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      Hepatic Injury in Nonalcoholic Steatohepatitis Contributes to Altered Intestinal Permeability

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          Abstract

          BACKGROUND & AIMS

          Emerging data suggest that changes in intestinal permeability and increased gut microbial translocation contribute to the inflammatory pathway involved in nonalcoholic steatohepatitis (NASH) development. Numerous studies have investigated the association between increased intestinal permeability and NASH. Our meta-analysis of this association investigates the underlying mechanism.

          METHODS

          A meta-analysis was performed to compare the rates of increased intestinal permeability in patients with NASH and healthy controls. To further address the underlying mechanism of action, we studied changes in intestinal permeability in a diet-induced (methionine-and-choline-deficient; MCD) murine model of NASH. In vitro studies were also performed to investigate the effect of MCD culture medium at the cellular level on hepatocytes, Kupffer cells, and intestinal epithelial cells.

          RESULTS

          Nonalcoholic fatty liver disease (NAFLD) patients, and in particular those with NASH, are more likely to have increased intestinal permeability compared with healthy controls. We correlate this clinical observation with in vivo data showing mice fed an MCD diet develop intestinal permeability changes after an initial phase of liver injury and tumor necrosis factor- α (TNF α) induction. In vitro studies reveal that MCD medium induces hepatic injury and TNF α production yet has no direct effect on intestinal epithelial cells. Although these data suggest a role for hepatic TNF α in altering intestinal permeability, we found that mice genetically resistant to TNF α-myosin light chain kinase (MLCK)–induced intestinal permeability changes fed an MCD diet still develop increased permeability and liver injury.

          CONCLUSIONS

          Our clinical and experimental results strengthen the association between intestinal permeability increases and NASH and also suggest that an early phase of hepatic injury and inflammation contributes to altered intestinal permeability in a fashion independent of TNF α and MLCK.

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          Most cited references 32

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          Caveolin-1–dependent occludin endocytosis is required for TNF-induced tight junction regulation in vivo

