Protease-activated receptor 2 promotes experimental liver fibrosis in mice and activates human hepatic stellate cells

Authors

  • Virginia Knight,

    1. Center for Inflammatory Diseases, Monash University Monash University, Melbourne, Victoria, Australia
    2. Gastroenterology and Hepatology Unit, Monash Medical Center, Melbourne, Victoria, Australia
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  • Jorge Tchongue,

    1. Center for Inflammatory Diseases, Monash University Monash University, Melbourne, Victoria, Australia
    2. Gastroenterology and Hepatology Unit, Monash Medical Center, Melbourne, Victoria, Australia
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  • Dinushka Lourensz,

    1. Center for Inflammatory Diseases, Monash University Monash University, Melbourne, Victoria, Australia
    2. Gastroenterology and Hepatology Unit, Monash Medical Center, Melbourne, Victoria, Australia
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  • Peter Tipping,

    1. Center for Inflammatory Diseases, Monash University Monash University, Melbourne, Victoria, Australia
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  • William Sievert

    Corresponding author
    1. Center for Inflammatory Diseases, Monash University Monash University, Melbourne, Victoria, Australia
    2. Gastroenterology and Hepatology Unit, Monash Medical Center, Melbourne, Victoria, Australia
    • Gastroenterology and Hepatology Unit, Monash Medical Center, 246 Clayton Road, Melbourne, Victoria 3168, Australia fax: 613 9594 6250
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  • Potential conflict of interest: Nothing to report.

  • This work was supported by grants from the National Health and Medical Research Council of Australia.

Abstract

Protease-activated receptor (PAR) 2 is a G-protein–coupled receptor that is activated after proteolytic cleavage by serine proteases, including mast cell tryptase and activated coagulation factors. PAR-2 activation augments inflammatory and profibrotic pathways through the induction of genes encoding proinflammatory cytokines and extracellular matrix proteins. Thus, PAR-2 represents an important interface linking coagulation and inflammation. PAR-2 is widely expressed in cells of the gastrointestinal tract, including hepatic stellate cells (HSCs), endothelial cells, and hepatic macrophages; however, its role in liver fibrosis has not been previously examined. We studied the development of CCl4-induced liver fibrosis in PAR-2 knockout mice, and showed that PAR-2 deficiency reduced the progression of liver fibrosis, hepatic collagen gene expression, and hydroxyproline content. Reduced fibrosis was associated with decreased transforming growth factor beta (TGFβ) gene and protein expression and decreased matrix metalloproteinase 2 and tissue inhibitor of matrix metalloproteinase 1 gene expression. In addition, PAR-2 stimulated activation, proliferation, collagen production, and TGFβ protein production by human stellate cells, indicating that hepatic PAR-2 activation increases profibrogenic cytokines and collagen production both in vivo and in vitro. Conclusion: Our findings demonstrate the capacity of PAR-2 activation to augment TGFβ production and promote hepatic fibrosis in mice and to induce a profibrogenic phenotype in human HSCs. PAR-2 antagonists have recently been developed and may represent a novel therapeutic approach in preventing fibrosis in patients with chronic liver disease. (HEPATOLOGY 2011)

Hepatic fibrosis occurs in response to acute and chronic liver injury from a variety of sources and may progress to end-stage liver disease with the development of portal hypertension, hepatocellular carcinoma, and liver failure. A substantial body of evidence has identified the hepatic stellate cell (HSC) as the principal source of collagen produced during hepatic fibrogenesis,1 and thus there is considerable interest in factors that regulate HSC activation and collagen expression.

