Yosuke Shimodaira,Shyam K. More,Hussein Hamade,Anna Y. Blackwood,Jay P. Abraham,Lisa S. Thomas,Jordan H. Miller,Dalton T. Stamps,Sofi L. Castanon,Noam Jacob,Connie W.Y. Ha,Suzanne Devkota,David Q. Shih,Stephan R. Targan,Kathrin S. Michelsen
DR3 Regulates Intestinal Epithelial Homeostasis and Regeneration After Intestinal Barrier Injury
Intestinal Permeability,IEC Proliferation,Epithelial Barrier,Tissue Regeneration
Background & Aims Tumor necrosis factor (TNF) superfamily member tumor necrosis factor–like protein 1A (TL1A) has been associated with the susceptibility and severity of inflammatory bowel diseases. However, the function of the tumor necrosis factor–like protein 1A and its receptor death receptor 3 (DR3) in the development of intestinal inflammation is incompletely understood. We investigated the role of DR3 expressed by intestinal epithelial cells (IECs) during intestinal homeostasis, tissue injury, and regeneration. Methods Clinical phenotype and histologic inflammation were assessed in C57BL/6 (wild-type), Tl1a-/- and Dr3-/- mice in dextran sulfate sodium (DSS)-induced colitis. We generated mice with an IEC-specific deletion of DR3 (Dr3ΔIEC) and assessed intestinal inflammation and epithelial barrier repair. In vivo intestinal permeability was assessed by fluorescein isothiocyanate dextran uptake. Proliferation of IECs was analyzed by bromodeoxyuridine incorporation. Expression of DR3 messenger RNA was assessed by fluorescent in situ hybridization. Small intestinal organoids were used to determine ex vivo regenerative potential. Results Dr3-/- mice developed more severe colonic inflammation than wild-type mice in DSS-induced colitis with significantly impaired IEC regeneration. Homeostatic proliferation of IECs was increased in Dr3-/- mice, but blunted during regeneration. Cellular localization and expression of the tight junction proteins Claudin-1 and zonula occludens-1 were altered, leading to increased homeostatic intestinal permeability. Dr3ΔIEC mice recapitulated the phenotype observed in Dr3-/- mice with increased intestinal permeability and IEC proliferation under homeostatic conditions and impaired tissue repair and increased bacterial translocation during DSS-induced colitis. Impaired regenerative potential and altered zonula occludens-1 localization also were observed in Dr3ΔIEC enteroids. Conclusions Our findings establish a novel function of DR3 in IEC homeostasis and postinjury regeneration independent of its established role in innate lymphoid cells and T-helper cells.
DR3 Regulates Intestinal Epithelial Homeostasis and Regeneration After Intestinal Barrier Injury

Tumor necrosis factor (TNF) superfamily member tumor necrosis factor–like protein 1A (TL1A) has been associated with the susceptibility and severity of inflammatory bowel diseases. However, the function of the tumor necrosis factor–like protein 1A and its receptor death receptor 3 (DR3) in the development of intestinal inflammation is incompletely understood. We investigated the role of DR3 expressed by intestinal epithelial cells (IECs) during intestinal homeostasis, tissue injury, and regeneration.

Clinical phenotype and histologic inflammation were assessed in C57BL/6 (wild-type), Tl1a-/- and Dr3-/- mice in dextran sulfate sodium (DSS)-induced colitis. We generated mice with an IEC-specific deletion of DR3 (Dr3ΔIEC) and assessed intestinal inflammation and epithelial barrier repair. In vivo intestinal permeability was assessed by fluorescein isothiocyanate dextran uptake. Proliferation of IECs was analyzed by bromodeoxyuridine incorporation. Expression of DR3 messenger RNA was assessed by fluorescent in situ hybridization. Small intestinal organoids were used to determine ex vivo regenerative potential.

Dr3-/- mice developed more severe colonic inflammation than wild-type mice in DSS-induced colitis with significantly impaired IEC regeneration. Homeostatic proliferation of IECs was increased in Dr3-/- mice, but blunted during regeneration. Cellular localization and expression of the tight junction proteins Claudin-1 and zonula occludens-1 were altered, leading to increased homeostatic intestinal permeability. Dr3ΔIEC mice recapitulated the phenotype observed in Dr3-/- mice with increased intestinal permeability and IEC proliferation under homeostatic conditions and impaired tissue repair and increased bacterial translocation during DSS-induced colitis. Impaired regenerative potential and altered zonula occludens-1 localization also were observed in Dr3ΔIEC enteroids.

Our findings establish a novel function of DR3 in IEC homeostasis and postinjury regeneration independent of its established role in innate lymphoid cells and T-helper cells.