          Introduction Infectious, ischemic, and immune-mediated intestinal diseases are characterized by a loss of epithelial paracellular barrier function (Hollander et al., 1986; Clayburgh et al., 2004; Turner, 2009), which, in the absence of gross epithelial destruction, reflects increased tight junction permeability (Gitter et al., 2001; Suenaert et al., 2002; Epple et al., 2009). Although the extracellular mediators that trigger tight junction regulation are incompletely defined, cytokines contribute to barrier loss by at least two mechanisms. For example, IL-13, which is increased in the mucosa of patients with ulcerative colitis and Crohn’s disease, is able to induce expression of claudin-2 (Prasad et al., 2005; Zeissig et al., 2007). In turn, claudin-2 expression increases tight junction permeability to both ions and small, nonionic solutes (Simon et al., 1999; Furuse et al., 2001; Van Itallie et al., 2001, 2008; Weber et al., 2010). In contrast, TNF, which is critical to Crohn’s disease pathogenesis and contributes significantly to infectious, ischemic, and immune-mediated intestinal diseases, regulates barrier function via myosin light chain (MLC) phosphorylation (Clayburgh et al., 2005; Blair et al., 2006) and tight junction remodeling (Shen et al., 2006). Although in vitro and in vivo studies have shown that acute TNF-induced barrier loss requires MLC kinase (MLCK)–dependent MLC phosphorylation (Zolotarevsky et al., 2002; Clayburgh et al., 2005; Ma et al., 2005; Wang et al., 2005), the only associated ultrastructural modification reported is condensation of the perijunctional actomyosin ring (Clayburgh et al., 2005). Immunofluorescence microscopy demonstrates marked internalization of the transmembrane protein occludin after TNF treatment (Clayburgh et al., 2005). The observations that TNF-induced MLC phosphorylation, occludin internalization, paracellular barrier loss, and diarrhea are all prevented by genetic or pharmacological MLCK inhibition (Clayburgh et al., 2005) suggest that these events are closely linked. Therefore, we sought to define the mechanisms of TNF-induced occludin internalization and to determine whether this endocytic event is required for in vivo barrier loss. In vitro studies have reported occludin endocytosis via macropinocytosis, clathrin-coated pits, and caveolae (Ivanov et al., 2004b; Bruewer et al., 2005; Shen and Turner, 2005; Schwarz et al., 2007). To define the mechanisms of TNF-induced occludin endocytosis in vivo, we developed mice expressing fluorescent occludin and ZO-1 fusion proteins within the intestinal epithelium. These were studied using high resolution in vivo imaging approaches. Our data show that TNF induces focal intrajunctional concentration of occludin followed by caveolin-1–dependent endocytosis. Moreover, both caveolin-1 knockout and pharmacologic inhibition of endocytosis prevented TNF-induced occludin internalization as well as tight junction barrier loss and water secretion. Finally, occludin overexpression limited barrier loss and prevented water secretion. Thus, caveolin-1–dependent occludin endocytosis is essential for in vivo immune-mediated tight junction regulation. Results We have previously shown that diarrhea induced by TNF requires the combined effects of protein kinase Cα, which inhibits Na+ absorption, and MLCK, which increases tight junction permeability (Clayburgh et al., 2005, 2006). The mechanisms by which Na+ absorption is inhibited, thereby reducing the transmucosal Na+ gradient that drives paracellular water absorption, have been studied extensively (Lee-Kwon et al., 2003; Clayburgh et al., 2006). In contrast, the processes by which MLCK activation leads to tight junction regulation are less well characterized. Occludin internalization precedes intestinal fluid secretion To determine whether tight junction protein redistribution was associated with TNF-induced barrier disruption and diarrhea, the localizations of claudin-1, -3, -4, -5, -7, -15, and E-cadherin were examined. These were unaffected by in vivo TNF treatment (Fig. 1 A). Claudin-2 was only detected in crypt epithelium, and claudin-12 was not detected at all, which is consistent with previous data (Holmes et al., 2006). In contrast, a marked increase in the number of occludin-containing cytoplasmic vesicles developed after TNF administration (Fig. 1 B). Although apical–basal-oriented sections (Fig. 1 B) could be interpreted to suggest that occludin synthesis is stimulated by TNF, images orthogonal to the apical–basal orientation showed that this increase in cytoplasmic occludin is associated with a decrease in tight junction–associated occludin (Fig. 1 C). This is confirmed by immunoblots showing that the total occludin content of isolated jejunal enterocytes was unchanged after TNF treatment (Fig. 1 D). Figure 1. Occludin endocytosis begins 90 min after TNF administration and precedes intestinal fluid accumulation. (A) Jejunum was harvested from wild-type mice at the indicated times after intraperitoneal injection of 5 µg TNF and labeled for claudin proteins or E-cadherin (green), ZO-1 or F-actin (red), and nuclei (blue). (B) As in A, jejunal sections were labeled for occludin (green), F-actin (red), and nuclei (blue). (C) Jejunal sections were harvested and labeled as in B. TNF treatment leaves large regions of the tight junction completely lacking in occludin (arrows). (D) Jejunal epithelia were isolated from wild-type mice 120 min after injection with saline or TNF and analyzed via immunoblotting. (E) Number of occludin-containing vesicles (black circles) was assessed morphometrically, and fluid accumulation (white boxes) was measured as weight/length ratio (n = 4). Error bars indicate mean ± SEM. Bars, 10 µm. As an initial assessment of the relationship between occludin internalization, barrier loss, and diarrhea, occludin vesicle number was correlated with intestinal fluid accumulation. Immunofluorescent analysis of jejunum harvested at intervals after intraperitoneal TNF injection identified significant occludin internalization