now report that in the small intestine, MUC2 plays an active role in immunological tolerance by activating a transcription factor in resident dendritic cells, thereby selectively blocking their ability to launch an inflammatory response. This work identifies MUC2 as a central mediator of immune tolerance to maintain homeostasis in the gut and possibly at other mucosal surfaces in the body.

Abstract

A dense mucus layer in the large intestine prevents inflammation by shielding the underlying epithelium from luminal bacteria and food antigens. This mucus barrier is organized around the hyperglycosylated mucin MUC2. Here we show that the small intestine has a porous mucus layer, which permitted the uptake of MUC2 by antigen-sampling dendritic cells (DCs). Glycans associated with MUC2 imprinted DCs with anti-inflammatory properties by assembling a galectin-3–Dectin-1–FcγRIIB receptor complex that activated β-catenin. This transcription factor interfered with DC expression of inflammatory but not tolerogenic cytokines by inhibiting gene transcription through nuclear factor κB. MUC2 induced additional conditioning signals in intestinal epithelial cells. Thus, mucus does not merely form a nonspecific physical barrier, but also constrains the immunogenicity of gut antigens by delivering tolerogenic signals.

Mechanisms whereby the gut mucosa tolerates commensal bacteria and food antigens without developing inflammation remain elusive. Though traditionally viewed as a nonspecific barrier between the host and the environment, mucus also regulates gut homeostasis. The building block of gut mucus is MUC2, a gel-forming mucin secreted by goblet cells (GCs) (1). In the large intestine (LI), MUC2 prevents inflammation by generating an outer nonattached mucus layer inhabited by the microbiota and an inner mucus layer adherent to intestinal epithelial cells (IECs) and impervious to bacteria (1). The structure and function of mucus in the small intestine (SI) are less well understood.

The SI harbors bacteria that promote homeostasis by inducing Foxp3+ T regulatory (Treg) cells and B cell production of immunoglobulin A (IgA) antibodies (Abs) in Peyer’s patches (PPs) and mesenteric lymph nodes (MLNs) (2, 3). These responses involve sampling of bacteria by CD11c+ dendritic cells (DCs) (4), including macrophage-like CD103CD11b+CX3CR1+ DCs and myeloid CD103+CD11b+CX3CR1 DCs (2, 5, 6). In the lamina propria (LP), myeloid DCs sample soluble antigens from GC-associated passages and cooperate with lymphoid CD103+CD11bCX3CR1 DCs to generate Treg cells (79). Such antigen-sampling activities require a porous mucus barrier, raising questions as to how MUC2 prevents inflammation in the SI.

MUC2 Mitigates Inflammatory Responses in DCs

We first analyzed the structure of gut mucus in wild-type C57BL/6 mice. Whereas LI mucus formed a dense bilayered barrier that segregated bacteria from IECs, SI mucus was less organized and thus permitted the adhesion of bacteria to IECs (fig. S1, A to C). Consistent with the presence of some MUC2-coated bacteria on IECs and inside DCs (fig. S2, A to C), CX3CR1+ DCs from both PPs and SI-LP captured carboxyfluorescein succinimidyl ester (CFSE)-labeled MUC2 bound to fluorescent bacteria (fig. S3, A to C). In agreement with recent studies (6), some CD103+ DCs too captured MUC2-coated bacteria (fig. S3, A to D). Similarly, human monocyte-derived DCs internalized MUC2-bound bacteria across IECs sealed by occludin-containing tight junctions (fig. S4, A to D). Moreover, MUC2 was detected in human SI-LP CD103+ DCs proximal to GCs (Fig. 1A and movies S1 and S2).

Fig. 1 MUC2 imprints DCs with tolerogenic properties.

(A) Confocal microscopy of human SI-LP stained for CD11c, MUC2, CD103 and DNA-capturing 4′-6- diamidino-2-phenylindole (DAPI). Boxes and arrowheads: MUC2+CD103+ DCs. Original magnification, ×63. (B) Enzyme-linked immunosorbent assay (ELISA) of IL-12p70, IL-10, and TGF-β1 and flow cytometry (FC) of active RALDH in human DCs cultured for 2 days with or without LPS and/or MUC2. (C to E) FC of IFN-γ, Foxp3, CD4, and CFSE in human naïve CD4+ T cells cultured for 4 days with allogeneic DCs stimulated with or without LPS and/or MUC2 for 2 or 5 days in the absence or presence of control (ctr) IgG Ab, ctr vehicle, neutralizing Abs to TGF-β1 or IL-10, or LE540. (F) FC of CD103 and CX3CR1 on DCs cultured for 2 days with or without LPS and/or MUC2. Data summarize three experiments (error bars, SD; unpaired t test, *P < 0.05) or show one of four experiments with similar results.

