β-catenin regulates innate and adaptive immunity in mouse liver ischemia-reperfusion injury

Authors

  • Bibo Ke,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Xiu-Da Shen,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Naoko Kamo,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Haofeng Ji,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Shi Yue,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Feng Gao,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Ronald W. Busuttil,

    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
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  • Jerzy W. Kupiec-Weglinski

    Corresponding author
    1. Dumont-UCLA Transplant Center, Department of Surgery, Division of Liver and Pancreas Transplantation, David Geffen School of Medicine at UCLA, Los Angeles, CA
    • M.D., Ph.D., Dumont-UCLA Transplant Center 77-120 CHS, 10833 Le Conte Ave., Los Angeles, CA 90095
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    • fax: 310-267-2358


  • Potential conflict of interest: Nothing to report.

Abstract

Dendritic cells (DCs) are critical mediators of immune responses that integrate signals from the innate immune system to orchestrate adaptive host immunity. This study was designed to investigate the role and molecular mechanisms of STAT3-induced β-catenin in the regulation of DC function and inflammatory responses in vitro and in vivo. STAT3 induction in lipopolysaccharide (LPS)-stimulated mouse bone marrow-derived DCs (BMDCs) triggered β-catenin activation by way of GSK-3β phosphorylation. The activation of β-catenin inhibited phosphatase and tensin homolog delete on chromosome 10 (PTEN) and promoted the phosphoinositide 3-kinase (PI3K)/Akt pathway, which in turn down-regulated DC maturation and function. In contrast, knockdown of β-catenin increased PTEN/TLR4 (Toll-like receptor 4), interferon regulatory factor-3 (IRF3), nuclear factor kappa B (NF-κB) activity, and proinflammatory cytokine programs in response to LPS stimulation. In a mouse model of warm liver ischemia and reperfusion injury (IRI), disruption of β-catenin signaling increased the hepatocellular damage, enhanced hepatic DC maturation/function, and PTEN/TLR4 local inflammation in vivo. Conclusion: These findings underscore the role of β-catenin to modulate DC maturation and function at the innate-adaptive interface. Activation of β-catenin triggered PI3K/Akt, which in turn inhibited TLR4-driven inflammatory response in a negative feedback regulatory mechanism. By identifying the molecular pathways by which β-catenin regulates DC function, our findings provide the rationale for novel therapeutic approaches to manage local inflammation and injury in IR-stressed liver. (HEPATOLOGY 2013)

Liver ischemia and reperfusion injury (IRI), a local inflammatory response driven by innate and supported by adaptive immune responses, represents an important cause of organ dysfunction and failure in liver transplantation.1 Our group was one of the first to document the essential function of Toll-like receptor 4 (TLR4) in the mechanism of liver IRI by promoting local inflammation and hepatocellular damage by way of the downstream interferon (IFN) regulatory factor (IRF) 3 pathway.2 It soon became evident that IR-induced liver damage triggers TLR4 endogenous ligands, such as high-mobility group box 1 (HMGB1), to activate dendritic cells (DCs) and facilitate inflammatory cytokine programs that further enhance TLR4-mediated local inflammation.3, 4

Although different cell types (hepatocytes, Kupffer cells, sinusoidal endothelial cells, and infiltrating T cells) contribute to IRI pathophysiology, hepatic DCs are well suited to modulate local immune responses that can bridge innate and adaptive immunity in the liver.5 Indeed, immature DCs in peripheral tissues function to capture and process antigens.5, 6 Upon exposure to pathogens and TLR ligands, however, DC rapidly acquire an activated phenotype and undergo maturation characterized by up-regulated expression of major histocompatibility complex (MHC) antigens, costimulatory CD80/CD86 molecules, and proinflammatory cytokines that stimulate naïve T-cell differentiation.5, 6 Hence, controlling DC differentiation is important to prevent hepatic innate and adaptive inflammatory development.

STAT3 is known to mediate many biological effects by regulating immune homeostasis and influencing cell proliferation/differentiation.7 Disruption of STAT3 during hematopoiesis activates innate immune response and promotes proinflammatory phenotype.8 STAT3 signaling may halt DC maturation in vitro,9 whereas STAT3 deficiency in interleukin (IL)-10−/− DCs was shown to increase nuclear factor kappa B (NF-κB) binding to the IL-12p40 promoter and to promote TLR-dependent IL-12 inflammation.10 As conditional deletion of STAT3 results in severe colitis and enhanced Th1-type activity,11 STAT3 may serve as an intrinsic negative regulator of DC function.12

The Wnt-β-catenin pathway is an important regulator of cell development, regeneration, and carcinogenesis.13, 14 In response to Wnt signaling, β-catenin is rapidly phosphorylated and enters the nucleus, where it interacts with T-cell factor / lymphoid enhancer factor (TCF/LEF) family members to regulate transcription of the target genes. Inhibition of STAT3 induces translocation of β-catenin from the nucleus to the cytoplasm, leading to decreased β-catenin transcription activity,15 suggesting β-catenin function might be mediated by STAT3. Moreover, activation of β-catenin was shown to regulate the local immunity and tolerance balance in murine intestinal mucosa.16 Despite its essential immunomodulatory functions, however, little is known of the molecular mechanisms by which β-catenin may regulate DC function and/or local inflammation responses in the liver.

