Cholestatic liver injury may activate HSCs (hepatic stellate cells) to a profibrogenic phenotype, contributing to liver fibrogenesis. We have previously demonstrated the involvement of TLR (Toll-like receptor) 7 in the pathogenesis of biliary atresia. In the present study we investigated the ability of TLR7 to modulate the profibrogenic phenotype in HSCs. Obstructive jaundice was associated with significant down-regulation of TLR7. Primary HSCs isolated from BDL (bile duct ligation) rats with obstructive jaundice exhibited reduced expression of TLR7 and increased expression of α-SMA (α-smooth muscle actin) and collagen-α1 compared with sham rats, reflecting HSC-mediated changes. Treatment of primary activated rat HSCs and rat T6 cells with CL075, a TLR7 and TLR8 ligand, significantly decreased expression of MCP-1 (monocyte chemotactic protein-1), TGF-β1 (transforming growth factor-β1), collagen-α1 and MMP-2 (matrix metalloproteinase-2), and inhibited cell proliferation and migration. In contrast, silencing TLR7 expression with shRNA (short hairpin RNA) in T6 cells effectively blocked the effects of CL075 stimulation, reversing the changes in MCP-1, TGF-β1 and collagen-α1 expression and accelerating cell migration. Our results indicate that obstructive jaundice is associated with down-regulation of TLR7 and up-regulation of profibrogenic gene expression in HSCs. Selective activation of TLR7 may modulate the profibrogenic phenotype in activated HSCs associated with cholestatic liver injury.
Persistent liver injury due to cholestasis or hepatitis may result in liver fibrosis that engages a range of cell types [1,2]. HSCs (hepatic stellate cells) are activated and undergo morphological and functional transdifferentiation, converting from vitamin A-storing cells to contractile myofibroblastic cells responsible for ECM (extracellular matrix) production in the injured liver [1–3]. This response is transient if the injury is acute and reversible; however, if the injury is chronic, activated HSCs will continue to produce profibrogenic and ECM proteins that ultimately compromise liver function [1–3].
The TLR (Toll-like receptor) family is the most well-characterized class of pattern recognition receptors that signal to host cells in the presence of infection in mammalian species . In a previous study, we demonstrated the involvement of TLRs, and TLR7 in particular, in the pathogenesis of biliary atresia . In addition, a TLR7 single nucleotide polymorphism has been shown to protect men with chronic HCV (hepatitis C virus) infection from advanced inflammation and fibrosis . The mechanism through which TLR7 activation protects patients from fibrosis is not clear. Since HSCs play a central role in liver fibrogenesis, it is important to determine whether TLR7 can modulate the activity of HSCs in cholestatic or hepatitis-associated liver injury. Other members of the TLR family, such as TLR4, have been shown to directly stimulate HSCs to induce the pro-inflammatory phenotype, including up-regulation of chemokines, adhesion molecules and profibrogenic genes [7,8]. However, it has also been demonstrated that TLR9 signalling may inhibit the migration of HSCs, despite enhancing collagen production in HSCs by apoptotic hepatocyte DNA .
Growing evidence suggests that either removal of the aetiological agent/condition or implementation of effective therapies can result in significant regression of liver fibrosis, in part as a result of the reversal of activated HSCs to a more quiescent phenotype through apoptosis, senescence or other mechanisms [10,11]. We hypothesize that TLR7 expression alleviates fibrogenesis through modulation of the HSC phenotype. In the present study we investigated the expression of TLR7 and profibrogenic proteins in an in vivo model of cholestatic liver injury and the in vitro effects of TLR7 agonist and antagonist treatment on the activation of HSCs and rat T6 HSCs.