In the presence of MUC2, human DCs exposed to bacteria or lipopolysaccharide (LPS) across IECs or directly incubated with LPS secreted less interleukin-12 (IL-12) (Fig. 1B and fig. S4E), a cytokine that induces proinflammatory interferon-γ (IFN-γ)-producing CD4+ T helper 1 (TH1) cells (2). This effect was comparably induced by human, murine, or porcine MUC2 and was not due to elevated endotoxin, impaired DC uptake of bacteria, or increased DC apoptosis (figs. S4F and S5, A to D). Unlike native MUC2, deglycosylated MUC2, a MUC2 peptide, or the mucin-interacting protein trefoil factor 3 did not inhibit LPS-induced IL-12 secretion (fig. S5E). MUC2 also impaired IL-12 as well as IL-6, IL-8, and tumor necrosis factor (TNF) transcription in response to bacterial Toll-like receptor (TLR) ligands such as LPS and flagellin or cytokines such as TNF (Fig. 1B and fig. S6, A to E). MUC2 elicited similar anti-inflammatory effects in monocyte-derived and myeloid CD1c+ DCs (Fig. 1B and fig. S6, A to E), which may include precursors of mucosal DCs (2, 8, 10). Thus, uptake of MUC2 causes carbohydrate-dependent attenuation of proinflammatory cytokine production by DCs.

MUC2 Delivers Tolerogenic Signals to DCs

Next, we established whether MUC2 induces IL-10, an anti-inflammatory cytokine that inhibits IL-12 and IFN-γ (2). Besides sustaining or augmenting IL-10 transcription and secretion in monocyte-derived DCs exposed to LPS or flagellin, MUC2 enhanced IL-10 secretion in LPS-activated myeloid CD1c+ DCs (Fig. 1B and fig. S6, C to E). MUC2 alone or combined with LPS also increased DC transcription and secretion of transforming growth factor–β1 (TGF-β1), a SMAD-signaling cytokine (Fig. 1B and figs. S6E and S7A) that helps the induction of Treg cells by tolerogenic CD103+ DCs (2, 7, 11). Moreover, MUC2 augmented DC transcription and activation of retinaldehyde dehydrogenase (RALDH or ALDH1), a 4-diethylaminobenzaldehyde (DEAB)-sensitive enzyme (Fig. 1B and figs. S6E and S7B) with A1-3 isoforms that help CD103+ DCs to induce Treg cells by converting dietary vitamin A into retinoic acid (RA) (2, 7, 11).

Accordingly, human DCs exposed to LPS in the presence of MUC2 decreased CD4+ T cell proliferation and IFN-γ production, but increased Foxp3 expression (Fig. 1, C and D, and fig. S8A). These effects were reversed by neutralizing Abs to IL-10 or TGF-β1 and by the RA antagonist LE540 (Fig. 1E). Despite up-regulating the antigen-presenting molecule human leukocyte antigen–DR (HLA-DR), MUC2 down-regulated the T cell costimulatory molecules CD80 and CD86 and the maturation molecule CD83 on LPS-activated DCs (fig. S8B). Consistent with its ability to autonomously induce TGF-β1 and RA, MUC2 induced regulatory DCs even in the absence of LPS priming (fig. S9, A and B). In addition to up-regulating CD103 and CX3CR1 on LPS-activated DCs (Fig. 1F), MUC2 stimulated CD103 expression and Treg cell–inducing signals in DCs undergoing transepithelial sampling of bacteria (fig. S10, A to C).

In the presence of MUC2, LPS-primed mouse bone marrow–derived DCs produced less IL-12 but more IL-10 (fig. S11A). When pulsed with the soluble protein ovalbumin (OVA) in the presence of MUC2, these DCs triggered less IFN-γ production by OVA-specific transgenic OT-II CD4+ T cells, which concurrently generated more Treg cells (fig. S11, B and C). MUC2 also attenuated flagellin-induced TNF but sustained or augmented IL-10 production in SI-LP CD103+ and CX3CR1+ DCs, respectively (Fig. 2A). Thus, MUC2 elicits tolerogenic IL-10, TGF-β1, and RA signals that somewhat vary in distinct subsets of DCs.