Here we report on the crucial regulatory function of STAT3-induced β-catenin on DC function and inflammatory responses in hepatic IRI. We demonstrate that β-catenin inhibits phosphatase and tensin homolog delete on chromosome 10 (PTEN) and promotes the PI3K/Akt pathway, which in turn down-regulates DC immune function and depresses TLR4-driven inflammation. Our data document β-catenin as a novel regulator of innate and adaptive immune responses in the mechanism of liver IRI.

Abbreviations

Ad-β-gal, recombinant adenovirus β-galactosidase reporter gene; BMDCs, bone marrow derived-dendritic cells; DC, dendritic cell; GSK-3β, glycogen synthase kinase 3β; HO-1, hemeoxygenase-1; IRF3, interferon regulatory factor-3; LPS, lipopolysaccharide; PI3K, phosphoinositide 3-kinase; PTEN, phosphatase and tensin homolog delete on chromosome 10; sGPT, serum glutamic-pyruvic transaminase; siRNA, small interfering RNA; TLR4, Toll-like receptor 4; TUNEL, terminal deoxyribonucleotidyl transferase (TdT)-mediated dUTP-digoxigenin nick end labeling.

Materials and Methods

Animals.

Male C57BL/6 wildtype (WT) mice at 6-8 weeks of age were used (Jackson Laboratory, Bar Harbor, ME). Animals, housed in UCLA animal facility under specific pathogen-free conditions, received humane care according to the criteria outlined in the “Guide for the Care and Use of Laboratory Animals” (NIH publication 86-23 revised 1985).

Cell Isolations, In Vitro Cultures.

Murine BMDCs and liver DCs were generated as described.17, 18 In brief, bone-marrow cells from femurs of WT mice were cultured in RPMI-1640 supplemented with 10% fetal bovine serum (FBS), 100 μg/mL of penicillin/streptomycin (Life Technologies, Grand Island, NY), in 12-well plates (1 × 106 cells/mL) with granulocyte-macrophage colony-stimulating factor (GM-CSF, 20 ng/mL, R&D Systems, Minneapolis, MN) and IL-4 (10 ng/mL, R&D Systems). Adherent immature DCs (purity ≥90% CD11c+) were recovered for in vitro experiments on day +7.

To separate hepatic DCs, mouse livers perfused with phosphate-buffered saline (PBS) followed by collagenase type IV/DNase 1 (Sigma-Aldrich, St. Louis, MO). After washing, the resuspended cells were incubated with antimouse CD11c-coated immunomagnetic beads (Stemcell Technologies) for 15 minutes at 4°C and positively selected by using a magnetic column according to the manufacturer's instruction. For DC maturation studies, CD11c-enriched cells were cultured for 24 hours with lipopolysaccharide (LPS; 0.5 μg/mL).

Preparation of Small Interfering RNA (siRNA).

siRNA against β-catenin was designed using the siRNA selection program.19 The sense and antisense strands of murine β-catenin siRNA were 5′-AGCUGAUAUUGAUGGACAG-3′ (sense) and 5′-CUGUCCAUCAAUAUCAGCU-3′ (antisense). The nonsilencing (NS)siRNA 5′-UUCUCCGAACGUGUCACGU-3′ (sense) and 5′-ACGUGACACGUUCGGAGAA-3′ (antisense) served as negative controls. The generation of siRNA against STAT3 was described.20 All siRNAs were synthesized in 2′-deprotected, duplexed, desalted and purified siRNA form (Qiagen, Chatsworth, CA).

In Vitro Transfections and Treatment.

On day 7, one ×106 cells/well of immature BMDCs were transfected with 100 nM of siRNA using lipofectamine 2000 reagent (Invitrogen, San Diego, CA) and incubated for 24 hours. Cells were then treated with 10 μg/mL of cobalt protoporphyrin (CoPP; HO-1 inducer) or tin protoporphyrin (SnPP; competitive HO-1 inhibitor) (Porphyrin Products, Logan, UT) or with 50 ng/mL of murine recombinant IL-10 (rIL-10; R&D Systems) and incubated for an additional 6 hours.20

Enzyme-Linked Immunosorbent Assay (ELISA) Assay.