The study protocol was approved by the Animal Ethics Committee of Chang Gung University, Taiwan. Male Sprague–Dawley rats were purchased from the National Laboratory Animal Center (Academia Sinica, Taipei, Taiwan) after weaning. The animals were maintained on standard laboratory rat chow and housed with a 12 h light–dark cycle. Rats were anaesthetized by intraperitoneal injection of 50 mg/kg thiopentone sodium (Pentothal, Abbott Laboratories). The blood for liver function tests was withdrawn through an indwelling catheter in the right femoral vein. For those assigned to the BDL (bile duct ligation) group in the obstructive jaundice model, the extrahepatic bile duct was identified, doubly ligated with 5-0 silk sutures and transected at a level 0.7–0.8 cm distal to the last bifurcation. For those assigned to the sham group, the silk suture was passed through the extrahepatic bile duct without ligation or transection. At 2 weeks after BDL or sham ligation, the animals were killed by overdose of thiopentone sodium. Blood tests for liver function were conducted and livers were collected. A portion of the liver from each rat was snap-frozen for protein and mRNA determination, whereas the remaining liver tissue was fixed in 4% paraformaldehyde and embedded in paraffin for histological and immunohistochemical studies. Complete datasets from 10 animals in each group (BDL and sham) were analysed.
Stellate cell isolation and culture
Primary HSCs were isolated from male Sprague–Dawley rats (weighing 450–750 g each) by sequential digestion of the liver with pronase and collagenase, followed by density gradient centrifugation in 8.5% Nycodenz (Sigma–Aldrich) as described previously . The purity of the HSCs was assessed by autofluorescence of stored retinoids in HSC lipid droplets. Cell viability determined by a Trypan Blue exclusion assay revealed that more than 95% of the cells were viable. Purity of the HSC culture was found to be 95–99%. HSCs were either used directly for RNA isolation, protein extraction or culture on uncoated polystyrene dishes. Cells were maintained in DMEM (Dulbecco's modified Eagle's medium) supplemented with 5% FBS (fetal bovine serum). After 1 day in culture, the HSCs had a quiescent phenotype and they developed an activated phenotype after 7–14 days. The passage of the cultured cells was conducted after reaching confluence and experiments were carried out using cells between passages 2 and 6.
Culture of T6 cells and treatment with CL075
The T6 cell line used in the present study was derived from immortalized rat HSCs transfected with the SV40 (simian virus 40) large T-antigen containing a Rous sarcoma virus promoter . T6 cells were maintained in Waymouth's medium (Invitrogen) supplemented with 10% heat-inactivated FBS, 100 units/ml penicillin and 100 mg/ml streptomycin and were routinely incubated at 37°C in 5% CO2. The cells were plated at a density of 5×106 cells per 10-cm culture dish and were treated with various concentrations of the TLR7/8 agonist CL075, a thiazoloquinoline compound (also known as 3M002, InvivoGen), and harvested at different time intervals for Western blot and RNA analysis.
Transfection of T6 cells with TLR7-targeting shRNA (short hairpin RNA)
T6 cells (1×105 cells in six-well plates) were transfected with either Turbo GFP (green fluorescent protein) shRNA as a non-targeting control or TLR7 shRNA (clone TRCN0000065984, Sigma–Aldrich) using ICAFectin® 441 reagent (Eurogentec Biologics). The shRNA clone was constructed within the lentivirus plasmid vector pLKO.1-Puro . The pLKO.1-Puro vector contains ampicillin and puromycin antibiotic resistance genes for selection of bacterial or mammalian cells respectively. At 48 h after transfection, we aspirated the medium and replaced it with fresh medium containing 5 μg/ml puromycin. This selection medium was refreshed every 2 days. We monitored the cells daily and observed the percentage of surviving cells stably expressing TLR7 shRNA or control shRNA plasmids. At approximately 4 weeks following transfection, stable cells were harvested and prepared for Western blot and qRT-PCR (quantitative real-time PCR) analysis.
