Gliadin induces an increase in intestinal permeability and zonulin release by binding to the chemokine receptor CXCR3

Abstract

Background & aims: Celiac disease is an immune-mediated enteropathy triggered by gliadin, a component of the grain protein gluten. Gliadin induces an MyD88-dependent zonulin release that leads to increased intestinal permeability, a postulated early element in the pathogenesis of celiac disease. We aimed to establish the molecular basis of gliadin interaction with intestinal mucosa leading to intestinal barrier impairment.

Methods: Alpha-gliadin affinity column was loaded with intestinal mucosal membrane lysates to identify the putative gliadin-binding moiety. In vitro experiments with chemokine receptor CXCR3 transfectants were performed to confirm binding of gliadin and/or 26 overlapping 20mer alpha-gliadin synthetic peptides to the receptor. CXCR3 protein and gene expression were studied in intestinal epithelial cell lines and human biopsy specimens. Gliadin-CXCR3 interaction was further analyzed by immunofluorescence microscopy, laser capture microscopy, real-time reverse-transcription polymerase chain reaction, and immunoprecipitation/Western blot analysis. Ex vivo experiments were performed using C57BL/6 wild-type and CXCR3(-/-) mouse small intestines to measure intestinal permeability and zonulin release.

Results: Affinity column and colocalization experiments showed that gliadin binds to CXCR3 and that at least 2 alpha-gliadin 20mer synthetic peptides are involved in this binding. CXCR3 is expressed in mouse and human intestinal epithelia and lamina propria. Mucosal CXCR3 expression was elevated in active celiac disease but returned to baseline levels following implementation of a gluten-free diet. Gliadin induced physical association between CXCR3 and MyD88 in enterocytes. Gliadin increased zonulin release and intestinal permeability in wild-type but not CXCR3(-/-) mouse small intestine.

Conclusions: Gliadin binds to CXCR3 and leads to MyD88-dependent zonulin release and increased intestinal permeability.

Figure 1. Colocalization of CXCR3 and PT-gliadin in CXCR3-transfected HEK293T cells

(A–C) Representative photomicrographs of PT-gliadin-stimulated CXCR3-expressing HEK293T cells. In CXCR3-transfected cells, CXCR3 (B) colocalized with gliadin (A) as indicated by a yellow appearance in the merged picture (C). A second cell expressing CXCR3 only in its upper pole showed gliadin binding only where CXCR3 was present. For details on immunofluorescence staining protocol, see Materials and Methods section. (D–F) Merged pictures of PT-gliadin-treated pcDNA vector-transfected cells stained with anti-gliadin and anti-CXCR3 specific Ab (D), gliadin-treated CXCR3-transfected cells stained with secondary anti-rabbit IgG-FITC and anti-CXCR3 specific Ab (E) and (F) BSA-treated CXCR3-transfected cells stained with anti-BSA and anti-CXCR3 specific Ab. None of the control stainings showed colocalization. Nuclei are in DAPI (blue). Original magnification, × 100. Photomicrographs are representative of 3 experiments.

Figure 2. Dose-response binding curve of PT-gliadin. Concentration-response curve of PT-gliadin binding to CXCR3

PT-gliadin caused a concentration-dependent displacement of ^[125]I-TAC binding from CXCR3, with 50% of ligand displacement obtained with PT-gliadin concentration of 1 mg/mL. The solid boxes represent the percentage of ^[125]I-TAC bound to CXCR3 at the indicated concentrations of PT-gliadin.

Figure 3. CXCR3 is expressed on intestinal epithelial cell lines

Isotype-matched control stainings in panels A and C, respectively. Basal CXCR3 (red) expression was detected on human CaCo-2 (B) and rat IEC6 cells (D). Nuclei are in DAPI (blue). Original magnification, × 100 (n = 2). Basal CXCR3 mRNA expression in CaCo-2 cells (1 ± 0.13) (n = 4) is shown in comparison with CXCR3 expression in HEK293T transfectants. No detectable (ND) CXCR3 mRNA was found in pcDNA-transfected cells. Expression in CXCR3-transfected cells was almost 6-fold higher (5.39 ± 0.68) (n = 9).