Murine BMDC culture supernatants were harvested for cytokine analysis. ELISA kits were used to measure IL12p40/TNF-α/IL-6 levels (eBiosciences, San Diego, CA).

Flow Cytometry Analysis.

Murine BMDCs were stained with anti-CD11c, CD40, CD80, and CD86 PE-conjugated monoclonal antibodies (mAbs) (eBiosciences). PE-labeled rat anti-IgG2a isotypes were used as negative controls. Measurements were performed using a FACSCalibur flow cytometer (BD Biosciences). Data analysis was performed using CellQuest software.

Malachite Green Phosphate Assay.

Murine BMDC and hepatic DC protein lysates were immunoprecipitated with anti-PTEN Ab and incubated with protein A/G agarose beads. The PTEN malachite green assay was performed with beads-bound PTEN (Echelon Biosciences, Salt Lake City, UT). The released phosphate was determined relative to a phosphatase standard curve.

Mouse Liver IRI Model and Treatment.

We used an established mouse model of warm hepatic ischemia followed by reperfusion.19, 20 Mice were injected with heparin (100 U/kg) and an atraumatic clip was used to interrupt the arterial/portal venous blood supply to the cephalad liver lobes. After 90 minutes of ischemia the clip was removed and mice were sacrificed at 6 hours of reperfusion. Some animals were injected by way of the tail vein with Ad-HO-1, Ad-IL-10, or Ad-β-gal (2.5 × 109 pfu) 24 hours prior to ischemia. β-Catenin siRNA or nonspecific siRNAs (2 mg/kg) was injected intravenously at 4 hours prior to ischemia.19, 20 Consistent with others,21 >40% of intravenously infused siRNA consistently accumulate in the ischemic lobes.19

Hepatocellular Function Assay.

Serum glutamic-pyruvic transaminase (sGPT) levels, an indicator of hepatocellular injury, were measured with an autoanalyzer (Antech Diagnostics, Los Angeles, CA).

Histology, Immunohistochemistry, Double Immunofluorescence Staining.

Liver sections (5 μm) were stained with hematoxylin and eosin (H&E). The severity of IRI was graded using Suzuki's criteria on a scale from 0-4.22 Liver DCs were detected using primary mAb against mouse CD11c (EMD Millipore, Billerica, MA) followed by incubation with secondary Ab, biotinylated goat antihamster IgG (Vector, Burlingame, CA). CD11c/β-catenin double-positive DCs were identified by immunofluorescence using hamster antimouse CD11c (Santa Cruz Biotechnology) and rabbit antimouse β-catenin (Cell Signaling Technology) mAb. After incubation with secondary goat antirabbit FITC-conjugated IgG (Sigma-Aldrich) and goat antihamster Texas Red-conjugated IgG (Vector), the samples were premounted with Vectashield medium with DAPI (Vector). Positive cells were counted blindly in 10 HPF/section (×200).

Caspase-3 Activity Assay.

Caspase-3 activity was performed and determined by an assay kit (Calbiochem, La Jolla, CA) as described.20

Terminal Deoxyribonucleotidyl Transferase (TdT)-Mediated dUTP-Digoxigenin Nick End Labeling (TUNEL) Assay.

The Klenow-FragEL DNA Fragmentation Detection Kit (EMD Chemicals, Gibbstown, NJ) was used to detect the DNA fragmentation characteristic of oncotic necrosis/apoptosis in formalin-fixed paraffin-embedded liver sections.19, 20 Results were scored semiquantitatively by averaging the number of apoptotic cells/microscopic field at 200× magnification. Ten fields were evaluated/sample.

Quantitative Reverse-Transcription Polymerase Chain Reaction (RT-PCR) Analysis.

Quantitative real-time PCR was performed using the DNA Engine with Chromo 4 Detector (MJ Research, Waltham, MA). In a final reaction volume of 25 μL, the following were added: 1× SuperMix (Platinum SYBR Green qPCR Kit; Invitrogen) cDNA and 10 μM of each primer. Amplification conditions were: 50°C (2 minutes), 95°C (5 minutes), followed by 40 cycles of 95°C (15 seconds) and 60°C (30 seconds). Primers used to amplify specific gene fragments were published.20, 23

Western Blot Analysis.

Proteins (30 μg/sample) from cell cultures or liver samples were subjected to 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to nitrocellulose membrane (Bio-Rad, Hercules, CA). Polyclonal rabbit antimouse TLR4 (Imgenex, San Diego, CA), phos-Stat3, Stat3, phos-β-catenin, β-catenin, phos-GSK-3β, GSK-3β, PTEN, phos-Akt, Akt, phos-IκBa, IκBa, phos-IRF3, IRF3, Bcl-2, Bcl-xl, cleaved caspase-3, and β-actin Abs (Cell Signaling Technology) were used. The relative quantities of proteins were determined by densitometer and expressed in absorbance units (AU).