Fig. 2 MUC2 delivers anti-inflammatory signals to gut DCs.

(A) ELISA of TNF and IL-10 from mouse SI-LP DCs cultured for 2 days with or without flagellin and/or MUC2. (B) Light microscopy of Alcian blue–stained mucin and fluorescence in situ hybridization (FISH) of bacterial 16S ribosomal RNA in DAPI-stained SI-LP from wild-type (WT) and Muc2/ mice. Original magnification, ×10. (C to E) Quantitative real-time polymerase chain reaction (qRT-PCR) of mRNAs for TNF, IL-12p35 (Il12a), IL-12p40 (Il12b), IL-10, RALDH1 (Aldh1a1), and TGF-β1 in SI-LP DCs and FC of Foxp3 and CD4 on SI-LP T cells from WT and Muc2/ mice before and after oral antibiotics. RE, relative expression compared to Gapdh encoding glyceraldehyde 3-phosphate dehydrogenase. (F) ELISA of proliferation-induced bromodeoxyuridine (BrdU) and IFN-γ from OT-II cells activated for 5 days by OVA-pulsed SI-LP DCs from WT and Muc2/ mice with or without MUC2. Data summarize two experiments with ≥3 mice per group (error bars, SD; unpaired Student’s t test, *P < 0.05) or show one of four experiments with similar results.

MUC2 Enhances Gut Homeostasis

The immunoregulatory function of MUC2 was further explored in MUC2-deficient (Muc2/) mice (12). Compared to wild-type controls, Muc2/ mice showed more IEC-adherent bacteria and their SI-LP CD103+ and CX3CR1+ DCs expressed more TNF and IL-12, but less IL-10, TGF-β1, and ALDH1A1 and thus induced fewer Treg cells (Fig. 2, B and C, and fig. S12A). Accordingly, Muc2/ mice had fewer SI-LP Treg cells but more proinflammatory TH1 and IL-17–producing TH17 cells (fig. S12B). These changes were associated with increased bacteria-bound IgA Abs and SI-LP DC expression of B cell–activating factor of the TNF family (BAFF) and a proliferation-inducing ligand (APRIL) (fig. S12, C and D), two IgA-inducing cytokines induced by microbial and inflammatory signals (3). Compared to wild-type mice, Muc2/ mice sterilized of gut bacteria normalized SI-LP DC expression of BAFF and APRIL, but neither augmented SI-LP DC expression of IL-10, TGF-β1, and ALDH1A2 nor increased SI-LP Treg cells (Fig. 2E and fig. S12, D and E). Thus, perturbations of gut homeostasis in Muc2/ mice cannot be solely ascribed to increased IEC-adherent bacteria.

We then verified whether MUC2 regulates IECs. Compared to controls, IECs from Muc2/ mice expressed less IL-10, TGF-β1, ALDH1A1, and thymic stromal lymphopoietin (TSLP) (fig. S13A), which generates tolerogenic DCs (13, 14). Gut sterilization failed to restore wild-type–like amounts of IL-10, ALDH1A1, and TSLP, but did normalize TGF-β1 and RegIIIγ (fig. S13A), an antimicrobial protein induced by bacteria (15). In humans, IECs expressed more TSLP in response to MUC2 (fig. S13A), which confirmed the tolerogenic function of MUC2 on IECs.

The immune and barrier functions of MUC2 were further uncoupled by gavaging Muc2/ mice with MUC2 from wild-type controls. When exposed to MUC2, SI-LP CD103+ and CX3CR1+ DCs from Muc2/ mice reduced OVA-specific CD4+ T cell proliferation and IFN-γ production, as did SI-LP DCs from wild-type mice (Fig. 2F). Gavaged MUC2 did not restore a visible barrier, but its capture not only augmented IL-10, TGF-β, and ALDH1A1 and decreased IL-12 in SI-LP DCs, but also increased SI-LP Treg cells and reduced SI-LP TH1 and TH17 cells (Fig. 3, A and B, and fig. S14A). Similarly, gavaged MUC2 helped SI-LP CD103+ DCs from Muc2/ mice to induce more OVA-specific Treg cells after intragastric OVA immunization (Fig. 3C).

Fig. 3 MUC2 enhances gut homeostasis and oral tolerance.