Frozen rat liver samples (0.1 g per sample) were homogenized in 1 ml TRIzol® (Invitrogen), and total RNA was isolated according to the manufacturer's instructions. For cDNA preparation, 2 μg of total RNA was added to 0.1 μg of oligo-(dT)15, according to the manufacturer's instructions (Invitrogen). Sequences of the PCR primers were designed based on cDNA sequences from GenBank® (Supplementary Table S1 at http://www.BiochemJ.org/bj/447/bj4470025add.htm). GAPDH (glyceraldehyde-3-phosphate dehydrogenase) was used as the internal control gene to analyse the relative mRNA expression of the following transcripts: TLR1–11, MCP-1 (monocyte chemotactic protein-1), TGF-β1 (transforming growth factor-β1) and MMP-2 (matrix metalloproteinase-2). qRT-PCR was performed using SYBR Green PCR Master Mix and each sample was analysed in duplicate. Quantification of the mRNA levels for each of the genes of interest was achieved using the 7500 Fast Real-Time PCR system (Applied Biosystems) and comparative methods. The quantity of mRNA was calculated using the ΔCt method. In this method, Ct values for each gene were normalized to the Ct value of a housekeeping gene (GAPDH) within the same reaction. The results are presented as 2−ΔΔCt (ΔCt=CtGAPDH–Cttarget) and expressed as the fold increase/decrease in gene expression compared with T6 or primary HSC untreated control. The primers for TLR1–11, MCP-1, TGF-β1, MMP-2, IL-6 (interleukin 6), type 1 collagen and GAPDH are shown in Supplementary Table S1.
Western blot analysis
Crude protein extracts (30 μg) were treated with sample buffer and then boiled for 10 min, separated by SDS/PAGE (12% gel) and transferred on to nitrocellulose membranes. The blots were incubated with primary antibodies targeting α-SMA (α-smooth muscle actin; 1:200 dilution; Abcam), collagen-α1 (Sigma–Aldrich), TLR7 (ENZO Life Sciences), MMP-2 (Santa Cruz Biotechnology) and GAPDH (Sigma–Aldrich) according to the manufacturer's recommended protocols. Detection was achieved using a chemiluminescence substrate (Santa Cruz Biotechnology) and exposure to film. Signals were quantified by densitometric analysis.
Liver tissues were embedded in TissueTek® OCT™ (optimal cutting temperature) compound (Sakura Finetek) and frozen at −80°C for storage. Frozen sections (4 μm thick) were prepared using a cryostat (CM3050 S, Leica). Cryosections and cell culture slides were fixed with isotonic PBS and 4% paraformaldehyde solution for 1 h. To block non-specific background staining, the samples were incubated in a solution containing 1% BSA for 30 min. After washing with PBS, the slides were incubated with primary antibodies for 1 h at room temperature (25°C) in the dark. The primary antibodies used in liver tissues were anti-GFAP (glial fibrillary acidic protein; 1:100 dilution; Abcam) and anti-TLR7 (1:200 dilution; ENZO Life Sciences), whereas an anti-collagen-α1 primary antibody (1:2000 dilution; Sigma–Aldrich) was used with cell culture slides. Alexa Fluor® 488- and Alexa Fluor® 594-conjugated secondary antibodies (Molecular Probes) were used. For F-actin (filamentous actin) staining, Alexa Fluor® 488 phalloidin (Molecular Probes) was used. Cells were co-stained with DAPI (4′,6-diamidino-2-phenylindole; Molecular Probes) to visualize the nuclei. The stained cells were mounted with fluorescent mounting medium (Dako Cytomation) and visualized by Olympus FluoView® confocal microscopy. All of the exposure gains and rates were consistent among samples. Fluorescence intensities were quantified on independent colour channels.
Cell migration detected by wound-healing assay
Cells were seeded into ibidi culture inserts at a concentration of 10000 cells per well. After allowing the cells to attach overnight, the culture insert was gently removed using sterile forceps. Cells were incubated with 5 μg/ml CL075 and monitored for 72 h. Images were taken at 0, 24, 48 and 72 h with a D5000 digital camera using inverted Nikon TE300 microscope and superposed using PhotoImpact (Adobe). The number of cells that migrated into the wound space were manually counted in three fields per well under a light microscope at ×50 magnification. Areas were quantified by image analysis using Wimasis image analysis software (ibidi).
Gelatin zymography was performed on protein extracts as described previously . Briefly, samples containing 50 μg of total protein were mixed with SDS sample buffer consisting of 0.4 M Tris (pH 6.8), 50 g/l SDS, 200 g/l glycerol and 0.3 g/l Bromophenol Blue and then subjected to gelatin zymography by SDS/PAGE (10% gel) copolymerized with 1 mg/ml gelatin (Sigma–Aldrich). The gels were washed in 2.5% Triton X-100 for 2 h at room temperature to remove the SDS, and then incubated in MMP activation buffer containing 50 mmol/l Tris/HCl (pH 7.5), 150 mmol/l NaCl, 10 mmol/l CaCl2 and 0.05% NaN3 overnight at 37°C. Following 0.25% Coomassie Blue staining and subsequent destaining with 45% methanol and 10% acetic acid, the detection of clear bands on the gel indicated the presence of gelatinase activity.