Protease-activated receptors (PARs) are a unique group of G-protein–coupled receptors activated by proteolytic cleavage of their extracellular N terminal domain to reveal a “tethered” ligand that binds with the second extracellular loop of the receptor to initiate signaling. PAR-1 was initially identified in the search for the cellular thrombin receptor, and, to date, four PARs have been identified. Thrombin activates PAR-1, 3, and 4, and factor Xa activates PAR-1 and 2. PAR-2 is also activated by trypsin, mast cell tryptase, and the tissue factor/factor VIIa and factor Xa complex.2 There is a strong linkage between inflammation, coagulation, and fibrosis,3 and a prothrombotic state appears to accelerate liver fibrogenesis.4 One proposed mechanism for this linkage is signaling by coagulation factors through their cellular receptors, PARs, to activate stellate cells.4, 5

PAR-2 is widely expressed in the gastrointestinal (GI) tract on epithelial cells and smooth muscle cells.6 It has been shown to have important, multifaceted roles in the regulation of GI physiology and in inflammatory processes, including pancreatitis, gastritis, and colitis. In the healthy liver, PAR-2 is expressed on hepatocytes, Kupffer cells, bile duct epithelial cells, and endothelial cells of large vessels. Rat HSC express PAR-2 under normal conditions and its expression is markedly increased in liver fibrosis.5 Mast cells are prominently recruited during hepatic fibrosis7 and have the potential to provide a potent source of mast cell tryptase, which can activate PAR-2 receptors. PAR-2 activation augments inflammatory cell recruitment and profibrotic pathways through the induction of genes encoding proinflammatory cytokines and proteins of the extracellular matrix (ECM). PAR-2 activation has been shown to promote pulmonary8 and renal9 fibrosis with increased expression in progressive liver injury,10 but the contribution of PAR-2 to liver fibrosis has not been reported.

We hypothesized that PAR-2 activation promotes hepatic fibrosis in mice and induces HSC proliferation and collagen synthesis. In this study, we show that deletion of PAR-2 diminishes CCl4-induced hepatic fibrosis and that PAR-2 agonists promote HSC proliferation and collagen production.

Abbreviations

CD, cluster of differentiation; cDNA, complementary DNA; ECM, extracellular matrix; ELISA, enzyme-linked immunosorbent assay; GI, gastrointestinal; HSC, hepatic stellate cell; IgG, immunoglobulin G; KO, knockout; MMP, matrix metalloproteinase; mRNA, messenger RNA; PAR, protease activated receptor; PDGF, platelet-derived growth factor; RT-PCR, reverse-transcriptase polymerase chain reaction; αSMA, alpha smooth muscle actin; TGFβ, transforming growth factor β; TIMP, tissue inhibitor of matrix metalloproteinase; WT, wild type.

Materials and Methods

Animals.

PAR-2−/− (PAR-2 knockout; KO) mice, derived on a mixed 129/SvJ and C57BL/6 background, were obtained from Dr. Shaun Coughlin (University of California, San Francisco, CA) and back-crossed 10 generations onto a C57BL/6 background. Their genotype was confirmed by reverse-transcriptase polymerase chain reaction (RT-PCR). Mice were allowed food and water ad libitum and were housed at a constant temperature in a 12-hour light and dark cycle. Experimental protocols were approved by the Monash University Animal Ethics Committee, and mice received humane care as specified under the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes.

CCl4-Induced Hepatic Fibrosis.

Liver fibrosis was induced in male mice by twice-weekly intraperitoneal injections of 1 μL/g body weight of CCl4 mixed with olive oil (1:10), starting between 8 and 10 weeks of age and continuing for 5-8 weeks. Six groups of mice were studied: Two groups received CCl4 for 5 weeks (PAR-2−/−, n = 6; wild-type [WT] C57BL/6, n = 9), and two groups received CCl4 for 8 weeks (PAR-2−/−, n = 8; WT, n = 10). Two control groups of WT C57BL/6 mice (n = 8 each) received olive oil alone for 5 and 8 weeks. Mice were killed 72 hours after the last dose of CCl4, and blood and tissue were collected for analysis.

Fibrosis Assessment.

Liver tissue was fixed in 2% paraformaldehyde for histological examination. Four-micron-thick sections from paraffin-embedded liver tissue were deparaffinized and stained with picrosirius red (Sirius red F3BA 0.1% [w/v] in saturated picric acid) for 90 minutes, washed in acetic acid and water (5:1,000), dehydrated in ethanol, and mounted in neutral DPX. Fifteen consecutive nonoverlapping fields were acquired for each mouse liver, the image was digitized, and fibrosis area was analyzed by Scion Image for Windows (vAlpha 4.0.3.2; Scion Corporation, Frederick, MD).