SummaryExpression of death receptor 3 in intestinal epithelial cells regulates intestinal permeability and cellular localization of tight junction proteins during homeostasis. As a result, Dr3ΔIEC mice are more susceptible to acute colitis and show severely impaired epithelial barrier regeneration. Intestinal homeostasis is tightly regulated by complex interactions between the intestinal mucosal immune network, intestinal epithelial cells (IECs), and intestinal microbiota. Dysfunction in any of these mechanisms can lead to the development of intestinal inflammation and inflammatory bowel diseases (IBD). IBD results from integrated interactions between genetic susceptibility and severity genes that have an impact on the intestinal microbial environment and dysregulate innate and adaptive immune responses of the host against commensal microorganisms. IBD-associated single-nucleotide polymorphisms are enriched in genes that regulate epithelial barrier function, innate and adaptive immunity that could affect immune responses to commensal bacteria. Tumor necrosis factor (TNF)-like ligand 1A (TL1A), a TNF superfamily member originally described as a cytokine produced by endothelial and antigen-presenting cells, provides co-stimulatory signals to activated T cells and enhances their function via its receptor death receptor 3 (DR3).3, 4, 5 Importantly, genetic polymorphisms in the genes TNFSF15 encoding TL1A and TNFRSF25 encoding DR3 have been associated with susceptibility to and severity of IBD.6, 7, 8, 9, 10 Crohn’s disease (CD) patients with TNFSF15 risk haplotypes have increased expression of TL1A in intestinal mucosa and peripheral blood monocytes and are more prone to develop complications such as stricturing CD.11, 12, 13 Mouse models that constitutively overexpress TL1A develop similar clinical phenotypes as human CD patients with risk variants. These mice develop spontaneous ileitis, intestinal fibrosis, and exacerbated dextran sulfate sodium (DSS)-induced chronic colitis.,14, 15, 16 However, mouse models of deficiencies of Tl1a or its receptor Dr3, as well as using neutralizing TL1A antibodies, have led to conflicting results. Although neutralizing TL1A antibodies prevent and reverse acute and chronic intestinal inflammation and reduce the development of intestinal fibrosis,,, Tl1a or Dr3 deficiency exacerbated acute DSS-induced colitis., The role of TL1A and DR3 in these models has been attributed mainly to its function on different Thelper (TH) subsets, regulatory T cells, and innate lymphoid cells (ILCs).,19, 20, 21 In addition, it has been shown that DR3 is expressed on nonimmune cells, including IECs and fibroblasts, which could contribute to intestinal inflammation and fibrosis., However, the exact mechanisms of how TL1A and DR3 affect intestinal inflammation, especially the role of DR3 expressed by IECs, and its contribution to the development of intestinal inflammation and tissue repair, still largely is unknown. This study aimed to investigate the role of TL1A–DR3 signaling on IECs and its contribution to intestinal inflammation. IECs regulate the intestinal barrier function and protect against pathogenic and commensal luminal organisms. One of the mechanisms of IECs to maintain the physical barrier between luminal contents and the lamina propria is the expression of tight junction proteins, which regulate an impenetrable intact barrier. Dysregulated tight junction expression or intracellular localization elicits increased barrier permeability and susceptibility to DSS-induced murine colitis.24, 25, 26 IEC proliferation, migration, and tight junction protein expression can be regulated by proinflammatory cytokines during inflammation. For example, TNF-α, via its receptor Tumor necrosis factor receptor type II, affects a multitude of IEC functions including expression of tight junction proteins and maintenance of barrier function.27, 28, 29 However, whether TL1A-DR3 signaling affects IEC function during homeostasis and inflammation still is unknown. Here, we show that Dr3 deficiency exacerbates acute and chronic DSS-induced colitis characterized by impaired intestinal epithelial regeneration, while Tl1a-/- mice developed a similar degree of colitis as wild-type (WT) mice. Under homeostatic conditions, we observed increased intestinal permeability, dysregulated expression of tight junction proteins, and uncontrolled IEC proliferation in Dr3-/- mice compared with WT mice. To elucidate the role of DR3 signaling in IECs, we generated mice with IEC-specific deletion of Dr3 (Dr3ΔIEC). Dr3ΔIEC mice recapitulate the phenotype of Dr3-/- mice under homeostatic conditions and during DSS-induced colitis. We observed increased intestinal permeability, altered tight junction protein expression, and increased IEC proliferation in Dr3ΔIEC mice under homeostatic conditions. Strikingly, acute DSS colitis is exacerbated in Dr3ΔIEC mice characterized by impaired intestinal epithelial regeneration, bacterial translocation to mesenteric lymph nodes (MLNs), and systemic inflammation. Consistent with these findings, we observed reduced regenerative potential of Dr3ΔIEC ex vivo enteroid cultures and altered cellular localization of tight junction protein zonula occludens 1 (ZO-1). Our data establish a critical novel function of DR3 in regulating IEC homeostasis and uncover a mechanistic role of DR3 in IEC regeneration during mucosal injury.

First, we investigated the role of TL1A–DR3 signaling in a chronic model of colitis. Administration of 4 cycles of DSS in drinking water leads to the development of chronic colonic inflammation with characteristic TH1 and TH17 responses. To investigate the specific roles of TL1A and DR3, we induced chronic DSS-induced colitis in TL1A-deficient (Tl1a-/-), DR3-deficient (Dr3-/-), and WT mice, and determined the clinical and pathologic characteristics of colitis. Interestingly, we observed bloody stool and severe diarrhea during each DSS cycle in Dr3-/- mice, while Tl1a-/- and WT mice had less severe symptoms (Figure 1A). Consistent with the stool phenotype, histologic scores as assessed by H&E staining showed that Dr3-/- mice developed more severe inflammation in the rectum and colon (Figure 1B and C). Interestingly, the significantly higher histologic score in Dr3-/- mice was driven mainly by a significantly higher regeneration subscore (score, 0–4, with 4 showing no tissue repair) compared with WT and Tl1a-/- mice, which is indicative of impaired epithelial regeneration in Dr3-/- mice (Figure 1C and D). DR3 is known to be expressed primarily in CD4+ T cells and TL1A engagement induces the activation of several TH subsets, including TH1, TH17, TH9, and regulatory T cells (Tregs).,30, 31, 32, 33, 34 However, we did not observe a significant difference in cell numbers or expression of the cytokines interferon gamma (IFN-gamma), interleukin (IL)17A, IL22, or IL10 between groups (Figure 2). We observed that forkhead box P3 (FoxP3+) Tregs were reduced significantly in MLNs of Dr3-/- mice but not in the lamina propria (LP), as has been reported previously in acute DSS-induced colitis (Figure 2C). Taken together, Dr3-/- mice develop exacerbated DSS-induced chronic colitis while intestinal inflammation in Tl1a-/- mice is comparable with WT mice. However, the more severe colonic inflammation in Dr3-/- mice was not associated with significant differences in CD4+ TH cell responses.