Statistical Analysis.

Data are expressed as mean ± standard deviation (SD). Statistical comparisons between groups were analyzed by Student's t test. Differences were considered statistically significant at P < 0.05.

Results

STAT3 Activates β-Catenin and Inhibits DC Maturation/Function.

We have shown that STAT3 exerts potent antiinflammatory activity both in vitro and in vivo.20 To delineate whether STAT3-induced β-catenin plays a role in DC maturation/function, mouse LPS-pulsed BMDCs were supplemented with CoPP (HO-1 inducer) or rIL-10. Western blot analysis showed that LPS slightly increased STAT3 phosphorylation (Fig. 1A; 0.5-0.7 AU), whereas addition of CoPP/rIL-10 markedly enhanced phosphorylated STAT3 (2.5-2.7 AU) in BMDCs. Interestingly, DC maturation after CoPP/rIL-10 was accompanied by up-regulation of β-catenin and GSK-3β phosphorylation (2.1-2.4 AU and 2.2-2.4 AU, respectively), compared with LPS-matured BMDCs (0.4-0.6 AU). FACS analysis revealed phenotypic changes in DC maturation program, as demonstrated by CoPP-/rIL-10-mediated depression of otherwise robust LPS-induced CD40, CD80, and CD86 phenotype (Fig. 1B). These data are consistent with a diminished ELISA profile for IL-12p40, TNF-α, and IL-6 in LPS-BMDCs after CoPP/rIL-10 treatment (Fig. 1C). Hence, STAT3-induced β-catenin inhibits BMDCs maturation and function.

Figure 1.

STAT3 activates β-catenin and inhibits DC maturation and function in murine LPS-stimulated BMDCs. Cells were incubated with CoPP (10 μg/mL) or rIL-10 (50 ng/mL) for 6 hours, and stimulated with/without LPS (0.5 μg/mL) for 24 hours. (A) Protein was extracted from BMDCs and the expression of STAT3, β-catenin, and GSK-3β was evaluated by western blots. Representative of three separate experiments. (B) BMDC phenotypic changes were evaluated by flow cytometry. Representative of three separate experiments. (C) The production of IL-12p40, TNF-α, and IL-6 was measured in cell culture supernatants by ELISA. Mean ± SD (n = 3-4 samples/group). *P < 0.0005, **P < 0.0001.

β-Catenin Regulates DC Function in a STAT3-Dependent Manner.

To elucidate the regulatory role of STAT3 and β-catenin on DC function, we disrupted STAT3 signaling in BMDCs by using a small interfering RNA (siSTAT3). This resulted in diminished CoPP-/rIL-10-mediated β-catenin expression (Fig. 2A; 0.2-0.4 AU), compared with nonspecific (NS) siRNA-transfected cells (2.5-2.8 AU). In addition, SnPP (HO-1 inhibitor)-treated cells showed decreased β-catenin levels (0.2-0.6 AU). Interestingly, specific knockdown of STAT3 in CoPP-/rIL-10-treated BMDCs promoted PTEN activation (Fig. 2A; 2.2-2.4 AU) but inhibited Akt phosphorylation (0.2-0.5 AU), as compared with NS siRNA-transfected cells (0.2-0.3 AU and 2.3-2.5 AU, respectively). Furthermore, disruption of STAT3 reversed CoPP or rIL-10-mediated inhibition of LPS-triggered DC maturation, evidenced by increased CD40, CD80, and CD86 expression (Fig. 2B). Consistent with flow cytometry data, the production of IL-12p40, TNF-α, and IL-6 was elevated after blockade of STAT3 in CoPP- or rIL-10-treated, but not in NS siRNA-treated BMDCs (Fig. 2C). Thus, STAT3 knockdown inhibits β-catenin signaling and triggers PTEN/PI3K and DC maturation, suggesting that β-catenin regulates DC function in a STAT3-dependent manner.

Figure 2.

β-Catenin-mediated regulation of DC function in murine LPS-stimulated BMDCs is STAT3-dependent. Cells were transfected with Stat3 siRNA/NS siRNA (10 0nM) and incubated for 24 hours. After washing, cells were treated with CoPP (10 μg/mL) or rIL-10 (50 ng/mL) for 6 hours, and stimulated with/without LPS (0.5 μg/mL) for 24 hours. (A) Western-assisted expression of β-catenin, PTEN, and Akt. Results are representative of three separate experiments. (B) BMDC phenotypic changes were evaluated by flow cytometry. Representative of three separate experiments. (C) ELISA-assisted production of IL-12p40, TNF-α, and IL-6 in cell culture supernatants. Mean ± SD (n = 3-4 samples/group). *P < 0.0001.