Determination of Hepatic Hydroxyproline Content.

Hepatic hydroxyproline content was quantified using liver tissue frozen in liquid nitrogen, as previously described, with minor modification.11 Briefly, liver samples were weighed and hydrolyzed in 2.5 mL of 6 N of HCl at 110°C for 18 hours in Teflon-coated tubes. The hydrolysate was centrifuged at 3,000 rpm for 10 minutes; the pH of the resulting supernatant was adjusted to 7.4, and absorbance was measured at 558 nm. Total hydroxyproline content was measured against a standard curve prepared with trans-4-hydroxy-L-proline (Sigma-Aldrich, St. Louis, MO) preparations in the range of 0.156-5.0 μg/mL and expressed per milligram of wet tissue weight.

RNA Purification, Reverse Transcription, and Real-Time Quantitative PCR.

Mouse liver RNA was purified from snap-frozen tissue using the Qiagen RNeasy mini kit, according to the manufacturer's instructions (Qiagen Pty Ltd., Hilden, Germany). RNA from cultured cell lines was isolated using TRIzol (Invitrogen, Carlsbad, CA), as previously described.12 RNA concentration was measured with a Nanodrop ND-100 spectrophotometer (Thermo Scientific, Waltham, MA), and complementary DNA (cDNA) was generated using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA), as per the manufacturer's instructions. Real-time PCR analysis was performed (FastStart Sybr Green; Roche, Mannheim, Germany) using a Rotor Gene 3000 light cycler (Qiagen Pty Ltd., Sydney, Australia), and the specific target messenger RNA (mRNA) of interest was quantified as a ratio relative to 18S RNA content of the sample. The following mouse primers were used: MMP-2 forward: ACC CAG ATG TGG CCA ACT AC, reverse: TCA TTT TAA GGC CCG AGC AA; TIMP-1 forward: ACG AGA CCA CCT TAT ACC AGC CG, reverse: GCG GTT CTG GGA CTT GTG GGC (from Dr. Scott Freidman, Mt. Sinai School of Medicine, New York, NY); 18S forward: GTA ACC CGT TGA ACC CCA TTC, reverse: GCC TCA CTA AAC CAT CCA ATC G (from Dr. Eric Morand, Monash University, Melbourne, Victoria, Australia); TGFβ forward: TGC CCT CTA CAA CCA ACA CA, reverse: GTT GGA CAA CTG CTC CAC CT (Primer 3 software); PAR-1 forward: CTC CTC AAG GAG CAG ACC CAC; reverse: AGA CCG TGG AAA CGA TCA AC (Primer 3 software); and PAR-2 and 18S primers from Applied Biosystems TaqMan probe (Mm00433160_m1, Hs03003631_g1) using TaqMan Gene Expression Master Mix (Applied Biosystems).

Immunohistochemistry.

Paraformaldehyde-fixed 4-micron-thick liver tissue sections were stained with primary antibody for alpha smooth muscle actin (αSMA) (monoclonal mouse antimouse α-SMA; Sigma-Aldrich), F4/80 (rat antimouse, 1:200; a gift of Dr. Richard Kitching, Monash University, Clayton, Victoria, Australia) and cluster of differentiation (CD)68 (rat antimouse CD68, FA11, 1:100; a gift of Dr. G. Koch, Cambridge, UK). The following secondary antibodies were used: αSMA biotinylated rabbit antimouse immunoglobulin G (IgG)2a antibody (1:300; Invitrogen, Carlsbad, CA), F4/80 and CD68 polyclonal rabbit antirat IgG (1:150; Dako, Carpinteria, CA). In brief, sections were dewaxed, rehydrated, and then blocked with 0.6% hydrogen peroxide and CAS protein blocking solution (Invitrogen). Primary antibody incubations for 30 minutes at room temperature (αSMA) and overnight at 4°C (F4/80, CD68) were followed by the application of secondary antibody. Staining was amplified using an avidin-biotin complex kit (Vector Laboratories, Burlingame, CA) and was detected with diaminobenzidine (Dako). Slides were counterstained with Harris hematoxylin. For quantitation of immunoreactivity, 15 consecutive nonoverlapping fields at 250× magnification (α-SMA, F4/80, and CD68) were scored using a graticule eyepiece in a blinded fashion. Negative controls consisted of a mouse IgG1 isotype control antibody (Dako, Glostrup, Denmark) and water substituting for the primary antibody.