within 90 min (P 90% sequence conservation between human and murine ZO-1; therefore, a validated human ZO-1 cDNA was used (Shen and Turner, 2005; Shen et al., 2008). The villin promoter, fusion protein sequence, and bovine growth hormone polyadenylation sequence were excised by restriction digestion and injected into C57BL/6 embryos by The University of Chicago Transgenic Mouse Core Facility. Mice were screened by PCR of genomic DNA. Live animal imaging Mice were anesthetized, injected intravenously with Hoechst 33342 dye, and the abdomens were opened by a midline incision. Electrocautery was used to open a 2-cm loop of jejunum along the antimesenteric border. The abdominal cavity was closed under the externalized loop of jejunum while taking care to protect the neurovascular supply. The mucosal surface of the jejunum was placed against the coverslip bottom of a 35-mm Petri dish containing 0.15 ml HBSS with the body of the mouse over the jejunum, and both were placed on a 37°C heated microscope stage. Mice were imaged using a multiphoton confocal inverted microscope (SP5; Leica) with a 40× 0.8 NA water immersion objective. EGFP was imaged using an Argon laser and spectral emission range of 490–568 nm, and mRFP1 was imaged using a laser (DPSS 561) with a spectral emission range of 577–679 nm, and a pinhole of 130 µm was used for both proteins. Hoechst dye was imaged using a multiphoton laser with a spectral emission range of 400–492 nm and pinhole of 600 µm. Scanning was performed at 200 Hz, and all image acquisition was controlled by LAS-AF software (version 2.1; Leica). Postacquisition image analysis was performed using MetaMorph (version 7; MDS Analytical Technologies). Immunofluorescence Mouse jejunum was snap frozen in optimal cutting temperature medium and stored at −80°C. 5-µm frozen sections were fixed in 1% paraformaldehyde and immunostained as described previously using primary mouse monoclonal antioccludin (Invitrogen), rabbit anti–caveolin-1 (Abcam), rabbit anticlathrin heavy chain (Santa Cruz Biotechnology, Inc.), rabbit anti-EEA1 (Thermo Fisher Scientific), affinity-purified rabbit anti–phosphorylated MLC (Berglund et al., 2001), mouse anti–claudin-1 (Invitrogen), rabbit anti–claudin-2 (Invitrogen), rabbit anti–claudin-3 (Invitrogen), rabbit anti–claudin-4 (Invitrogen), rabbit anti–claudin-5 (Invitrogen), rabbit anti–claudin-7 (Invitrogen), rabbit anti–claudin-12 (Invitrogen), rabbit anti–claudin-15 (Invitrogen), rat anti–E-cadherin (Invitrogen), or monoclonal rat anti–ZO-1 (Stevenson et al., 1986) primary antibodies followed by Alexa Fluor 488– or 594–conjugated secondary antibodies (Invitrogen) along with Alexa Fluor 488– or 594–conjugated phalloidin (Invitrogen) and Hoechst 33342 (Invitrogen). Rabbit antioccludin (Invitrogen) was used in some experiments. Stained sections were mounted in Prolong gold (Invitrogen) and imaged using an epifluorescence microscope (DM4000; Leica) equipped with DAPI, Endow GFP, and Texas red zero-pixel shift filter sets (Chroma Technology Corp.), a 63× 1.32 NA oil immersion objective, and a camera (CoolSNAP HQ; Roper Industries) controlled by MetaMorph. Z stacks were collected at 0.2-µm intervals and deconvolved using AutoDeblur (version X1; Media Cybernetics) for 10 iterations. For morphometric analysis, deconvolved z stacks were merged after pseudocolor assignment, and vesicles were defined as round or oval structures <2 µm in greatest diameter present in two or more planes. Determination of colocalization required at least 50% overlap of the two signals within at least two adjacent image planes. The number of vesicles was counted over the entire cell volume. 30 representative surface enterocytes were counted for each condition. Electron microscopy and morphometry At indicated times after intraperitoneal saline or TNF injection, mice were sacrificed, and jejunal segments were minced and fixed in 2.5% glutaraldehyde and 4% paraformaldehyde in 0.1 M sodium cacodylate and processed as described previously (Clayburgh et al., 2005). Samples were embedded in SPURR (Electron Microscopy Sciences), and two blocks from each of two mice were processed per time point. At least 15 well-oriented sections including apical cytoplasm of 15–20 villous enterocytes were examined for each mouse. Images were collected using a scanning transmission electron microscope (Tecnai F30; FEI). For morphometry, images taken at 5,900× were imported into MetaMorph, vesicles were counted and measured, and cytoplasmic area was assessed by an observer blinded to experimental condition. At least 1,000 µm2 of apical cytoplasm was examined for each condition. Data were modeled as the sum of four Gaussian equations, where a, b, and c are the amplitudes, position of the center of peak, and width of the vesicle pools, respectively. Analysis with a1 − a4 as the only free parameters was used to determine the amplitude of each pool. Constant values were b1 = 80.5, b2 = 124.8, b3 = 168.6, b4 = 240, c1 = 18.5, c2 = 18.2, c3 = 18.0, and c4 = 8.7. The number of vesicles was calculated by the equation a 1 e ( − ( ( x − b 1 ) 2 / 2 c 1 2 ) + a 2 e ( − ( ( x − b 2 ) 2 / 2 c 2 2 ) + a 3 e ( − ( ( x − b 3 ) 2 / 2 c 3 2 ) + a 4 e ( − ( ( x − b 4 ) 2 / 2 c 4 2 ) . Immunoelectron microscopy 100 min after intraperitoneal saline or TNF injection, mice were anesthetized, and 1–2-mm jejunal segments were transferred to aluminum sample holders, cryoprotected with 150 mM sucrose, and frozen in a high pressure freezer (HPM 010 RMC; BAL-TEC). High pressure–frozen samples were freeze substituted in 0.2% uranyl acetate (Electron Microscopy Sciences) plus 0.25% glutaraldehyde (Electron Microscopy Sciences) in acetone at −80°C for 84 h and warmed to −50°C. After three 15-min acetone washes, the samples were removed from their holders and slowly infiltrated under controlled time and temperature conditions in an AFS system (Leica) at −50°C with resin (Lowicryl HM20; Electron Microscopy Sciences) according to the following schedule: 25, 50, 75, and 100% (12 h at each concentration) followed