Knockdown of β-Catenin Activates PTEN/TLR4 Signaling in DCs.

To further dissect putative mechanisms by which β-catenin may regulate DC function, we disrupted β-catenin signaling in BMDCs by using a small interfering RNA (siβ-cat). As shown in Fig. 3A, LPS-stimulated BMDCs readily induced PTEN (2.3-2.5 AU) and TLR4 (2.6-2.8 AU). Interestingly, disruption of β-catenin in CoPP or rIL-10 pretreated BMDCs led to enhanced expression of PTEN and TLR4 (2.2-2.4 AU and 1.9-2.1 AU, respectively) compared to nonspecific siRNA (siNS)-treated controls (0.3-0.7 AU and 0.2-0.4 AU, respectively). Furthermore, knockdown of β-catenin in CoPP or rIL-10 pretreated BMDCs increased the phosphorylation of IRF3 and IκBα (Fig. 3A; 1.5-1.7 AU and 1.6-1.8 AU, respectively). Similar findings were recorded in LPS-stimulated BMDC without adjunctive CoPP/rIL-10 (Supporting Fig. 3). As PTEN/PI3K signaling regulates TLR4 activation in DCs,24 we used the PTEN phosphate release assay, in which β-catenin knockdown was found to increase PTEN activity (Fig. 3B) in CoPP- or rIL-10-treated LPS-stimulated DCs. These results were consistent with increased expression of CCR2, CCR5, and CXCR3 in siβ-cat-treated DCs, compared with those without β-catenin-silenced cells (Fig. 3C). Thus, disruption of β-catenin activates PTEN/TLR4 signaling in DCs.

Figure 3.

Knockdown of β-catenin in LPS-stimulated BMDCs activates PTEN/TLR4 signaling. Cells transfected with β-catenin siRNA/NS siRNA (100 nM) were treated with CoPP (10 μg/mL) or rIL-10 (50 ng/mL), and then stimulated with or without LPS (0.5 μg/mL). (A) Western-assisted expression of β-catenin, PTEN, TLR4, IRF3, and IκBa. Representative of three experiments. (B) PTEN activity was measured by malachite green phosphate assay. Mean ± SD; n = 4-6 samples/group. *P < 0.05. (C) Quantitative RT-PCR-assisted chemokine expression in LPS-stimulated BMDCs. Each column represents mean ± SD (n = 3-4 samples/group). *P < 0.05.

Knockdown of β-Catenin Increases Hepatocellular Damage in Liver IRI.

Next, we investigated whether disruption β-catenin signaling may affect local inflammatory responses in a mouse liver IRI model. The hepatocellular damage at 6 hours of reperfusion following 90 minutes of partial liver warm ischemia was evaluated by Suzuki's histological grading (Fig. 4A/B). Livers in mice treated with NS siRNA (siNS) plus Ad-HO-1 or Ad-IL-10 showed mild to moderate edema without necrosis (Fig. 4Af/h; score = 1.2 ± 0.42 and 1.1 ± 0.3, P < 0.0001). In contrast, livers in mice after adjunctive β-catenin siRNA (siβ-cat) and Ad-HO-1 or Ad-IL-10 revealed significant edema, severe sinusoidal congestion/cytoplasmic vacuolization, and extensive (30%-50%) necrosis (Fig. 4Ae/g; score = 3.3 ± 0.48 and 3.2 ± 0.42). These data are consistent with hepatocellular function, assessed by sGPT levels (IU/L). Indeed, disruption of β-catenin in Ad-HO-1/Ad-IL-10-transfected mice increased sGPT levels, compared to NS siRNA-treated controls (Fig. 4C; 9,518 ± 3,797 and 9,061 ± 3,374 vs. 781 ± 442 and 561 ± 284, respectively, P < 0.005).

Figure 4.

Knockdown of β-catenin increases hepatocellular damage after liver IR. Mice were subjected to 90 minutes of partial liver warm ischemia, followed by 6 hours of reperfusion. (A,B) The severity of liver IRI was evaluated by Suzuki's histological grading. (a) Sham control; (b) WT (2.8 ± 0.42); (c) Ad-β-gal (3.5 ± 0.53); (d) WT+siβ-cat (3.6 ± 0.7); (e) siβ-cat+Ad-HO-1 (3.3 ± 0.48); (f) nonspecific siRNA+Ad-HO-1 (1.2 ± 0.42); (g) siβ-cat+Ad-IL-10 (3.2 ± 0.42); (h) nonspecific siRNA+Ad-IL-10 (1.1 ± 0.3). Representative of n = 6 mice/group (*P < 0.05; **P < 0.0001); original magnification ×200. (C). Hepatocellular function in serum samples was evaluated by sGPT levels (IU/L). Results expressed as mean ± SD (n = 6 samples/group). *P < 0.05; **P < 0.005.