(A) FC of CD103 and qRT-PCR of Il12a, Il12b, Il10, Aldh1a1, and Tgfb1 in SI-LP CD103+ DCs from WT or Muc2/ mice gavaged for 5 days with phosphate-buffered saline (PBS) or MUC2. RE, relative expression compared to Gapdh. (B) FC of Foxp3, IFN-γ, IL-17, and CD4 in SI-LP T cells from WT or Muc2/ mice treated as in (A). (C) FC of Foxp3 and CD4 in naïve OT-II cells cultured for 5 days with SI-LP CD103+ DCs from WT or Muc2/ mice treated as in (A) and intragastrically immunized with OVA. CD4+CD25+ OT-II cells from these cultures were incubated for 5 days with CFSE-labeled naïve OT-II cells and Abs to CD3 and CD28; divided CFSElow cells were quantified by FC. (D) ELISA of fecal OVA-specific IgG from WT and Muc2/ mice tolerized with PBS, OVA, or OVA plus MUC2 for 5 days and immunized as in (C). (E) ELISA of IFN-γ from OT-II cells incubated for 5 days with MLN CD103+ DCs from WT or Muc2/mice tolerized and immunized as in (D). (F) OVA-induced DTH in WT or Muc2 / mice tolerized as in (C) and subcutaneously immunized with OVA. (G) ELISA of proliferation-induced BrdU from SPL CD4+ T cells activated for 5 days with OVA-pulsed SPL DCs from WT or Muc2/ mice tolerized and immunized as in (F). (H to J) DTH, OVA-specific serum IgG and IgE, and SPL CD4+ T cell proliferation and IFN-γ secretion in WT or Muc2/ mice immunized and tolerized as in (F) after oral antibiotics. Data summarize two experiments with ≥4 mice per group (error bars, SD; unpaired Student’s t test, *P < 0.05) or show one of four experiments with similar results.

Notably, gavaged MUC2 enhanced the resistance of Muc2/ mice to dextran sodium sulfate (DSS), a colitogenic agent that disrupts the epithelial barrier. Although unable to attenuate inflammation-induced shortening of the LI, gavaged MUC2 ameliorated both clinical symptoms and histological lesions in DSS-treated Muc2/ mice, which showed less weight loss than wild-type or Muc2/ mice challenged with DSS in the absence of exogenous MUC2 (fig. S14, B to D). Thus, MUC2 attenuates bacteria-induced gut inflammation by delivering DC and IEC conditioning signals.

MUC2 Promotes Oral Tolerance

Oral tolerance consists of the attenuation of T and B cell responses to an antigen by prior oral administration of that antigen and involves MLN induction of Treg cells by migratory CD103+ DCs (2). Wild-type mice gavaged and later intragastrically immunized with OVA showed decreased gut B cell production of OVA-specific IgG Abs, which correlated with reduced IFN-γ production by OVA-reactive CD4+ T cells in response to OVA-pulsed MLNs, PPs, or SI-LP CD103+ DCs (Fig. 3, D and E, and fig. S15, A and B).

Tolerization also reduced OVA-induced delayed-type hypersensitivity (DTH)—an inflammatory skin reaction involving TH1 cells—as well as DC-dependent splenic CD4+ T cell proliferation and IFN-γ secretion following systemic immunization of Muc2/ mice with OVA (Fig. 3, F and G, and fig. S15, C and D). Unlike wild-type mice, Muc2/ mice gavaged with OVA developed neither intestinal nor systemic tolerance (Fig. 3, D to G, and fig. S15, A to D). However, Muc2/ mice restored oral tolerance when gavaged with OVA in the presence of MUC2 (Fig. 3, D to G, and fig. S15, A to D). The impairment of tolerance in Muc2/ mice was not merely due to bacteria-induced inflammation, as Muc2/ mice lacking gut bacteria did not attenuate OVA-specific DTH, IgG, and IgE responses after gavage with OVA (Fig. 3, H to J). Yet, tolerance was restored when OVA was gavaged in combination with MUC2 (Fig. 3, H to J). Thus, mucus actively constrains the immunostimulating properties of an oral soluble antigen.