Based on our results showing that the severe colonic inflammation in DSS-induced chronic colitis in Dr3-/- mice correlated with impaired intestinal epithelial regeneration, we hypothesized that Dr3-/- mice have impaired IEC function. To test for impaired epithelial barrier function, we first determined intestinal permeability for these mice during DSS colitis. Dr3-/- mice showed significantly higher fluorescein isothiocyanate (FITC) dextran permeability compared with WT mice (Figure 3A). To test whether the intestinal barrier is impaired in Dr3-/- mice under homeostatic conditions, we next performed intestinal permeability assays in 8-week-old untreated mice that did not show any signs of spontaneous intestinal inflammation. We observed that Dr3-/- mice have significantly increased permeability even under homeostatic uninflamed conditions (Figure 3B). Next, we investigated whether DR3 is involved in the maintenance of IECs. First, we assessed the expression of DR3 in IECs. A previous publication showed expression of DR3 in epithelial cell lines. However, antibody-based detection of DR3 has been difficult in the past because of a lack of specificity of available anti-DR3 antibodies and we were unable to specifically detect DR3 with commercially available antibodies by immunofluorescence. Therefore, we performed single-molecule fluorescent in situ hybridization (sm-FISH) using a custom-made specific probe for murine Dr3 messenger RNA (mRNA) and observed expression of DR3 in ileal and colonic IECs that was not observed in Dr3-/- intestinal tissue (Figure 4A and B). Furthermore, Dr3 mRNA also was expressed in IECs from noninflamed areas of surgical specimens collected from CD patients (Figure 4C). We compared the expression of Dr3 mRNA in inflamed and noninflamed ileal tissue sections from CD patients and did not observe differences in DR3 expression in IECs. Major differences in DR3 expression between inflamed and noninflamed tissue were observed in the LP, most likely as a result of infiltration of immune cells (Figure 4D). Furthermore, we analyzed the expression of Dr3 mRNA in colonic tissue sections from a patient with diverticulitis (Figure 4E). Next, we assessed the baseline phenotype of IECs in Dr3-/- mice. We observed significantly increased villi length in the ileum and crypt length in the colon of Dr3-/- mice compared with WT mice (Figure 4F and G). Next, we assessed the proliferation and migration of IECs in Dr3-/- mice after bromodeoxyuridine (BrdU) incorporation under homeostatic conditions. Under homeostatic conditions, proliferating cells can be visualized by BrdU incorporation in transit-amplifying zones 2.5 hours after BrdU injection. At baseline, Dr3-/- mice have significantly higher numbers of proliferating IECs compared with WT mice in the ileum and colon (Figure 4H and I). These data indicate that DR3 regulates the proliferation of IECs. To determine if the increase in homeostatic proliferation in Dr3-/- mice is microbiota-dependent, we treated Dr3-/- mice with broad-spectrum antibiotics and assessed BrdU incorporation. We did not observe any significant differences in the numbers of proliferating IECs in the ileum or colon of antibiotic-treated Dr3-/- mice (Figure 4J). To investigate molecular mechanisms leading to increased permeability in Dr3-/- mice, we next studied the expression of several tight junction proteins at baseline. Compared with WT and Tl1a-/-, ZO-1 expression in Dr3-/- mice is less continuous at the apical side of IECs and we observed more pronounced expression of ZO-1 and claudin-1 in the cytoplasm and at the basal–lateral membrane in Dr3-/- IECs (Figure 5A). We did not observe differences in the localization of claudin-2 or occludin in Dr3-/- compared with WT IECs (Figure 5A). We observed significantly increased mRNA expression of claudin-1, claudin-4, and myosin light-chain kinase (Mlck), and increased protein expression for claudin-1 in Dr3-/- IECs, but no differences in the expression of other tight junction proteins (claudin-2, -3, and -15) (Figure 5B and C). These data indicate that DR3 but not TL1A deficiency affects the subcellular location of tight junction proteins and their expression level. Collectively, DR3 deficiency affects IEC homeostasis, altering the rate of proliferation and turnover and distribution of tight junction proteins throughout the small and large intestine, leading to increased intestinal permeability.

To further investigate the increased IEC proliferation in Dr3-/- mice, we used the acute DSS model to assess epithelial cell regeneration. During acute DSS colitis, Dr3-/- mice developed more severe intestinal inflammation compared with WT or Tl1a-/- mice, and had a higher stool score driven by a higher stool blood subscore (Figure 6A). Although microscopically injured tissue was covered by regenerating epithelial cells in WT mice, regeneration was strikingly impaired in Dr3-/- mice, which also was reflected by a significantly increased histology and regeneration score (Figure 6B). We also observed an increase in the size of spleens and significantly increased splenic cell numbers in Dr3-/- mice (Figure 6C), suggesting systemic inflammation caused by epithelial damage in these mice. In contrast, we did not observe any significant differences in LP total cell numbers, CD4+ T-cell numbers, secretion of IFN-gamma, IL17A, or IL22 from lamina propria mononuclear cells (LPMCs), percentages of IL22+ CD4+ T cells, or IL22+ retinoid orphan receptor gamma t (RORγt+) innate lymphoid cell type 3 (ILC3s) in Dr3-/- compared with WT mice (Figure 6D–G). Interestingly, BrdU+ cells were decreased in Dr3-/- compared with WT mice under inflammatory conditions (Figure 7A and B). Although WT mice responded to acute injury of the epithelial layer by significantly increased proliferation, in Dr3-/- mice the number of BrdU+ cells was not increased over homeostatic proliferation, suggesting a blunted compensatory proliferation in response to injury in Dr3-/- mice (Figure 7C). These data indicate that while under homeostatic conditions DR3 deficiency leads to an increase in IEC proliferation, the regeneration of IECs is impaired in Dr3-/- mice after tissue injury. Next, we generated intestinal enteroids from the ileum of WT, Tl1a-/-, and Dr3-/- mice to assess the functions of intestinal stem cells in ex vivo enteroid cultures. DR3 but not TL1A deficiency significantly reduced the enteroid surface area (Figure 8A and B). In contrast, the number of crypts was reduced significantly in Tl1a-/- and Dr3-/- enteroids (Figure 8A and B). Next, we analyzed ZO-1 expression by immunofluorescence staining in these enteroids. Tl1a-/- and Dr3-/- enteroids had significantly decreased ZO-1 fluorescence intensity compared with WT enteroids (Figure 8C). These results suggest compromised intestinal stem cell functions and expression of tight junction protein in Dr3-/- enteroids, while Tl1a-/- enteroid formation and surface area, indicative of proliferative response, is comparable with WT enteroids. However, Tl1a-/- enteroids have reduced de novo crypt formation, indicative of IEC differentiation, and ZO-1 expression compared with WT enteroids. Next, we analyzed mRNA expression of TL1A and DR3 in enteroids. Both TL1A and DR3 mRNA are expressed in WT enteroids (Figure 8D). To investigate direct effects of TL1A on enteroids, we first treated WT enteroids with TL1A. We observed significantly reduced enteroid surface area after TL1A treatment (Figure 8E). Next, we treated WT enteroids with a combination of TL1A and IL22. Although IL22 increased the enteroid surface area in a dose-dependent manner, as has been shown previously, addition of TL1A resulted in a similar increase in enteroid surface area compared with IL22 treatment alone (Figure 8E). These data suggest that IL22 treatment reverses the negative effects TL1A has on organoid formation. Finally, we treated enteroids derived from WT and Dr3-/- mice with IL22. We observed a significant increase in enteroid surface area and crypts per enteroid in both WT and Dr3-/- enteroids treated with IL22, suggesting that IL22 signaling is independent of DR3 signaling (Figure 8F).