Hepatic TGFβ1 Content.

Extracts were prepared from snap-frozen liver by homogenization in lysis buffer (Tris-HCl 50 mM, NaCl 150 mM, ethylenediaminetetraacetic acid 1 mM, 1% Triton X-100, 0.5% Tween-20, and 0.1% sodium dodecyl sulphate), containing a protease-inhibitor cocktail (Roche), followed by centrifugation at 14,000×g for 15 minutes at 4°C. Supernatants were collected and activated with acetic acid/urea before analysis. Transforming growth factor β (TGFβ1) content of liver protein extracts were measured using a mouse TGFβ1 enzyme-linked immunosorbent assay (ELISA) kit (R&D Systems, Inc., Minneapolis, MN). Plates were read using the Bio-Rad (Hercules, CA) microplate reader at 450 nm (with a 540-nm reference filter), and TGFβ1 concentrations were calculated from the standard curve by the plate-reader software.

PAR-1- and PAR-2-Stimulated HSC Collagen and TGFβ1 Production In Vitro.

Immortalized human HSCs (LX-2 cells; a gift from Dr. Scott Friedman) were seeded for 3 days into six-well plates at a density of 1 × 105 cells per well in M199 medium (Gibco, Grand Island, NY) with 5% fetal calf serum. Media were changed at day 3, and human PAR-1 agonist hexapeptide SFLLRN-NH2 (Sigma-Aldrich) and/or human PAR-2 agonist hexapeptide SLIGKV (Sigma-Aldrich) were added at varying concentrations. A scrambled hexapeptide (Auspep, Melbourne, Victoria, Australia) was used as a control. A further dose of either agonist or scrambled peptide was added at 24 and 48 hours, and culture medium and cells were harvested after 72 hours of peptide exposure. The collagen content of the cell-culture supernatant was measured using the Sircol Sirius red dye colorimetric assay (Biocolor, Newtown Abbey, Northern Ireland), as previously described,11 and TGFβ1 content was measured by ELISA.

HSC Proliferation in Response to PAR Activation.

LX-2 cells were seeded onto 96-well plates at a density of 1 × 104 per well in 5% FCS/M199 media and cultured overnight. The PAR-2 agonist peptide, SLIGKV, was added at concentrations from 0 to 100 μM at 24 and 48 hours. Human platelet-derived growth factor (PDGF)-BB (R&D Systems, Minneapolis, MN) was used as a positive control at a concentration of 25 ng/mL. Proliferation of activated HSCs was assessed using a colorimetric bromodeoxyuridine ELISA (Roche), according to the manufacturer's instructions.

Statistical Analysis.

Data are expressed as mean ± standard error of the mean. Statistical significance was determined by one-way analysis of variance with the Newman-Keuls post-test for multiple comparisons or the Student's t test for comparisons between two groups, as appropriate, using GraphPad Prism 5.03 for Windows (GraphPad Software, Inc., La Jolla, CA).

Results

PAR-2 Deficiency Prevents Progression of Histological Hepatic Fibrosis.

WT mice developed significant hepatic collagen deposition in response to CCl4 administration (Fig. 1A). No fibrosis was observed in WT mice given olive oil alone (data not shown). Quantitative analysis of histological fibrosis by computer-assisted morphometry in CCl4-treated WT mice showed marked fibrosis at 5 weeks (1.97% ± 0.16% liver area), which progressed with continued CCl4 exposure over 8 weeks (3.39% ± 0.26%) (Fig. 1C). In PAR-2 KO mice, CCl4 administration induced similar fibrosis to that of WT mice at 5 weeks (2.07% ± 0.26%). However, there was no progression of liver fibrosis with continued CCl4 exposure between 5 and 8 weeks in the PAR-2 KO mice (2.09% ± 0.28%). At 8 weeks, there was significantly less hepatic fibrosis in the PAR-2 KO, compared to WT, mice (P = 0.004) (Fig. 1B,C).