MUC2 Binds a Galectin-3–Dectin-1–FcγRIIB Receptor Complex on DCs

Carbohydrates account for 80% of the weight of MUC2 (1) and may thus mediate its binding to DCs. Indeed, unlabeled glycosylated MUC2 inhibited the binding of CFSE-labeled native MUC2 to human DCs, whereas unlabeled deglycosylated MUC2 or a MUC2 peptide did not (Fig. 4A and fig. S16A). C-type lectin receptors (CLRs) and soluble galectins form carbohydrate-binding DC platforms with tolerogenic function (1620). Saturation of CLRs and galectins with mannan and lactose, respectively, attenuated MUC2 binding to DCs (Fig. 4A). DCs express galectin-1, -3 and -9 (21), but only galectin-3 interacted with glycosylated MUC2 and increased its binding to DCs through a process that was inhibited by MUC2 deglycosylation or Ab-mediated blockade of galectin-3 (Fig. 4, B to D). In agreement with their ability to bind galectin-3 following its secretion (16, 21), DCs increased both soluble and membrane-bound galectin-3 in response to LPS (fig. S16B). Thus, DCs may form a galectin-3–based MUC2-binding platform after sensing gut microbial signals.

Fig. 4 MUC2 binds galectin-3, Dectin-1, and FcγRIIB on DCs.

(A) FC of CFSE-MUC2 on human DCs preincubated with unlabeled native MUC2, deglycosylated (dgl) MUC2, MUC2 peptide, mannan, or lactose. Percent of MUC2 binding compared to medium alone. (B) ELISA of native or dgl MUC2 interaction with galectins. (C) CFSE-MUC2 or CFSE-dgl MUC2 binding to DCs preincubated with PBS, human serum albumin (HSA), or galectin-3. (D) CFSE-MUC2 binding to DCs before and after preincubation with a fluorescent Ab to galectin-3. (E) IFA of CD11c, galectin-3, Muc2, and DAPI in mouse PP sections. Original magnification, ×5 (upper panel) and ×63 (bottom panel). (F) qRT-PCR of mRNA for galectin-3 in DC subsets from mouse PPs, SI-LP, and SPL. RE, relative expression compared to Gapdh. (G) Immunoprecipitation (IP) with control or anti–galectin-3 Abs of proteins from human DCs treated without (ctr) or with MUC2 for 30 min, followed by immunoblotting of FcγRIIB, Dectin-1, and galectin-3. (H) ELISA of IL-12p70 from human DCs exposed to scrambled (ctr) or LGALS3 (galectin-3), FCGR2B (FcγRIIB), or CLEC7A (Dectin-1) small interfering RNAs (siRNAs) and cultured with or without LPS and/or MUC2 for 2 days. (I) Binding of CFSE-MUC2 to SI-LP DCs from WT, Lgals3/, Clec7A/, or Fcgr2b/ mice. Data summarize experiments with three donors or three mice from each strain (error bars, SD; unpaired t test, *P < 0.05) or show one of three experiments with similar results.

Accordingly, PP and SI-LP CD103+ and CX3CR1+ DCs produced more galectin-3 than splenic DCs did and showed intracellular galectin-3 colocalized with MUC2 (Fig. 4, E and F, and fig. S16, C to E). SI-LP DCs also displayed more surface galectin-3 (Fig. 4F and fig. S16F), part of which may have an epithelial origin (21). Indeed, galectin-3 was detected in both IECs and GCs, and its secretion augmented upon IEC exposure to bacteria (fig. S16, G and H). Consequently, DCs acquired more surface galectin-3 after exposure to IEC supernatant or migration across IECs (fig. S16, I and J).

We then determined how soluble galectin-3 anchors itself and MUC2 to DCs. Galectin-3 binds Dectin-1, a CLR that induces tolerogenic DCs and enhances gut homeostasis by recognizing fungal carbohydrates (16, 17, 20). Dectin-1 further interacts with the anti-inflammatory receptor FcγRIIB in response to glycans from therapeutic IgG Abs (22). In human embryonic kidney 293 cells, MUC2 bound transfected Dectin-1 and FcγRIIB in the presence of galectin-3 (fig. S17, A and B). In DCs, MUC2 binding correlated with increased surface amounts of galectin-3, Dectin-1, and FcγRIIB and caused Dectin-1 recruitment to a preformed galectin-3–FcγRIIB complex (Fig. 4G and fig. S17C). Accordingly, DCs lacking galectin-3, Dectin-1, or FcγRIIB neither effectively bound MUC2 nor decreased IL-12 production in response to MUC2 (Fig. 4, H and I, and fig. S17, D to G).