Colony formation assays are based on the principle that the stable expression of certain proteins can cause either cell-cycle arrest or cell death, hence leading to a reduction in colony number. T6 cells (200) were seeded into a 12-well plastic plate with 1 ml of Waymouth's medium containing 10% FBS and incubated overnight at 37°C. The cells were incubated with various concentrations of CL075 (0, 1, 5 or 10 μg/ml) and cultured for 5–10 days. The cells were then stained with 0.5% Crystal Violet and pictures of Petri dishes were taken using a compact camera. The number of colonies in each plate was counted, and the size of each colony was measured using ImageJ (http://rsbweb.nih.gov/ij/). Additionally, the number of colonies growing in each sample plate was counted and compared with the number of colonies growing in control cultures.
Cell proliferation was assessed using a WST-1 (water-soluble tetrazolium salt 1) assay. T6 cells were plated at 5×103 cells/well in 96-well plates in Waymouth's medium containing 10% FBS and incubated overnight at 37°C. The cells were incubated with various concentrations of CL075 (0, 1, 5 or 10 μg/ml). Untreated cells served as control. After 24 or 48 h of incubation, 10 μl of WST-1 (Roche) was added to each well and incubated for an additional 2 h. The absorbance of samples was measured at 450 nm. Each assay was performed in triplicate and repeated three times. The cell proliferation rate [proliferation rate=(A450 experiment group/A450 control group)×100%] was plotted against time.
All of the results are expressed as means±S.E.M. Student's t test (unpaired, 2-tailed) was used for comparison between experimental groups with continuous variables. A P value of less than 0.05 was considered statistically significant.
Up-regulation of α-SMA and collagen-α1 is associated with down-regulation of TLR7 expression in cholestatic rat liver
Up-regulation of α-SMA and collagen-α1 protein can be used as an indicator of the activation state of HSCs and liver fibrosis. Using Western blot analysis, we found significantly higher expression of α-SMA and collagen-α1 protein, but lower expression of TLR7 protein, in tissues from the BDL group than in tissues from the sham group (Figures 1A–1C). To further characterize TLR7 protein expression in vivo immunofluorescence staining was performed. To verify TLR7 expression in HSCs, immunofluorescence images were merged to allow for the colocalization of TLR7 and GFAP within the same cell. As illustrated in Figure 1(D), liver sections from both the sham and BDL groups were stained for TLR7 (green) and GFAP (red), a specific marker of HSCs (arrow) . Compared with cells isolated from BDL rats, hepatocytes, bile ductular epithelial cells and double-stained non-parenchymal cells morphologically identical with HSCs (arrowhead) isolated from sham rats exhibited stronger TLR7 immunoreactivity.
Expression of α-SMA, collagen-α1 and TLR7 in livers of rats with BDL or sham controls
The liver contains many different types of cells that could serve as sources of TLR expression. Thus we further examined the expression of TLR mRNA transcripts by qRT-PCR in activated HSCs isolated from the BDL rats. Activated HSCs from the BDL group showed differential expression of TLR mRNAs. We found that whereas the mRNA levels of TLR4, -7 and -9 were significantly down-regulated in the BDL group compared with the sham group (Figure 2), the mRNA levels of TLR1, -2, -3, -5, -6, -8, -10 and -11 were not significantly different between the two groups.
Expression of TLR genes in BDL-activated HSCs and sham-operated controls
TLR7 regulates the activation of primary HSCs and transformation of T6 cells
The activation of HSCs is known to lead to increased expression of several profibrogenic genes, including MCP-1, TGF-β, and MMP-2 during the progression of hepatic inflammation and fibro-genesis [17,18]. We treated activated primary HSCs (14-day cultures) isolated from Sprague–Dawley rats with CL075 (a ligand for TLR7 and TLR8) and found a significant increase in TLR7 transcript levels after 1 and 3 h of treatment (Figure 3A), and this was also associated with a significant decrease in MCP-1, TGF-β1, MMP-2 and collagen-α1 transcript levels (Figure 3B). The same trend was also found in the T6 cell line (Figure 3C), which essentially exhibits characteristics of activated HSCs.