To address the functional importance of DR3 expressed on IECs we generated mice with DR3 deficiency in IECs by crossing Villin-Cre+ with Dr3fl/fl mice (Dr3ΔIEC). We confirmed the cell-type–specific deletion of DR3 in Dr3ΔIEC mice (Figure 9A). We observed significantly increased intestinal permeability in Dr3ΔIEC compared with Dr3fl/fl mice under homeostatic conditions (Figure 9B). Furthermore, we also observed an increased serum IgA concentration in Dr3ΔIEC mice while serum IgG concentrations were not significantly different between Dr3ΔIEC and Dr3fl/fl mice, further corroborating an impaired intestinal barrier (Figure 9C). We did not observe any differences in fecal IgA concentrations between genotypes (Figure 9C). We observed more pronounced expression of claudin-1 in the cytoplasm and at the basal–lateral membrane (Figure 9D), while apical claudin-1 expression in tight junctions was diminished. Overall, claudin-1 protein expression in Dr3ΔIEC IECs was increased significantly (Figure 9E). IEC proliferation was increased significantly in Dr3ΔIEC mice (Figure 9F and G). DR3 signaling activates caspase 8 through the Fas Associated Via Death Domain pathway, which can induce apoptosis in certain cell types., To determine the rate of homeostatic cell death, we performed terminal deoxynucleotidyl transferase–mediated deoxyuridine triphosphate nick-end labeling (TUNEL) staining, which detects apoptotic and necroptotic cell death. The number of TUNEL-positive cells was similar in ileal and colonic sections of Dr3fl/fl and Dr3ΔIEC mice, indicating that cell death does not contribute to the compromised intestinal barrier in Dr3ΔIEC mice (Figure 9H). These data indicate that DR3 expressed by IECs contributes directly to homeostatic proliferation and barrier function. Next, we determined if DR3 deficiency in IECs leads to differences in LP immune cell compositions. We did not observe any significant differences in LP total cell numbers; CD4+ T-cell numbers; secretion of IFN-gamma, IL17A, or IL22 from LPMCs; percentages of IFN-gamma+, IL17A+, IL22+, IL10+ CD4+ T cells; or IL22+ RORγt+ ILC3s under homeostatic conditions (Figure 10A–D). We also did not observe any significant difference in the relative abundance of segmented filamentous bacteria (SFB), a main inducer of IL22, between genotypes (Figure 10E). Similar to the results we obtained from Dr3-/- mice, we did not observe any significant differences in the number of proliferating IECs in the ileum or colon of antibiotic-treated Dr3ΔIEC mice (Figure 10F).

The regenerative potential of intestinal stem cells was assessed by ileal ex vivo enteroid cultures. Enteroids prepared from Dr3ΔIEC mice had significantly reduced surface area compared with Dr3fl/fl enteroids, indicating reduced proliferative potential (Figure 11A and B). The overall enteroid formation potential also was reduced in Dr3ΔIEC enteroids (Figure 11C). We observed significantly reduced numbers of enteroids with 3 or more than 4 buds per enteroid in Dr3ΔIEC compared with Dr3fl/fl enteroids (Figure 11D). Reduced surface area and impaired development of crypts in Dr3ΔIEC enteroids suggest an impaired regenerative response. Staining of these enteroids with ZO-1 confirmed our findings of mislocalized tight junction proteins in Dr3-/- and Dr3ΔIEC ileal tissues (Figures 5A and 7E). We observed discontinuous staining of ZO-1 in Dr3ΔIEC enteroids, while ZO-1 staining was continuous and intact in Dr3fl/fl enteroids (Figure 11E).

We next induced acute DSS-induced colitis in Dr3ΔIEC mice. During acute colitis, we observed a more severe stool phenotype in Dr3ΔIEC mice characterized by bloody diarrhea similar to the phenotype we observed in Dr3-/- mice (Figure 12A). We also observed more severe intestinal inflammation and a significantly higher histology score that was driven mainly by a significantly higher regeneration subscore, indicative of impaired IEC regeneration (Figure 12B). Consistent with our observations in Dr3-/- mice, we observed splenomegaly and an increased number of splenocytes in Dr3ΔIEC mice (Figure 12C), suggesting systemic inflammation. Next, we assessed the degree of bacterial translocation from the LP to the draining lymph nodes by culturing MLN homogenates under anaerobic conditions. We observed a significant increase in the bacterial colonies in MLN homogenates from Dr3ΔIEC mice compared with Dr3fl/fl mice (Figure 12D). Consistent with our data in Dr3-/- mice, the number of BrdU+ cells was similar in Dr3ΔIEC compared with Dr3fl/fl under inflammatory conditions (Figure 12E and F). Although Dr3fl/fl mice responded to acute injury by increased proliferation, in Dr3ΔIEC mice the number of BrdU+ cells was not increased over homeostatic proliferation, suggesting a blunted compensatory proliferation in response to injury in Dr3ΔIEC mice (Figure 12G). Next, we determined the LP immune cell compositions after acute DSS injury in Dr3fl/fl and Dr3ΔIEC mice. We did not observe any significant differences in LP total cell numbers; CD4+ T-cell numbers; secretion of IFN-gamma, IL17A, or IL22 from LPMCs or ileal explants; percentages of IFN-gamma+, IL17A+, IL10+ CD4+ T cells; or IL22+ RORγt+ ILC3s after acute DSS injury (Figure 13). We did observe a significant increase in LP IL22+ CD4+ T cells in Dr3ΔIEC mice (Figure 13D). Collectively, these results show a key role for DR3 signaling on IECs in regulating homeostatic IEC proliferation and barrier function and protecting from bacterial dissemination and promoting tissue repair after intestinal injury.