In wild-type mice, SI-LP CD103+ DCs expressed less Dectin-1 but more galectin-3 and FcγRIIB than CX3CR1+ DCs did (Fig. 5A). Compared to Lgals3 (galectin-3)/, Clec7a (Dectin-1)/, or Fcgr2b (FcγRIIB)/ mice, wild-type mice had a comparable SI mucus layer, but showed more SI-LP DCs containing MUC2 (Fig. 5B and fig. S18, A and B). Unlike IECs, these SI-LP DCs did not express MUC2 (Fig. 5C), which confirmed that gut DCs acquire MUC2 from the external environment. Compared to controls, Lgals3/, Clec7a/, or Fcgr2b/ mice had SI-LP DCs that produced more IL-12, but less ALDH1A1 and TGF-β1 and did not respond to MUC2 (Fig. 5D and fig. S19). These mice also had fewer SI-LP Treg cells, more SI-LP TH1 cells, and impaired tolerogenic T and B cell responses to OVA (Fig. 5, E to G). Thus, MUC2 may enhance gut homeostasis and tolerance by assembling a signal-transducing Dectin-1–FcγRIIB complex on DCs with the help of soluble galectin-3.

Fig. 5 Galectin-3, Dectin-1, and FcγRIIB enhance gut homeostasis and oral tolerance.

(A) FC of galectin-3, FcγRIIB, and Dectin-1 on SI-LP DCs from WT mice. Galectin-3 was measured in permeabilized DCs. MFI, mean fluorescence intensity. (B) Quantification of MUC2+ SI-LP DCs from WT, Lgals3/, Clec7A/, or Fcgr2b/ mice by IFA of 10 to 12 SI-LP sections per group. (C) qRT-PCR of Muc2 in IECs and SI-LP DCs from WT mice. RE, relative expression compared to Gapdh. (D and E) qRT-PCR of Il12a, Il12b, Aldh1a1, and Tgfb1 in SI-LP DCs and FC of Foxp3 and IFN-γ in SI-LP CD4+ T cells from WT, Lgals3/, Clec7A/, or Fcgr2b/ mice. RE, relative expression compared to Gapdh. (F) DTH and ELISA of OVA-specific IgE in WT, Lgals3/, Clec7A/, or Fcgr2b/ mice intragastrically tolerized with PBS or OVA for 5 days and subcutaneously immunized with OVA. (G) ELISA of proliferation-induced BrdU from SPL CD4+ T cells activated for 5 days by OVA-pulsed SPL DCs from WT, Lgals3 /, Clec7A/, or Fcgr2b/ mice tolerized and immunized as in (F). Data summarize two experiments with ≥4 mice per group (error bars, SD; unpaired Student’s t test, *P < 0.05) or show one of four experiments with similar results.

MUC2 Tolerizes DCs by Inducing β-Catenin

How do Dectin-1 and FcγRIIB convey tolerogenic signals to DCs exposed to MUC2? Dectin-1 phosphorylates AKT (23) and may thus induce β-catenin, an AKT-regulated transcription factor required by gut tolerogenic DCs (24). Indeed, MUC2 alone or combined with LPS phosphorylated AKT and glycogen synthase kinase–3β (GSK-3β) (Fig. 6A), two events that cause GSK-3β inactivation and inhibition of β-catenin degradation (24, 25). Consequently, MUC2 elicited cytoplasmic accumulation and nuclear translocation of β-catenin in wild-type but not Clec7a/ DCs (Fig. 6A and fig. S20, A and B).

Fig. 6 MUC2 impairs NF-κB–driven inflammatory signals via β-catenin.

(A) WB of cytoplasmic or nuclear phospho (p)-AKT, AKT, pGSK-3β, GSK-3β, β-catenin (β-cat), dephospho (dp)-β-cat, actin, and octamer-1 (Oct-1) from human DCs cultured with or without LPS and/or MUC2 for 10 min. (B) Electrophoretic mobility gel shift assay of IL12A-bound NF-κB p65-p50 and consensus DNA-bound Oct-1 in DCs cultured as in (A) for 3 hours. (C) Chromatin IP of IL12A-bound NF-κB p65 in DCs cultured as in (A) for 3 hours. RDQ, relative DNA quantity. (D) IP with control (ctr) IgG Ab or anti–dp-β-cat Ab of nuclear proteins from DCs cultured with or without LPS and/or MUC2 for 10 min, followed by WB of NF-κB p65 and dp-β-cat. (E) IL12A transcription in DCs cultured as in (A) for 2 days. (F and G) NF-κB–mediated transcription and ELISA of IL-12p70 and IL-10 in DCs cultured as in (E) in the presence of scrambled (ctr) or CTNNB1 (β-cat) siRNAs. (H) IFA of pAKT, pGSK-3β, β-cat, galectin-3, CD11c, and DAPI in mouse PPs. Original magnification, ×5. Data show one of three experiments yielding similar results or summarize three experiments (error bars, SD; unpaired t test, *P < 0.05).