Figure 1.

Hepatic collagen deposition in WT mice (A) and PAR-2 KO mice (B) administered CCl4 for 8 weeks (Sirius red staining, 400×).

Computer-assisted morphometry of Sirius-red–stained liver sections shows no difference in hepatic fibrosis area between WT and KO mice at 5 weeks; however, in contrast to WT mice, there was no progression in fibrosis area in PAR-2 KO mice by 8 weeks (C). Procollagen mRNA expression (D) and hepatic collagen content (E) were significantly lower in PAR-2 KO mice, compared to WT mice, at 8 weeks.

Hepatic Procollagen mRNA and Hydroxyproline Content Are Reduced in PAR-2 KO Mice Exposed to CCl4.

Histological assessment of fibrosis correlated closely with other indices of hepatic collagen content in mice given CCl4. At 8 weeks, PAR-2 KO mice showed significantly less induction of procollagen mRNA (1.8- ± 0.23-fold above untreated mice), compared with WT mice (5.9- ± 1.08-fold; P = 0.002) (Fig. 1D). After 5 weeks of CCl4 administration, similar increases in hepatic hydroxyproline were observed in WT and PAR-2 KO mice (0.45 ± 0.02 μg/mg and 0.43 ± 0.009 μg/mg, respectively) (Fig. 1E). However, after 8 weeks, whereas hepatic hydroxyproline content increased significantly in WT mice, there was no increase in PAR-2 KO mice, compared to levels at 5 weeks. PAR-2 KO mice (0.42 ± 0.026) had significantly less hepatic hydroxyproline, compared to WT mice (0.63 ± 0.03) at 8 weeks (P < 0.002).

PAR-2 Deficiency Is Associated With Reduced Stellate Cell Activation.

αSMA is a marker of HSC activation and myofibroblast differentiation. In WT mice, hepatic fibrosis induced by the administration of CCl4 was accompanied by a progressive increase in αSMA expression at 8 weeks, compared to untreated mice. In PAR-2 KO mice receiving CCl4, induction of αSMA was similar to WT mice treated with CCl4 at 5 weeks (Fig. 2A), but did not increase further, resulting in significantly less αSMA expression, compared to WT mice at 8 weeks (P = 0.014).

Figure 2.

After CCl4 administration, the number of cells expressing αSMA was significantly lower in PAR-2 KO mice at 8 weeks, compared to WT mice (A). TGFβ mRNA expression was significantly lower in PAR-2 KO mice, compared to WT controls, at 8 weeks (B). TGFβ protein levels were lower, remaining at control levels, in PAR-2 KO mice, compared to WT mice (C).

PAR-2 Deficiency Reduces Hepatic TGFβ Expression and Decreases Matrix Metalloproteinase/Tissue Inhibitor of Metalloproteinase mRNA.

CCl4-induced hepatic fibrosis was associated with up-regulation of TGFβ mRNA (3.44- ± 0.72-fold greater than control) and protein (9.2 ± 0.9 pg/mg liver, control 6.9 ± 0.19 pg/mg) in WT mice at 8 weeks. In PAR-2 KO mice, TGFβ mRNA up-regulation was significantly reduced (1.38- ± 0.23-fold of control; P = 0.016, compared to WT) (Fig. 2B), as was TGFβ protein, which was similar to control levels (Fig. 2C).

Matrix metalloproteinases (MMPs) and their specific tissue inhibitors, tissue inhibitors of metalloproteinase (TIMPs), regulate ECM composition and their expression is altered in response to liver injury. In WT mice treated with CCl4 for 8 weeks, both MMP-2 and TIMP-1 mRNA increased, consistent with active ECM remodeling during the development of hepatic fibrosis (Fig. 3A,B). Both MMP-2 and TIMP-1 mRNA expression were significantly reduced in PAR-2 KO mice, compared to WT mice, suggesting that ECM remodeling is reduced in association with the arrest in progression of fibrosis between 5 and 8 weeks in PAR-2 KO mice.