Figure 4. CXCR3 expression in mouse intestinal tissues

Laser capture microdissection followed by realtime RT-PCR revealed that CXCR3 mRNA was detectable in the epithelium. As expected, strong CXCR3 expression was detected in the lamina propria where immune cells are localized. Horizontal bars in the graphs indicate median values (n = 5).

Figure 5. Differential mucosal CXCR3 expression in nonceliac and CD patients

In normal subjects in whom the number of intraepithelial lymphocytes is limited (A), CXCR3 is expressed both by immune cells and enterocytes (B). Incubation of the tissue with secondary antibodies alone (C) was performed to show specificity of the staining. Compared with controls (D), CXCR3 expression in CD subjects is highly increased both at the epithelial level and in the lamina propria (E). The increase in intraepithelial lymphocytes typical of CD (F) clearly cannot entirely account for the diffuse over expression of CXCR3. Original magnification, × 60. (G) In active CD, CXCR3 gene expression as determined by real-time PCR was elevated significantly compared to non-CD patients (*P = .004), and this expression returned to baseline after the implementation of a gluten-free diet (n = 3–9).

Figure 6. The increased intestinal permeability and zonulin release in response to PT-gliadin challenge is CXCR3-dependent

(A) Apical application of PT-gliadin increased intestinal permeability in wild-type mice but not in CXCR3^−/− mice. In wild-type mice, a significant decrease in TEER (a measure of increased intestinal permeability) was observed after 90 minutes of PT-gliadin exposure (*P = .001) and decreased further over time (t = 120 minutes, *P = .001) (n = 9). (B) Wild-type C57BL/6 mice responded to PT-gliadin challenge by an increase in the zonulin release that was significant after 30 minutes (solid bar) as compared with baseline values (open bars) (*P < .05). Conversely, zonulin release was unchanged after PT-gliadin challenge in CXCR3^−/− mice. Baseline and post-PT-gliadin exposure TEER values are shown in open and solid bars, respectively (n = 9). (C) Zonulin pathway was intact in CXCR3^−/− mice. Apical challenge of CXCR3^−/− intestinal segments with AT1002, a synthetic peptide derived from Vibrio cholerae protein Zot (the eukaryotic analog of zonulin), induced a decrease in TEER that became significant after 90 minutes (*P = .001) and continued to decrease up to 120 minutes (*P < .001) (n = 7). (D) Four peptides from the α-gliadin synthetic peptide library were applied to the luminal side of wild-type C57BL/6 intestinal segments. A significant decrease in TEER was observed 90 minutes after challenge with the 2 CXCR3-binding peptides A (*P = .05) and B (*P = .02), whereas the nonbinding peptides C and D failed to do so. Baseline and post-PT-gliadin exposure TEER values are shown in open and solid bars, respectively (n = 4–8). (E) Pretreatment of wild-type intestinal segments with IP-10/CXCL10 partially induced tachyphylaxis. In these tissues, PT-gliadin induced a decrease in TEER comparable with tissue that was not pretreated, but the onset was delayed 30 minutes. P values IP-10/gliadin, P = .08 (t = 60 minutes); P = .02 (t = 90 minutes); gliadin, P = .004 (t = 60 minutes), P = .001 (t = 90 minutes) (n = 4). (F) The PT-gliadin-induced TEER changes required G-protein signaling because pertussis toxin (a G-protein-coupled receptor inhibitor) pretreatment abrogated responsiveness to PT-gliadin challenge. Under control conditions, eg, pretreatment with medium alone (*P = .016) or inactivated pertussis toxin (*P = .018), PT-gliadin challenge reduced TEER (n = 10).

Figure 7. PT-gliadin binding to CXCR3 induces recruitment of MyD88

IEC6 cells were incubated at different concentrations (A) and for various times (B) with PT-gliadin. Immunoprecipitation with anti-CXCR3 mAb was performed, and the blot was probed for MyD88. Equal loading was checked by stripping and reprobing the blot for CXCR3. PT-gliadin induced the association of CXCR3 with MyD88, which was observed best when PT-gliadin was applied at a concentration of 1 mg/mL and after an incubation period of 45 minutes (n = 2–3).