Our data highlight the role of the IBD-associated genes TNFSF15 (TL1A) and its receptor TNFRSF25 (DR3) as major regulators of intestinal epithelial barrier function. Previous publications that focused on the role of mucosal TL1A–DR3 signaling in innate and adaptive immune responses reported conflicting results, suggesting a complex, dual role of this pathway under homeostatic and inflammatory conditions. Although neutralizing TL1A antibodies prevented and reversed acute and chronic intestinal inflammation,, Tl1a or Dr3 deficiency exacerbated acute DSS-induced colitis., Overexpression of TL1A in transgenic mice causes spontaneous ileitis and fibrosis, thereby mimicking the phenotype of CD patients with TNFSF15 risk polymorphisms.,, The role of TL1A and DR3 in these models has been attributed mainly to its function on different TH subsets, regulatory T cells, and particularly ILCs.17, 18, 19, 20 In this study, we investigated the contribution of TL1A–DR3 signaling in epithelial cells to intestinal pathologies by generating cell type–specific genetic deletion of DR3 on IECs. Our results help to reconcile some of the conflicting results observed in these models by showing an important role for DR3 in intestinal epithelial barrier maintenance under homeostatic conditions and repair after injury. Although neutralizing anti-TL1A antibodies transiently lowers pathophysiological TL1A concentrations in mucosal tissues to physiological concentrations and thereby has therapeutic effects, a complete lack of either TL1A or DR3 has detrimental effects in DSS colitis.,, It is plausible that a complete lack of TL1A or DR3 in Tl1a-/- or Dr3-/- mice has an impact on intestinal homeostasis that renders these mice more susceptible to colitis. Our data support this hypothesis by showing that deficiency of DR3 on IECs plays an essential role in impaired barrier function under homeostatic conditions and impaired IEC repair after tissue injury. Although we observed a reduced frequency of Tregs in Dr3-/- mice that was consistent with a previous report using the acute DSS-induced colitis model and with an important function of DR3 on Treg expansion and function,, we did not observe any increase in T-cell–derived proinflammatory cytokines or significantly increased inflammatory infiltrates into the LP. Although we did not investigate the immunosuppressive properties of Tregs from Dr3-/- mice, a previous publication failed to detect functional impairment of Tregs from Dr3-/- mice in an adoptive transfer model. Our data suggest that immune responses might play a secondary role in the exacerbation of DSS-induced colitis in Dr3-/- mice and that impaired IEC regeneration leads to the severe stool phenotype we observed. Recent studies have reported an indirect effect of the TL1A/DR3 pathway on IEC regeneration by inducing IL22 by ILC3s and T-helper cells., IL22 has been shown to have a profound effect on mucosal healing by inducing the production of antimicrobial peptides and the acute phase protein serum amyloid A.38, 39, 40 Here, we report a direct and cell-intrinsic effect of DR3 expressed by IECs on maintaining intestinal barrier integrity under homeostatic conditions and promoting mucosal healing after injury. Using a highly specific and sensitive probe for Dr3, we detected Dr3 mRNA expression on murine IECs by sm-FISH. Moreover, we also observed Dr3 expression on human IECs in surgical mucosal sections of CD patients. Mice with a deletion of DR3 on IECs (Dr3ΔIEC) showed increased intestinal permeability and uncontrolled proliferation of IECs under homeostatic conditions that also was observed in Dr3-/- mice. Furthermore, we observed increased expression and altered cellular location of several tight junction proteins in these mice. Claudin-1 regulates IEC proliferation and its overexpression worsens DSS-induced colitis and impairs epithelial regeneration. Our findings of dysregulated claudin-1 expression support the idea that Dr3ΔIEC mice are more susceptible to DSS-induced colitis by modulating expression and cellular localization of tight junction proteins. Low-dose TL1A has been shown to induce MLCK2 mRNA and MLCK protein phosphorylation to regulate tight junctions in the human cancer coli-2 cell line via the phosphoinositide 3-kinases/serine-threonine kinase 1 signaling pathway. Further studies are warranted to determine the signaling pathways downstream of DR3 in IECs leading to alteration in tight junction protein expression and localization. Although DR3 signaling has been reported to activate caspase 8 and induce apoptosis in certain cell types, we did not observe any differences in IEC cell death in Dr3ΔIEC mice. These data suggest that IEC cell death does not contribute to the compromised barrier function in Dr3ΔIEC mice under homeostatic conditions. Epithelial regeneration and wound repair are dependent on the self-renewal, proliferation, and differentiation of intestinal stem cells. We observed significant growth defects, impairment of crypt formation, and compromised ZO-1 staining in Dr3ΔIEC enteroids, highlighting the novel role of DR3 in the regulation of intestinal stem cell regenerative potential during homeostasis. We confirmed a direct effect of TL1A–DR3 signaling on enteroids and observed that TL1A stimulation leads to significant growth defects of WT enteroids. Our ex vivo data of impaired organoid formation of Dr3ΔIEC enteroids and our observation that TL1A treatment of WT enteroids results in decreased enteroid surface area and crypts/enteroids are consistent with our in vivo data of impaired induction of IEC regeneration in Dr3ΔIEC mice after DSS-induced injury and indicate impaired proliferation and differentiation in the absence of DR3 signaling in IECs. Although TL1A and IL22 have opposing effects on enteroid growth, our data suggest that IL22 prevents the inhibitory effects TL1A has on organoid formation and growth. Intestinal microbiota contributes significantly to the inflammation seen in DSS-induced colitis. To exclude an impact of differences in intestinal microbiota on the susceptibility and severity of DSS colitis in our mice, we co-housed all experimental mice before and during DSS colitis. Nevertheless, we observed significant translocation of intestinal bacteria to MLNs in Dr3ΔIEC mice that was not observed in WT mice and is consistent with the systemic inflammation that we observed in Dr3ΔIEC mice. To determine if the increase in homeostatic IEC proliferation in Dr3-/- or Dr3ΔIEC mice was microbiota-dependent, we treated Dr3-/- or Dr3ΔIEC mice with broad-spectrum antibiotics and assessed BrdU incorporation. We did not observe any significant differences in the numbers of proliferating IECs in the ileum or colon of antibiotic-treated Dr3-/- or Dr3ΔIEC mice compared with water controls, suggesting that the differences in homeostatic IEC proliferation in Dr3-/- or Dr3ΔIEC mice are microbiome-independent. Our data show a discrepancy of the response to acute DSS between Dr3-/- and Tl1a-/- mice that has been reported previously in these mice. Although Dr3-/- mice present with impaired epithelial barrier integrity, altered expression and localization of tight junction proteins, and increased homeostatic proliferation, Tl1a-/- mice are similar to WT mice. One possible explanation that has been suggested is the presence of additional DR3 ligands, and some potential ligands have been identified in in vitro assays and in vivo models.43, 44, 45 However, their contribution to acute intestinal inflammation remains elusive. Nevertheless, the similar phenotypes of complete Dr3-/- mice and Dr3ΔIEC mice in regard to their IEC function suggests a key role of DR3 on IECs in maintaining homeostatic barrier function and mucosal healing after injury. Further studies using cell-type–specific deletions of TL1A are warranted to elucidate the complex functional network of the TL1A–DR3 pathway, including potential pleiotropic effects of TL1A on different cell types. One limitation of our study was the use of the acute DSS model, which is widely used in the field as an epithelial injury/repair model, but it does not reflect all aspects of human IBD or, more specifically, human ulcerative colitis. However, this model is useful in elucidating mechanisms of IEC regeneration or failure thereof as seen in ulcerative colitis. Our data showed that DR3 is expressed in mouse and human IECs. In addition to the known IBD risk loci in TL1A (TNFSF15), DR3 (TNFRSF25) has been identified recently as an IBD risk loci in a meta-analysis, supporting the importance of our findings. In conclusion, our findings uncovered a novel molecular mechanism underlying the intestinal pathology mediated through DR3 signaling specifically in IECs. These findings provide new insights into the mechanism of the TL1A–DR3 pathway in the pathogenesis of IBD and suggest that therapeutic approaches targeting TL1A rather than DR3 could provide anti-inflammatory therapy while preserving the repair mechanisms downstream of DR3 in IECs.