Figure 3.

Expression of MMP-2 mRNA (A) and TIMP-1 mRNA (B) was significantly lower in PAR-2 KO mice, compared to WT mice, at week 8 of CCl4 administration.

PAR-1 mRNA Is Up-regulated at Week 5, but Not at Week 8, in PAR-2 KO Mice.

The temporal pattern of PAR-1 mRNA expression was examined to investigate the potential mechanisms for the lack of early protection against hepatic fibrosis observed in PAR-2 KO mice. In PAR-2 knockout mice at 5 weeks, PAR-1 mRNA expression was significantly up-regulated, compared to CCl4-treated WT mice or untreated controls (Fig. 4A). However, at 8 weeks, PAR-1 expression in the PAR-2 KO mice was not significantly different from WT controls (Fig. 4B). Thus, up-regulation of PAR-1 mRNA may compensate for lack of PAR-2 in the early stages of CCl4-induced fibrogenesis, but this compensatory mechanism is not maintained as fibrosis progresses, resulting in significantly less fibrosis in PAR-2 KOs at 8 weeks.

Figure 4.

PAR-1 mRNA expression was significantly greater in PAR-2 knockout mice, compared to untreated controls and CCl4-treated WT mice, at 5 weeks (A). However, at 8 weeks, PAR-1 expression was similar in the PAR-2 KO mice, compared to WT controls (B).

Decrease in Activated Hepatic Macrophages in Week 8 PAR-2 KO Mice.

We also examined the nature of the inflammatory infiltrate at weeks 5 and 8 to investigate the difference in hepatic fibrosis between PAR-2 KO mice and WT mice observed at week 8. Significantly fewer F4/80+ macrophages were observed at both 5 and 8 weeks in PAR-2 KO mice, compared to CCl4-treated WT mice (Fig. 5A). In addition, at week 8, there were significantly fewer CD68+ macrophages in PAR-2 KO mice, compared to CCl4-treated WT mice, which is a difference that was not observed at week 5 (Fig. 5B). These observations are consistent with a role for PAR-2 in the recruitment, and later activation of, macrophages in CCl4-induced hepatic fibrosis.

Figure 5.

There were significantly fewer F4/80+ macrophages at both 5 and 8 weeks in PAR-2 KO mice, compared to CCl4-treated WT mice (A). At week 8, but not week 5, there were significantly fewer CD68+ macrophages in PAR-2 KO mice, compared to CCl4-treated WT mice (B).

PAR-2 Activation Stimulates HSC Proliferation.

To study the effect of PAR-2 activation directly and specifically in HSCs, we used an immortalized human stellate cell line (LX-2), which has been previously well characterised. Subconfluent cultures of LX-2 cells were stimulated with a specific PAR-2 agonist peptide (SLIGKV) for 48 hours or a scrambled hexapeptide control. The PAR-2 agonist peptide stimulated dose-dependent proliferation of LX-2 cells (Fig. 6A). At the maximum dose of 100 μM, the PAR-2 agonist peptide caused proliferation equivalent to PDGF (25 ng/mL), the most potent inducer of HSC proliferation.

Figure 6.

Stimulation of human LX-2 cells with a PAR-2 agonist peptide significantly increased cell proliferation to levels equivalent to PDGF (**values compared to 0 μM) (A). HSC collagen production was significantly increased by both PAR-2 and PAR-1 agonist peptide (100 μM) and the combination of the two; there was no effect from a scrambled peptide used as control (*values compared to untreated controls) (B). Both PAR-2 and PAR-1 agonist peptides significantly increased TGFβ protein production by HSC; there was no effect from a scrambled peptide used as control (all values compared to untreated control). The combination of PAR-1 and PAR-2 peptides significantly increased TGFβ production by HSC, compared to control peptide and untreated controls, but not to PAR-1 or PAR-2 alone (C).

PAR-2 Activation Increases HSC Collagen Production.