by three 1-h 100% resin washes. The samples were placed into flat-bottomed molds (Electron Microscopy Sciences) and polymerized at −50°C under UV light for 20 h. 80-nm-thin sections were mounted on Formvar-coated gold grids and single or double immunolabeled using a previously described approach (Otegui et al., 2006). In brief, for single labeling, sections were incubated with 0.1 N HCl for 10 min, blocked with a 5% (wt/vol) solution of nonfat milk in PBS containing 0.1% Tween 20, incubated with 1:5 rabbit antioccludin antibodies directed against the carboxy terminus (Invitrogen) for 3 h or 1:40 rabbit anti–caveolin-1 antiserum (Abcam) for 1 h, washed in a stream of PBS plus 0.5% Tween 20 (PBS-HT), and incubated with 1:20 goat anti–rabbit IgG antiserum conjugated to 10 nm gold particles (Ted Pella, Inc.) for 1 h followed by a wash with PBS-HT. At least 10 grids were examined for each staining condition. Label was only detected at the apical junctional complex and in association with intracellular vesicles (primarily after TNF treatment). No label was detected over the basal cytoplasm, nucleus, mitochondria, or luminal space. For double labeling, sections were incubated with HCl, blocked as described for single labeling, incubated with 1:5 rabbit antioccludin antibodies directed against the carboxy terminus (Invitrogen) for 3 h, washed, and incubated with 1:20 goat anti–rabbit IgG antiserum conjugated to 10 nm gold particles for 1 h. After a wash with PBS-HT, the sections were fixed with 0.1% glutaraldehyde for 10 min, incubated with blocking solution followed by 1:40 rabbit anti–caveolin-1 antiserum (Abcam) for 1 h, washed, and incubated with 1:20 goat anti–rabbit IgG conjugated to 15 nm gold particles (Ted Pella, Inc.) for 1 h. The distribution of each protein was identical in both single- and double-labeling experiments (single-antibody labeling shown in Fig. 7 and Fig. S2). Two control experiments were performed to test the specificity of the individual antibodies when used together, one in which only anti–caveolin-1 was omitted and only antioccludin staining was detected (only 10 nm gold present; Fig. S1 A), and the second in which only antioccludin was omitted and only anti–caveolin-1 staining was detected (only 15 nm gold detected; Fig. S1 B). In each case, no gold particles of the incorrect size, i.e., corresponding to the omitted primary antibody, were detected in at least 10 grids examined per condition. In addition, the identical distributions of occludin and caveolin-1 in single- and double-label experiments strongly argue that the detection observed is specific and not an artifact of antibody aggregation. Epithelial isolation The jejunum was opened lengthwise, washed in 4°C Ca2+- and Mg2+-free HBSS, and epithelial cells were isolated as described previously (Clayburgh et al., 2005). Cells were lysed in Laemmli sample buffer, sonicated, boiled, and separated by SDS-PAGE. After transfer to PVDF, membranes were probed using primary antibodies against phosphorylated MLC or total MLC (Wang et al., 2005), occludin, β-actin (Sigma-Aldrich), caveolin-1, or clathrin heavy chain followed by the appropriate secondary antibodies. Blots were analyzed using an Odyssey system (LI-COR Biosciences) and ImageJ (National Institutes of Health). Fluid accumulation and in vivo permeability assays Fluid accumulation was assessed as jejunal weight/length ratio as described previously (Clayburgh et al., 2005). Paracellular permeability and water transport were measured in vivo (Clayburgh et al., 2005). In brief, 20 min after intraperitoneal injection of 5 µg recombinant murine TNF (PeproTech) or saline, mice were anesthetized and injected intravenously with 250 µl 1 mg/ml Alexa Fluor 488–conjugated BSA (Invitrogen). The abdomen was opened, and a 5-cm loop of jejunum was cannulated and flushed to remove luminal material. The jejunal loop was perfused with test solution (50 mM NaCl, 5 mM Hepes, 2 mM sodium ferrocyanide, 2.5 mM KCl, and 20 mM glucose, pH 7.4) in a recirculating manner at 1 ml/min for 2 h, encompassing the interval from 1.5 to 3.5 h after initial intraperitoneal injection. Inhibitors were included in the perfusion solution where indicated. Ferrocyanide and Alexa Fluor 488–conjugated BSA concentrations in the perfusate were assessed and used to determine paracellular BSA flux and net water transport (Clayburgh et al., 2005). When used, fluorescent-conjugated tracers of endocytosis (Alexa Fluor 594–conjugated WGA [Invitrogen] and DyLight 594–conjugated mouse IgG [Jackson ImmunoResearch Laboratories, Inc.]) were included in an initial 30-min perfusion after which the perfusion solution was replaced with fresh buffer that was identical save for the exclusion of the fluorescent tracer. At least three replicates are reported for each condition. Statistical analysis All data are presented as means ± SEM and represent at least three independent experiments. P-values were determined by Student’s t test and were considered to be significant if P ≤ 0.05. Online supplemental material Fig. S1 shows controls for double-label immunoelectron microscopy. The double-labeling protocol was used as in Figs. 7 and 8, but either antioccludin or anticaveolin was omitted. There was no background signal for the omitted antigen under any of these conditions. Fig. S2 shows single-label immunoelectron microscopy to detect occludin or cavelin-1 in jejunal enterocytes of TNF-treated wild-type mice. Videos 1–4 show in vivo imaging of murine intestinal mucosa. Video 1 shows blood flow within villus capillaries. Videos 2 and 3 show the distribution of EGFP-occludin and mRFP1–ZO-1 transgenically expressed within murine intestinal epithelium. Video 4 shows EGFP-occludin endocytosis in vivo after systemic TNF. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200902153/DC1.
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            Interferon-gamma and tumor necrosis factor-alpha synergize to induce intestinal epithelial barrier dysfunction by up-regulating myosin light chain kinase expression.