In parallel experiments, we studied whether β-catenin modifies liver IRI under baseline conditions, i.e., in the absence of adjunctive IL-10 or HO-1. Indeed, knockdown of endogenous β-catenin in otherwise untreated WT mice exacerbated the hepatocellular damage as compared with β-catenin proficient controls, and evidenced by Suzuki's histological grading (Fig. 4Ab/d,B; Suzuki's score = 2.8 ± 0.42 and 3.6 ± 0.7, respectively, P < 0.05) and sGPT levels (Fig. 4C: 7,162 ± 2,657 IU/L in β-catenin proficient and 13,604 ± 6,971 IU/L in β-catenin-deficient WT, P < 0.05).

Knockdown of β-Catenin Enhances DC Activation and PTEN/TLR4-Inflammation in Liver IRI.

To investigate the regulatory role of β-catenin in DC function, we analyzed CD11c+ DC in the ischemic liver lobes by immunohistochemistry (Fig. 5A,B). Indeed, disruption of β-catenin in Ad-HO-1 or Ad-IL-10-transfected livers increased CD11c+ DC infiltration (Fig. 5Ac/e; 25.3 ± 6.9 and 23.6 ± 7.3) compared to the NS siRNA-group (Fig. 5Ad/f: 11.6 ± 3.4 and 9.5 ± 4.3, P < 0.005). Moreover, knockdown of β-catenin in Ad-HO-1/Ad-IL-10-treated livers increased mRNA levels coding for IL-12p40, TNF-α, IL-6, and CXCL-10, as compared with NS siRNA controls (Fig. 5C). These results were supported by western analysis, in which β-catenin knockdown in mice subjected to Ad-HO-1 or Ad-IL-10 diminished the expression of β-catenin (Fig. 5D, 0.2-0.5 AU) in the ischemic liver lobes, whereas NS siRNA followed by Ad-HO-1 or Ad-IL-10 did not affect β-catenin levels (2.0-2.3 AU). Interestingly, the expression of PTEN, TLR4, and phosphorylated IκBα markedly increased after disruption of β-catenin in Ad-HO-1- or Ad-IL-10-treated (2.2-2.4 AU, 2.1-2.3 AU and 2.0-2.2 AU, respectively) but not in NS siRNA-treated (0.5-0.7 AU, 0.2-0.4 AU, and 0.2-0.5 AU, respectively) groups (Fig. 5D).

Figure 5.

Knockdown of β-catenin enhances hepatic DC activation and PTEN/TLR4-mediated inflammation in liver IRI. Liver DCs were detected by immunohistochemical staining using mAb against mouse CD11c (A,B). (a) Sham control; (b) Ad-β-gal; (c) siβ-cat+Ad-HO-1; (d) nonspecific siRNA+Ad-HO-1; (e) siβ-cat+Ad-IL-10; (f) nonspecific siRNA+Ad-IL-10. Results scored semiquantitatively by averaging number of positively stained cells (mean ± SD)/field at 200× magnification. Representative of 4-6 mice/group (*P < 0.005). (C) Western-assisted detection of β-catenin, Akt, PTEN, TLR4, and IκBα. Representative of three experiments. (D). Quantitative RT-PCR-assisted detection of cytokines in mouse livers. Each column represents the mean ± SD (n = 3-4 samples/group). *P < 0.05.

We used immunofluorescence staining to identify and quantify β-catenin (green) and CD11c (red) double-positive cells in IR-stressed livers (Fig. 6A,B). Knockdown of β-catenin decreased (P < 0.005) the frequency of hepatic β-catenin+ DCs in Ad-HO-1/Ad-IL-10-treated mice (Fig. 6Ac/e; mean = 1.8-2.3 cells/HPF) as compared with nonspecific siRNA-conditioned controls (Fig. 6Ad/f; mean = 12.2-15.3 cells/HPF). We observed marginal β-catenin expression in hepatic SEC or hepatocytes after Ad-HO-1/Ad-IL-10 gene transfer. To detect whether knockdown of endogenous β-catenin affected DC function, we isolated DCs from ischemic liver lobes subjected to β-catenin siRNA versus NS siRNA pretreatment. Although disruption of β-catenin signaling did not affect the frequency of CD4+ DC versus CD8α+ DC populations in the liver (Supporting Fig. 4), it did increase (P < 0.005) PTEN activity (Fig. 6C) and IL-12p40 mRNA expression (Fig. 6D) in hepatic DCs, as compared with NS siRNA controls.

Figure 6.