HSCs spontaneously produce collagen during culture on plastic tissue-culture plates. PAR-2 agonist peptide (100 μM) stimulated a significant increase in collagen production by LX-2 cells, whereas the control hexapeptide failed to stimulate collagen production (Fig. 6B). Similarly, PAR-1 agonist peptide (100 μM) stimulated a significant increase in collagen production. The combination of PAR-1 and PAR-2 agonist peptide significantly increased collagen production, compared to control peptide and untreated controls, but not more than the individual agonists alone.

PAR-2 Activation Stimulates TGFβ Production.

TGFβ is spontaneously produced by HSCs in culture. PAR-2 agonist peptide (at 3 different doses) caused a significant increase in TGFβ production by LX-2 cells, compared to the control peptide and untreated controls (Fig. 6C). The threshold for the stimulation of TGFβ production (25 μM) was lower than that for stimulation of collagen production. As expected, TGFβ production also increased after stimulation with PAR-1 agonist peptide. The combination of PAR-1 and PAR-2 agonist peptides caused a significant increase in TGFβ production by LX-2 cells, compared to control peptide and untreated controls. Again, the effect of combined agonist peptides on TGFβ production was not significantly greater than the effect of the individual agonists, suggesting a maximal response at the selected doses.

Discussion

We observed that PAR-2 deficiency in experimental liver fibrosis leads to a reduction in hepatic collagen content and histological fibrosis accompanied by decreased HSC activation, as demonstrated by the reduced expression of αSMA. These findings were paralleled by a decrease in gene and protein expression of the principal profibrogenic cytokine, TGFβ, and altered MMP and TIMP gene expression. We confirmed a specific effect on HSC in vitro by showing that PAR-2 activation stimulated proliferation, collagen production, and TGFβ protein production. These data suggest that PAR-2 activation promotes hepatic fibrosis by inducing a profibrogenic phenotype in HSCs.

PAR-1 has been studied in animal models of hepatic necroinflammation and fatty liver disease10 and in human and murine lung injury.13 PAR-1-deficient mice appear to be protected from CCl4-induced liver fibrosis.14 Thus, there is compelling evidence that thrombin/Xa-induced PAR-1 signaling plays an important role in tissue fibrogenesis.4, 5 Interest in the role of PAR-2 in hepatic fibrosis has developed based on evidence that PAR-2 activation is associated with inflammatory and fibrogenic events in the kidney and pancreas9, 15 and its expression is increased in models of lung injury,8, 16 suggesting an important role for PAR-2 in mediating tissue repair. Cellular mechanisms underlying this role have been proposed by Borensztajn et al., who showed that Factor Xa signaling via PAR-2 induced fibroblast proliferation, migration, and differentiation into myofibroblasts.17

The role of PAR-2 in hepatic inflammation and fibrosis has been examined, to date, only in HSC derived from experimental animals. Gaca et al. demonstrated PAR-2 expression in rat HSC, and showed that PAR-2 agonists induced HSC proliferation and collagen production.18 Fiorucci et al. similarly showed that PAR-2 agonist stimulation of rat HSCs resulted in proliferation and activation.10 To our knowledge, the current study is the first to explore the role of the PAR-2 receptor in liver fibrosis in vivo in PAR-2 knockout mice and in vitro in human HSCs. The use of the KO model is a particular strength of the study that allows us to ascribe a profibrogenic role to PAR-2 unequivocally, because antagonist studies can be troubled by a lack of molecular specificity. These findings significantly expand the evidence linking PAR-2 ligation with hepatic fibrogenesis that occurs most likely through a direct effect on HSC proliferation and collagen production.

We confirmed the role of PAR-2 in HSC activation through studies using the human HSC line, LX-2, which expresses PAR-2. We observed a significant dose response to a specific PAR-2 agonist that achieved a proliferative response comparable to PDGF, the most potent cytokine in regard to stimulating HSC proliferation. In keeping with the overall effect that we saw in the PAR-2 KO mice, we showed that PAR-2 ligation of human HSCs led to increased TGFβ and collagen production. PAR-2 appears to have equivalent effects to PAR-1 in regard to the HSC expression of TGFβ and collagen. We did not demonstrate an additive effect on these responses with the combination of agonists, suggesting maximal stimulation of the common downstream effector pathways of these two receptors under the agonist doses and conditions of in vitro stimulation.