In some tumors, β-catenin interacts with nuclear factor κB (NF-κB) p65 to impede the binding of activating p50-p65 complexes to death-inducing genes (25). In DCs, MUC2 alone or coupled with LPS enhanced nuclear β-catenin interaction with NF-κB p65 and decreased NF-κB p50-p65 binding to both minimal and IL12 gene promoters, thus impairing their transcription (Fig. 6, B to E, and fig. S20C). Accordingly, β-catenin deficiency impeded MUC2-mediated inhibition of LPS-induced IL-12 but not IL-10 transcription and/or secretion, whereas β-catenin overexpression dampened TNF-induced NF-κB–driven transcription (Fig. 6, F and G, and fig. S20, D to F). Consistent with these data, galectin-3–expressing DCs from PPs contained abundant β-catenin in addition to activated AKT and inactive GSK-3β (Fig. 6H).

Besides AKT, Dectin-1 phosphorylates SYK, which activates NF-κB through a pathway mitigated by FcγRIIB via SH2 domain–containing inositol 5-phosphatase-1 (SHIP-1) (22, 26). SYK also activates cAMP responsive element binding protein (CREB), a calcium-dependent IL-10–inducing protein that removes the co-activator CREB-binding protein (CBP) from DNA-bound NF-κB (27, 28). Besides triggering SYK and SHIP-1 phosphorylation, MUC2 alone or combined with LPS decreased NF-κB p65 but not p50 nuclear translocation and increased galectin-dependent calcium fluxes, phosphorylation of CREB-targeting AKT, ERK1/2 and p38 kinases, phosphorylation and nuclear translocation of CREB, binding of CREB to IL10 and IL12 promoters, and loss of CBP from the IL12 promoter (figs. S20G and S21, A to G). Thus, similar to hyperglycosylated IgG Abs used to treat autoimmune disorders (22), MUC2 may recruit SHIP-1 via FcγRIIB to constrain proinflammatory NF-κB but not tolerogenic CREB signals emanating from Dectin-1 and SYK. In addition to inducing IL-10, CREB may cooperate with Dectin-1–induced β-catenin to inhibit NF-?B–dependent IL-12 production.

Conclusions

We have shown here that MUC2 enhances gut homeostasis and oral tolerance by conditioning DCs and IECs. Antigen-sampling DCs assemble galectin-3, Dectin-1, and FcγRIIB to acquire MUC2 across IECs and possibly from GCs (fig. S22A). This MUC2 receptor complex suppresses inflammatory but not tolerogenic DC responses by inhibiting NF-κB via β-catenin (fig. S22B). How DCs tune down these signals during infection remains unclear, but pathogen-induced perturbations of MUC2 glycosylation and polymerization patterns may be involved (1). A full understanding of the immunoregulatory function of MUC2 could help to devise better vaccines and treatments against infections and food allergies and to unravel how alterations of MUC2 and its receptors aggravate inflammatory bowel disease (1, 20), thus leading to safer therapies against this disorder.

Supplementary Materials

www.sciencemag.org/content/342/6157/447/suppl/DC1

Materials and Methods

Acknowledgments

Figs. S1 to S22

Tables S1 to S8

References (2937)

Movies S1 and S2

References and Notes

  1. Acknowledgments: This study was supported by the National Institute of Allergy and Infectious Diseases, NIH (AI61093, AI57653, AI95613, AI96187 and AI74378 to A.C. and AI073899, DK072201 and AI095245 to J.M.B.) and by Redes Temáticas de Investigación Cooperativa en Salud/Fondo Europeo de Desarrollo Regional (RD12/0036/0054 to A.B). pRSETb-mRFP used for red bacteria is under a materials transfer agreement with R. Y. Tsien at the University of California, San Francisco, and Howard Hughes Medical Institute. The data presented in this manuscript are tabulated in the main paper and the supplementary materials.
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