Immunofluorescence staining of β-catenin and CD11c double-positive cells in ischemic liver lobes (A,B). Goat antihamster CD11c (stained red) and rabbit antimouse β-catenin (stained green) mAbs were used. (a) Sham control; (b) Ad-β-gal; (c) siβ-cat+Ad-HO-1; (d) nonspecific siRNA+Ad-HO-1; (e) siβ-cat+Ad-IL-10; (f) nonspecific siRNA+Ad-IL-10. Results scored semiquantitatively by averaging frequency of double-positive cells. Mean ± SD/ HPF (200× magnification). Representative of 4/group (*P < 0.005). (C) PTEN activity measured by malachite green phosphate assay. Mean ± SD; n = 4/group. *P < 0.05. (D) Quantitative RT-PCR-assisted IL-12p40 expression in LPS-stimulated DCs. Mean ± SD (n = 4/group). *P < 0.05.

Knockdown of β-Catenin Increases Apoptosis in IR-Stressed Liver.

We investigated the regulatory role of β-catenin on apoptosis pathways by western blots. By 6 hours of reperfusion after 90 minutes of ischemia, knockdown of β-catenin in Ad-HO-1 or Ad-IL-10-transfected livers down-regulated Bcl-2/Bcl-xL (0.1-0.3 AU and 0.3-0.6 AU, respectively), yet up-regulated cleaved caspase-3 (2.4-2.7 AU) (Fig. 7A). In contrast, the expression of Bcl-2/Bcl-xL strongly up-regulated in NS siRNA-treated livers after Ad-HO-1 or Ad-IL-10 (2.0-2.2 AU and 2.1-2.3 AU, respectively), whereas the expression of cleaved caspase-3 was inhibited in NS siRNA-treated controls (0.3-0.5 AU). These results were confirmed by increased caspase-3 activity in siβ-cat- but not NS siRNA-treated mice (Fig. 7B: 4.12 ± 0.42 and 4.01 ± 0.4 versus 1.19 ± 0.29 and 1.08 ± 0.32, respectively, P < 0.001). We further analyzed IR-induced hepatic oncotic necrosis/apoptosis by TUNEL staining (Fig. 7C,D). Livers in mice treated with siβ-cat showed increased frequency of TUNEL+ cells (Fig. 7Cc/e: 28.6 ± 10.8 and 26.1 ± 11.1, respectively), compared with NS siRNA controls (Fig. 7Cd/f: 6.5 ± 3.6 and 5.5 ± 3.2, respectively, P < 0.0001).

Figure 7.

Knockdown of β-catenin increases IR-induced apoptosis. Mice were subjected to 90 minutes of partial liver warm ischemia, followed by 6 hours reperfusion. (A) Western-assisted analysis of Bcl-2, Bcl-xl, and cleaved caspase-3. Representative of three experiments. (B) Caspase-3 activity in mouse livers. Results expressed as mean ± SD; n = 4-6 samples/group. *P < 0.001. (C/D). Liver apoptosis by TUNEL staining. (a) Sham control; (b) Ad-β-gal; (c) siβ-cat+Ad-HO-1; (d) nonspecific siRNA+Ad-HO-1; (e) siβ-cat+Ad-IL-10; (f) nonspecific siRNA+Ad-IL-10. Results scored semiquantitatively by averaging the number of apoptotic cells (mean ± SD) per field at 200× magnification. Representative of 4-6 mice/group (**P < 0.0001).

Discussion

Both innate and adaptive immune responses are essential in the mechanism of liver IRI.1 By regulating the initial response in damaged/necrotic cells by way of TLR4 signaling, DCs are key mediators of immune homeostasis,25 yet by amplifying innate responses they may also promote the development of adaptive immunity.5, 6 Our results highlight the regulatory role of β-catenin to orchestrate local inflammation, PTEN/PI3K and TLR4 signaling in IR-stressed liver. Our in vitro data support the regulatory function of STAT3-induced β-catenin in DC activation and PTEN/TLR4 signaling. Previous studies have implicated STAT3-mediated antiinflammatory phenotype in LPS-stimulated DCs.26 We found that CoPP- or rIL-10-induced STAT3 triggered translocation of β-catenin from the cytoplasm to the nucleus, and transcription of its target genes in BMDCs. Activation of β-catenin inhibited IL-12p40, TNF-α, and IL-6 expression, as well as DC maturation by down-regulating costimulatory CD40, CD80, and CD86. In addition, our findings suggest that GSK-3β may play a role in β-catenin activation and DC maturation. Interestingly, STAT3 knockdown in LPS-stimulated BMDCs depressed β-catenin and Akt but enhanced PTEN expression, leading to increased DC expression of proinflammatory mediators and costimulatory molecules, suggesting STAT3 can mediate β-catenin activation to program DC functions. We found that knockdown of β-catenin in DCs augmented proinflammatory mediators, and enhanced PTEN/TLR4, to initiate downstream increased CCR2, CCR5, and CXCR3 chemokine program. Moreover, β-catenin knockdown promoted IRF3 activation and phosphorylated IκB to enhance NF-κB activity. Thus, DC proinflammatory phenotype arose from direct control of β-catenin-TLR4 axis.