Interestingly, we observed that the protective effect of PAR-2 gene deletion was apparent during more advanced stages of fibrosis, in this case at 8 weeks of CCl4 exposure, rather than at 5 weeks. This raises the question of the nature of the factor(s) leading to PAR-2 activation during continued hepatic injury. PAR-2 activation can be stimulated by trypsin and mast cell tryptase as well as coagulation proteases, such as factor VIIa, Xa, and tissue factor. Mast cells are recruited to the liver during fibrogenesis and their numbers can increase by up to 9-fold in the cirrhotic liver.7 Tryptase accounts for approximately 25% of mast cell protein, and its levels progressively increase with liver injury.19 Thus, we postulated that PAR-2 activation in the injured liver might occur through tryptase generation, given the interval between injury and mast cell accumulation. However, we did not observe any difference in histological staining or gene expression of mast cell chymase, a marker for mast cells, between mice treated for 5 or 8 weeks (data not shown). We then investigated PAR-1 expression in the PAR-2 KO mice and found significant up-regulation of PAR-1 mRNA in PAR-2 KO mice at 5 weeks, which was not observed in WT mice exposed to CCl4 or the vehicle control. Interestingly, at 8 weeks, PAR-1 up-regulation was not evident in the KO mice. Thus, there appears to be compensatory PAR-1 signaling early in fibrogenesis in the PAR-2 KO mice that is lost as fibrosis progresses, which may account for the difference in hepatic fibrosis observed at 8 weeks that was not evident at 5 weeks.

Macrophages play an important role in hepatic fibrogenesis,20 and therefore, we also examined the extent of macrophage infiltration at 5 and 8 weeks. We found that the number of F4/80+ macrophages in PAR-2 KO mice was lower than that in WT mice at both 5 and 8 weeks; however, the number of activated macrophages (CD68+ cells) was significantly lower in the KO mice, compared to WT controls, at 8 weeks. A recent study has shown that PAR-2 and Toll-like receptor 4, which is highly expressed on Kupffer cells and forms a component of the lipopolysaccharide receptor, cooperate to enhance the release of proinflammatory cytokines.21 Fewer activated macrophages observed at 8 weeks in the PAR-2 KO mice may therefore lead to alterations in the inflammatory hepatic microenvironment that could contribute to the decrease in hepatic fibrosis observed in PAR-2 deficiency. Thus, multiple lines of evidence suggest that it is likely that as inflammatory liver disease progresses, increasing expression of PAR-210, 18 and its ligands, such as factor Xa,17 potentiate HSC activation and collagen deposition.

Changes in the expression of MMPs and their specific tissue inhibitors, TIMPs, are complex and vary over time with liver injury. MMP-2 is an autocrine proliferation and migration factor for HSC,22 whose expression can be induced by TGFβ and is typically increased after liver injury. TIMP-1, which inhibits MMP activity, is produced by activated HSCs and its expression is also up-regulated with liver injury and leads to the net accumulation of ECM.23 The decrease in gene expression of MMP-2 and TIMP-1 in PAR-2-deficient mice reflects a relatively static phenotype associated with the failure of either fibrosis progression or regression that we observed in these animals.

Recently, peptide-mimetic compounds have been formulated that bind to PAR-2 and inhibit intracellular responses, including nuclear factor kappa light-chain enhancer of activated B cell activation and interleukin-8 expression as well as PAR-2-induced tissue responses, such as vascular (rat aorta) relaxation.24 These newly developed, specific PAR-2 antagonists may represent a novel therapeutic approach in preventing fibrosis in patients with chronic liver disease and support the need for further research into these unique receptors.

In conclusion, we have demonstrated that deletion of the PAR-2 gene in mice chronically exposed to CCl4 leads to a significant reduction in hepatic fibrosis. The mechanism of this effect is likely to be through a reduction in HSC proliferation and collagen production. These novel findings suggest that PAR-2 may be an important therapeutic target for the treatment of human hepatic fibrosis.

Ancillary