The following primary antibodies were used: anti–E-cadherin (1:100, cat. Asp155-Ile707; R&D Systems, Minneapolis, MN), anti-BrdU (1:100, cat. BU1/75 [ICR1]; NOVUS Biologicals, Centennial, CO), anti–ZO-1 (1:100, cat. 61-7300; Thermo Fisher Scientific, Waltham, MA), anti–claudin-1 (1:200, cat. ab15098; Abcam, Waltham, MA), anti–claudin-2 (1:200, cat. ab53032; Abcam), and anti–occludin-1 (1:50, cat. 40-4700; Thermo Fisher Scientific). The following secondary antibodies were used: anti-goat (1:500, A11058; Life Technology, Carlsbad, CA), anti-rat (1:500, cat. ab150160; Abcam), and anti-rabbit (1:500, cat. A-11037; Thermo Fisher Scientific). 4’,6-diamidino-2-phenylindole (cat. 0100-20; Southern Biotech, Birmingham, AL) was used for counterstaining for immunofluorescence and FISH. Fluorophore-conjugated antibodies against CD4 (GK1.5, RM4-5), CD45 (30-F11), CD90.2 (53-2.1), CD127 (A7R34), IFN-gamma (XMG1.2), IL17A (TC11-18H10.1), and corresponding isotype controls were from BioLegend (San Diego, CA), and antibodies against FoxP3 (FJK-16s), IL22 (1H8PWSR), RORγt (B2D), Natural Killer Cell P46-Related Protein (29A1.4), hematopoietic lineage cocktail (88-7772-72), IL10 (JES5-16E3), and corresponding isotype controls were from Thermo Fisher Scientific.

Tl1a-deficient (Tl1a-/-), Dr3-deficient (Dr3-/-), and mice with a floxed Dr3 allele (Dr3fl/fl) on a C57BL/6J background were generated as previously described. Villin-Cre mice were purchased from Jackson Laboratory and crossed with Dr3fl/fl to generate Dr3ΔIEC mice. All mice were maintained under specific pathogen-free conditions and handled according to the guidelines and approved protocols of the Cedars-Sinai Medical Center Animal Care and Use Committee (protocol 7430, 8793). Littermates were co-housed throughout the length of the experiments. Female age-matched mice were used for the DSS-induced colitis model, intestinal permeability assay, and assessment of the expression of tight junction genes/proteins.

Female 8- to 12-week-old mice were used for DSS-induced colitis. DSS (40,000–50,000 molecular weight) (cat. 160110; MP Biomedicals, Irvine, CA) was added to the drinking water at a concentration of 2.5% and given ad libitum on days 1–5, 8–12, 15–19, and 22–26 for the induction of chronic colitis, and 3% on days 1–5 for the induction of acute colitis. Body weights and disease activity index were monitored 3 times a week. The disease activity index was scored and calculated as described using freshly collected stool to determine the stool scores. In brief, the combined stool scores were calculated by adding the stool consistency score (0, firm dry stool; 1, moist stool; 2, soft adherent stool; 3, large soft pliable stool; and 4, liquid stool) and the stool blood score (0, no color; 1, flecks of blue; 2, up to 50% blue; 3, >50% blue; and 4, gross red blood). The amount of blood in the stool was quantified using Hemoccult Sensa tests (cat. 64151; Beckman Coulter, Brea, CA). Mice were killed on day 29 for the chronic model and on day 8 for the acute model.

Mice received antibiotic treatment from the time of weaning until 8 weeks of age. A combination of 4 antibiotics (1 g/L ampicillin, 500 mg/L vancomycin, 1 g/L neomycin sulfate, and 1 g/L metronidazole) was added to the drinking water and changed once per week.

Intestinal tissues were fixed in 10% neutral buffered formalin, embedded in paraffin, and stained with H&E for histologic analysis. Histopathologic scores were assigned in a blinded manner by 2 trained animal pathologists as previously described and the average of both scores was reported., In brief, the chronic DSS model was scored as follows: inflammation: 0, none; 1, mild; 2, moderate; and 3, severe; extent: 0, none; 1, mucosa; 2, mucosa and submucosa; and 3, transmural; regeneration: 0, complete regeneration; 1, almost complete regeneration; 2, regeneration with crypt depletion; 3, surface epithelium not intact; and 4, no tissue repair; crypt damage: 0, none; 1, basal one-third damaged; 2, basal two-thirds damaged; 3, only surface epithelium intact; and 4, entire crypt and epithelium lost; and percentage involvement: 1, 1% to 25%; 2, 26% to 50%; 3, 51% to 75%; and 4, 76% to 100%. The total histology score is given as inflammation + extent + regeneration + crypt damage + percentage involvement. Acute DSS colitis was scored as described.

Colons were excised swiftly and opened longitudinally, followed by washing with PBS. Epithelial cells were scraped off with a glass slide and collected for RNA isolation or protein extraction. LPMCs were isolated as previously described.