Next, we determined whether β-catenin signaling is essential for hepatic homeostasis. Although Wnt transcription regulates the cellular redox balance and hepatocytes that overexpress β-catenin were found resistant to IR-damage by way of hypoxia inducible factor (HIF)-1α,27 the crosstalk between β-catenin and host immune responses, pivotal in the mechanism of hepatic IR, remains to be elucidated. We have shown that HO-1-induced STAT3 is required for regulating innate immunity in hepatic IRI.20 In the current study, we used a mouse model of partial liver warm IRI to demonstrate that siRNA-induced β-catenin deficiency exacerbated the hepatocellular damage, assessed by sGPT levels and Suzuki's liver histological grading, in Ad-HO-1/Ad-IL-10-pretreated as well as at baseline conditions in otherwise untreated WT mice. In addition, β-catenin knockdown increased local CD11c+ DC infiltration, implicating β-catenin as a key regulator of inflammatory responses in IR-stressed hepatic DCs. Several factors may contribute to the regulatory function of β-catenin signaling. First, although myeloid/conventional DC (mDC/cDC) become activated in liver IRI by hepatocyte DNA by way of TLR9,28 this DC subset can also crosslink TLR4 ligand to promote adaptive immune activation.5, 6 Indeed, β-catenin knockdown in Ad-HO-1/Ad-IL-10-treated livers enhanced local inflammation by augmenting PTEN/TLR4, IRF3, and NF-κB expression. Thus, β-catenin down-regulates hepatic DC function and downstream signaling that control inflammation in the liver. Second, during IRI, DCs rapidly enter hepatic parenchyma in response to endogenous TLR ligands,4 resulting in TLR4/NF-κB activation and increased production of IL-12, the key cytokine at the innate-adaptive immune interface.29 Indeed, DCs are one of the major IL-12 producers.5, 30 Our results show that β-catenin knockdown in Ad-HO-1/Ad-IL-10-treated livers increased DC-mediated IL-12p40 expression, which further enhanced intrahepatic adaptive immune cascades. Hence, β-catenin is a crucial regulator of TLR4-mediated IL-12 production in IR-stressed liver.

Consistent with our in vitro data, we found that disruption of β-catenin signaling enhanced PTEN activation but inhibited Akt phosphorylation, suggesting the PTEN/PI3K/Akt pathway as an important regulatory mechanism in β-catenin function. Indeed, β-catenin knockdown promoted IκB phosphorylation and increased the TLR4-driven proinflammatory gene program, suggesting that β-catenin may affect TLR4 signaling by way of a negative feedback regulatory mechanism. Furthermore, Akt known to act as an antiapoptotic molecule that promotes cell survival, can also inhibit caspase-mediated cell death through phosphorylation of Bcl-2/Bcl-xL-associated death promoter (BAD), releasing Bcl-2 family members, and directly phosphorylating caspase protease.31 Our in vivo results further support the role of β-catenin-mediated PI3K/Akt in the regulation of hepatic oncotic necrosis/apoptosis. Thus, defective β-catenin down-regulated Bcl-2/Bcl-xL but up-regulated cleaved caspase-3 and its activity, which in turn enhanced apoptotic cell death in IR-stressed livers. Thus, our results highlight the function of β-catenin to trigger PI3K/Akt signaling and ameliorate liver cell death in IRI pathology.

Figure 8 depicts putative molecular mechanisms by which β-catenin signaling may regulate immune responses in the mechanism of liver IRI. STAT3 triggers β-catenin activation by way of GSK-3β phosphorylation. After translocating to the nucleus, β-catenin activates transcription of its target genes, depresses PTEN activity, and promotes PI3K/Akt signaling, to provide a negative TLR4 regulatory feedback to inhibit NF-κB/IRF3 activity, and ultimately suppress proinflammatory gene programs in the liver. Furthermore, PI3K/Akt inhibits IL-12 production and promotes antiapoptotic Bcl-2/Bcl-xL function, which may also limit the hepatocyte death.

Figure 8.

Schematic representation of signaling pathway by which β-catenin may regulate intricate inflammatory responses in liver IRI. See text for details.

In conclusion, this study extends our recent findings on the role of Akt/β-catenin/Foxo1 axis in the mechanism of macrophage innate activation32 by demonstrating that β-catenin may program DC development and regulate innate-adaptive interface in IR-stressed liver. By identifying molecular pathways critical for β-catenin function, our study provides the rationale for novel therapeutic approaches to ameliorate IR-triggered liver inflammation and damage.

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