            Numerous intestinal diseases are characterized by immune cell activation and compromised epithelial barrier function. We have shown that cytokine treatment of epithelial monolayers increases myosin II regulatory light chain (MLC) phosphorylation and decreases barrier function and that these are both reversed by MLC kinase (MLCK) inhibition. The aim of this study was to determine the mechanisms by which interferon (IFN)-gamma and tumor necrosis factor (TNF)-alpha regulate MLC phosphorylation and disrupt epithelial barrier function. We developed a model in which both cytokines were required for barrier dysfunction. Barrier dysfunction was also induced by TNF-alpha addition to IFN-gamma-primed, but not control, Caco-2 monolayers. TNF-alpha treatment of IFN-gamma-primed monolayers caused increases in both MLCK expression and MLC phosphorylation, suggesting that MLCK is a TNF-alpha-inducible protein. These effects of TNF-alpha were not mediated by nuclear factor-kappaB. However, at doses below those needed for nuclear factor-kappaB inhibition, sulfasalazine was able to prevent TNF-alpha-induced barrier dysfunction, MLCK up-regulation, and MLC phosphorylation. Low-dose sulfasalazine also prevented morphologically evident tight junction disruption induced by TNF-alpha. These data show that IFN-gamma can prime intestinal epithelial monolayers to respond to TNF-alpha by disrupting tight junction morphology and barrier function via MLCK up-regulation and MLC phosphorylation. These TNF-alpha-induced events can be prevented by the clinically relevant drug sulfasalazine.
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              Role of innate immunity and the microbiota in liver fibrosis: crosstalk between the liver and gut.