Cells were restimulated with 50 ng/mL phorbol 12-myristate 13-acetate, 500 ng/mL ionomycin, and Monensin (cat. 00-4505; Thermo Fisher Scientific) for 4 hours, stained with live/dead dye (Thermo Fisher Scientific) and anti-CD4, fixed and permeabilized using the FoxP3 staining buffer set (Thermo Fisher Scientific), and stained with antibodies against murine IL17A, IFN-gamma, IL10, IL22, and FoxP3. For ILC3 staining, cells were stimulated with IL23 (40 ng/mL), IL1β (100 ng/mL), and Brefeldin A for 4 hours, and then stained with hematopoietic lineage cocktail and antibodies against murine Natural Killer Cell P46-Related Protein, IL22, RORγt, CD45, CD90.2, and CD127. The following gating strategy was used to analyze ILC3: immune cells were gated based on forward and side scatter, excluding cell aggregates. Then, ILC3 was identified as CD45+ Lin- CD127+ CD90.2High Rorγt+ IL22+. Samples were acquired using a LSRII flow cytometer (BD Biosciences, Franklin Lakes, NJ) or the Sony ID7000 Spectral Cell Analyzer and analyzed using FlowJo software (TreeStar, Inc, Ashland, OR).

Blood was collected by retro-orbital bleeding and serum was stored at -80ºC until use. Serum IgA (cat. 88-50450) and IgG (cat. 88-50400, both Thermo Fisher Scientific) were assayed by enzyme-linked immunosorbent assay according to the manufacturer’s instructions.

Fecal pellets were collected from 8-week-old Dr3fl/fl and Dr3ΔIEC mice and reconstituted in 1 mL phosphate-buffered saline (PBS) containing 1% bovine serum albumin (BSA) (w/v) overnight at 4ºC. Samples were processed as described. In brief, cell-free supernatants of fecal matter were used for IgA enzyme-linked immunosorbent assay. Secreted IgA concentrations were normalized to the total weight of fecal pellets.

Eight-week-old female mice were fasted overnight and orally gavaged with FITC-dextran (molecular weight, 3000–5000 daltons, Sigma; 500 mg/kg body weight). Blood was collected by retro-orbital bleeding 1 hour after gavage and serum was obtained and stored at -80ºC until use. The serum FITC-dextran concentration was measured with a fluorescence spectrophotometer (SpectraMax; Molecular Devices) at an excitation/emission of 485 nm/520 nm and analyzed using a standard curve of FITC dextran diluted in PBS. Mouse serum from untreated mice was used as a background control.

The enteroid culture was performed using the ileal crypts isolated from Dr3fl/fl and Dr3ΔIEC mice as described previously. Briefly, the ileum was collected, washed in ice-cold Dulbecco's phosphate-buffered saline (DPBS) without Ca2+ and Mg2+, cut open vertically, and luminal content was washed out with ice-cold DPBS. Tissue pieces (2–3 mm) were washed in DPBS and incubated for 20 minutes at room temperature in DPBS/2 mmol/L EDTA with agitation. After the intestinal pieces settled down, DPBS/EDTA buffer was replaced with DPBS/0.1% BSA, gently vortexed for 30–60 seconds, and filtered through a 70-μm cell strainer to remove the tissue chunks. This step was repeated 2 more times and a total of 3 fractions were collected. Fractions 2 and 3 were pooled, centrifuged at 290 × g for 5 minutes, 4ºC, and washed with DPBS/0.1% BSA by centrifuging at 200 × g for 3 minutes, 4ºC. Crypts then were resuspended in Advanced Dulbecco’s modified Eagle medium/F12 (12034; Gibco). Epidermal growth factor (EGF)/Noggin/R-spondin-1 (ENR) media (Advanced Dulbecco’s modified Eagle medium/F12, 10 mmol/L HEPES, 1× GlutMAX supplement, 100 U/mL penicillin/streptomycin, 1 mmol/L N-acetylcysteine (A9165; Sigma Aldrich), 1 × B-27 supplement (17504; Life Technologies), 1 × N-2 supplement (17502; Thermo Fisher Scientific), 50 ng/mL recombinant human epidermal growth factor (AF10015), 100 ng/mL recombinant murine noggin (20538), 1 μg/mL recombinant human R-spondin-1 (12038; all Peprotech, Rocky Hill, NJ)), and Matrigel 1:1 (356231; Corning, Corning, NY) were added to the crypts at a seeding density of 100 crypts per 20 μL. Cell suspension (20 μL) was added to a single well of a 48-well plate (pre-incubated for 30 minutes at 37ºC) and incubated for 10 minutes at 37ºC for polymerization and dome formation. After polymerization of Matrigel, 250 μL ENR media was added to each well. The medium was replaced every 2–3 days with fresh media. Images were acquired using a Zeiss Axio Observer.Z1 inverted microscope at 10× magnification on days 1, 3, and 6. On day 1, enteroids were treated with either TL1A (1896; R&D Systems) or IL22 (Peprotech). RNA from organoids was extracted by adding RNA lysis buffer lysis buffer (Qiagen) directly to the enteroids on day 6 of culture and following the manufacturer’s instructions. The day 6 images were used for the quantification of enteroid surface area and counting the buds/crypts. For enteroid formation potential, the live, intact, and viable enteroids were counted on day 6 and then normalized to the counted enteroids on day 1 of the same well. At least 70–120 enteroids were included in the analysis. The enteroid surface area was calculated using ZEN2 software (Carl Zeiss microscopy).

Intestines were excised from 8-week-old mice and immediately fixed in 10% neutral buffered formalin followed by paraffin-embedding. Tissue sections were deparaffinized, boiled in citrate buffer (10 mmol/L sodium citrate, 0.05% Tween-20; pH 6.0) for antigen retrieval, and incubated with primary antibodies overnight. Fluorophore-conjugated secondary antibodies were incubated for 1 hour. Images were captured with a TCS SP5 X confocal microscope (Leica) and analyzed using LAS AF Lite software (Leica). BrdU-positive cells were counted in a well-oriented crypt–villi axis by an observer blinded to the genotypes. Whole-mount enteroid staining was performed as previously described. Briefly, enteroids were fixed in 4% paraformaldehyde, blocked with PBS/0.1% Triton X-100/0.2% BSA (enteroid wash buffer), and incubated with rabbit anti–ZO-1 (1:100, 61-7300; Thermo Fisher Scientific) and goat anti–E-cadherin (1:200, AF648; R&D Systems) antibodies overnight at 4°C. The enteroids were incubated overnight with donkey anti-goat AF 594 (1:500, A11058; Life Technology) and donkey anti-rabbit Dylight 650 (1:500, ab96922; Abcam), followed by nuclei staining with Hoechst 33342 dye (62249; Thermo Fisher Scientific). The enteroids were mounted on glass slides using fructose–glycerol solution. Images were acquired using a Leica Stellaris 8 confocal microscope and analyzed using ImageJ software (Fiji).

For cell death detection, the TUNEL assay was performed using the Click-iT Plus TUNEL assay kit according to the manufacturer’s instructions (C10618; Thermo Fisher Scientific). Fixed ileal and colonic tissues from Dr3fl/fl and Dr3ΔIEC mice were used.

Tissues were prepared as described for immunofluorescence staining. The DR3 transcripts were detected using the RNAscope Multiplex Fluorescent Reagent Kit v2 (ACD, Newark, CA) according to the manufacturer’s instruction followed by immunofluorescence staining for E-cadherin. The probe for murine Dr3 transcripts was custom designed to hybridize between exons 2 and 5 of murine Dr3, which was deleted in the process of generating Dr3-/- mice. Probes for Peptidylprolyl Isomerase B (Cyclophilin B) and Hela cell pellets were used as quality controls in this assay and a probe for Bacillus subtilis dihydrodipicolinate reductase was used as a negative control. The probe for human DR3 was purchased from Advanced Cell Diagnostics (cat. 448571). Deidentified formalin-fixed, paraffin-embedded tissue taken from the unaffected margin of small-bowel tissue resected during small-bowel resection for complicated CD was used. Tissue samples were obtained by the Material and Information Resources for Inflammatory and Digestive Diseases IBD Biobank after the patients’ informed consent and approval by the Institutional Review Board of the Cedars-Sinai Medical Center (protocol #3358).

RNA was extracted from isolated epithelial cells using RNeasy Mini Kit (Qiagen, Germantown, MD) and complementary DNA was synthesized using the Omniscript RT kit (Qiagen). Complementary DNA was amplified with the QuantiTect SYBR Green Polymerase Chain Reaction Kit (Qiagen) and specific primers using Eppendorf MasterCycler RealPlex (Eppendorf, Enfield, CT). Fecal samples were collected from 8-week-old mice, snap-frozen, and stored at -80°C. DNA was isolated using the DNeasy PowerSoil DNA Isolation Kit (Qiagen) following the manufacturer’s instructions. The abundance of SFB was measured by quantitative polymerase chain reaction using 25 ng fecal bacterial DNA and specific 16S ribosomal RNA primers for SFB. The relative abundance for SFB was quantified by normalizing the quantity of SFB 16S ribosomal RNA gene to the total amount of 16S bacterial DNA. Sequences of primers used in this study were as follows: Gapdh sense: 5’-ACAGTCCATGCCATCACTGCC-3’, antisense: 5’-GCCTGCTTCACCACCTTCTTG-3’, Atcb sense: 5’-GACGGCCAGGTCATCACTATT-3’, antisense: 5’-AGGAAGGCTGGAAAAGAGCC-3’, claudin-1 sense: 5'-CTGGAAGATGATGAGGTGCAGAAGA-3, antisense: 5’-CCACTAATGTCGCCAGACCTGAA-3’, claudin-2: 5’-GGCTGTTAGGCACATCCAT-3’, antisense: 5’-TGGCACCAACATAGGAACTC-3’, claudin-3 sense: 5’-AAGCCGAATGGACAAAGAA-3’, antisense: 5’-CTGGCAAGTAGCTGCAGTG-3’, claudin-4 sense: 5’-CGCTACTCTTGCCATTACG-3’, antisense: 5’-ACTCAGCACACCATGACTTG-3’, claudin-15 sense: 5’-CCACCAGGGCTGGGCTT-3’, antisense: 5’-TCCTGGAGACAGTGGGACAA-3’, Mlck sense: 5’-GGGAAGGCATCACTGAGGTTT-3’, antisense: 5’-GCTCTCAGCAGGCACAGGTGA-3’, Tnfsf15 sense 5’-CAGCAGAAGGATGGCAGAGG-3’, antisense: 5’-CTCTGGCCTGTGTCTACAGC-3’, Tnfrsf25 sense: 5’-GCGTTCTTTGGGGCTATC-3’, antisense: 5’-TCCCAGTACTGCTTGGAGGT-3’, 16S ribosomal RNA universal primer sense: 5’-ACTCCTACGGGAGGCAGCAGT-3’, antisense: 5’-ATTACCGCGGCTGCTGGC-3’, and SFB sense: 5'-GACGCTGAGGCATGAGAGCAT-3', antisense: 5'-GACGGCACGGATTGTTATTCA-3'. Expression levels for each gene were normalized to that of the housekeeping genes Gapdh or Actb and calculated using the 2-ΔΔCT method.

Isolated epithelial cells were washed with DPBS containing sodium orthovanadate and lysed in 50 mmol/L HEPES, 250 mmol/L NaCl, 20 mmol/L β-glycerophosphate, 1 mmol/L sodium orthovanadate, 1 mmol/L EDTA, 1% NP-40, and protease inhibitors (Calbiochem). Lysates were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis through 14% Tris-glycine gels (Invitrogen), transferred to polyvinylidene difluoride membranes, blocked (Odyssey Blocking Buffer, cat. 927-70001; Li-COR Biosciences), and incubated with anti–claudin-1 (1:150), anti–claudin-2 (1:1000), or anti–glyceraldehyde-3-phosphate dehydrogenase (1:2500, ab9485; Abcam) overnight at 4ºC followed by incubation with IRDye goat anti-rabbit antibody (1:5000, cat. 926-32211; Li-COR Biosciences). Images were visualized with the Odyssey Infrared Imaging System (Li-COR Biosciences). Expression levels for each protein were analyzed using ImageJ software (National Institutes of Health, Bethesda, MD) and normalized to that of glyceraldehyde-3-phosphate dehydrogenase.

MLNs were homogenized in 500 μL prereduced sterile PBS using a disposable pellet pestle. A total of 100 μL homogenate was plated on prereduced chocolate blood agar. Growth was recorded 1 week out. Tissue processing and cultivation were performed anaerobically.

Data were analyzed using GraphPad Prism software (GraphPad Software, La Jolla, CA) or JMP pro13 (JMP Statistical Discovery) and presented as the means ± SEM. Differences between the 2 groups were analyzed using the unpaired, 2-tailed Student t test or the Mann–Whitney U test. Analysis of variance with the Tukey honestly significant difference test was used to analyze statistical significance for more than 2 groups. Chi-squared analysis was used to analyze bacterial translocation in Dr3fl/fl and Dr3ΔIEC mice. P values less than .05 were considered statistically significant. The graphical abstract was created with BioRender. All authors had access to the study data and reviewed and approved the final manuscript.