              Liver fibrosis occurs as a wound-healing scar response following chronic liver inflammation including alcoholic liver disease, non-alcoholic steatohepatitis, viral hepatitis, cholestatic liver disease and autoimmune liver diseases. The liver has a unique vascular system within the gastrointestinal tract, as the majority of the liver's blood supply comes from the intestine through the portal vein. When the intestinal barrier function is disrupted, an increase in intestinal permeability leads to the translocation of intestine-derived bacterial products such as lipopolysaccharide (LPS) and unmethylated CpG containing DNA to the liver via the portal vein. These gut-derived bacterial products stimulate innate immune receptors, namely Toll-like receptors (TLRs), in the liver. TLRs are expressed on Kupffer cells, endothelial cells, dendritic cells, biliary epithelial cells, hepatic stellate cells, and hepatocytes. TLRs activate these cells to contribute to acute and chronic liver diseases. This review summarizes recent studies investigating the role of TLRs, intestinal microbiota and bacterial translocation in liver fibrosis, alcoholic liver disease and non-alcoholic steatohepatitis.
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                Author and article information

                Journal
                101648302
                43543
                Cell Mol Gastroenterol Hepatol
                Cell Mol Gastroenterol Hepatol
                Cellular and molecular gastroenterology and hepatology
                2352-345X
                10 September 2015
                15 January 2015
                March 2015
                22 September 2015
                : 1
                : 2
                : 222-232
                Affiliations
                [1 ]Gastrointestinal Unit, Department of Medicine, Massachusetts General Hospital, Harvard Medical School, Boston, Massachusetts
                [2 ]Division of Gastroenterology, Department of Medicine, Case Western Reserve University, Cleveland, Ohio
                [3 ]Center for Engineering in Medicine, Department of Surgery, Massachusetts General Hospital, and the Shriners Burns Hospital, Boston, Massachusetts
                [4 ]Department of Pathology, University of Chicago, Chicago, Illinois
                Author notes
                Address correspondence to: Suraj J. Patel, MD, PhD, Center for Engineering in Medicine, Massachusetts General Hospital, Shriner’s Burn Hospital, Boston, Massachusetts 02114. sjp27@ 123456alumni.mit.edu ; fax: (617) 573-9471
                [*]

                Authors share corresponding authorship.

                Article
                NIHMS721437
                10.1016/j.jcmgh.2015.01.001
                4578658

                This is an open access article under the CC BY-NC-ND license ( http://creativecommons.org/licenses/by-nc-nd/4.0/).

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