Expression of TLR7 protein and profibrotic genes in activated HSCs after treatment with the TLR7 agonist CL075
To test the effects of TLR7 inhibition on the expression of MCP-1 and TGF-β1, T6 cells stably expressing TLR7 shRNA, control T6 or control shRNA (GFP-labelled non-targeting shRNA) were used. As expected, TLR7 shRNA significantly down-regulated TLR7 mRNA expression (Figure 3D) and this was associated with up-regulation of MCP-1 (Figure 3E) and TGF-β1 (Figure 3F) mRNA. Moreover, expression of TLR7 shRNA also led to a significant up-regulation in α-SMA mRNA (Figure 4A) and protein (Figure 4B) expression compared with the control. Immunofluorescence staining also confirmed the findings of the Western blotting (Figures 4C and 4D).
Expression of α-SMA in T6 cells after treatment with CL075
Since TLR8 belongs to the same family as TLR7, we studied the expression of TLR8 in the T6 cell line. Baseline TLR7 and TLR8 expression was almost the same in T6 cells, but stimulation of T6 cells with CL075 caused a significant increase in TLR7, but not TLR8, expression (Supplementary Figure S1A at http://www.BiochemJ.org/bj/447/bj4470025add.htm). Furthermore, down-regulation of TLR7 with TLR7 shRNA, but not down-regulation of TLR8 with TLR8 shRNA, was associated with a significant increase in both α-SMA and collagen-1 expression (Supplementary Figure S1B).
Treatment with a TLR7 agonist reduces T6 cell proliferation
To examine the effects of treatment with the TLR7/8 agonist CL075 on cell proliferation, we conducted a WST-1 test; this revealed no significant effects of CL075 on cell proliferation during the 2-day study period (Figure 5A).
Effects of TLR7 agonist on proliferation and colony formation in T6 cells
Next, to test the effects of CL075 on HSC colony formation, equal numbers of T6 cells were seeded at very low densities, treated with various concentrations of CL075 (0, 1, 5 or 10 μg/ml) and allowed to grow for 10 days. Equal numbers of colonies were found in cells treated with 0, 1 and 5 μg/ml CL075, but larger colony sizes were observed in T6 cells without CL075 treatment and in T6 cells treated with 1 μg/ml CL075 than in T6 cells treated with 5 μg/ml CL075. At 10 μg/ml CL075, the number of formed colonies decreased 2.8-fold and the colony size decreased 32-fold when compared with the control cells (Figure 5B).
To test whether down-regulation of TLR7 could inhibit HSC proliferation, equal numbers of T6 cells and T6 cells stably expressing TLR7 shRNA or GFP-labelled no-targeting shRNA were seeded at very low densities, treated with various concentrations of CL075 (0, 1, 5 or 10 μg/ml), and allowed to grow for 5 days (for the evaluation of colony size) or 10 days (for the quantification of colony number). More rapid proliferation with larger colony size was observed in T6 cells stably expressing TLR7 shRNA without CL075 treatment (Figure 5C). No dose response was observed when the cells were treated with 0–5 μg/ml CL075 with respect to colony number (Figure 5D), but a dose-dependent decrease in colony size was observed in T6 cells, irrespective of TLR7 expression (Figure 5E). However, when cells received the same dose of CL075, T6 cells stably expressing TLR7 shRNA still had more cells per colony than the other groups (Figures 5C and 5E).
TLR7 regulates T6 cell migration
In order to assess whether TLR7 signalling may regulate HSC migration, a wound-healing assay was performed using primary HSCs and T6 cells. Confluent monolayers of primary HSCs, T6 cells and T6 cells stably expressing TLR7 shRNA or GFP-labelled non-targeting shRNA were incubated with or without CL075. Migration assays were conducted using ibidi inserts. After overnight incubation, the inserts were removed and the cells were allowed to proliferate and migrate into the wound for up to 72 h. The results of the present study showed that CL075 treatment inhibited the migration of primary HSCs (Figures 6A and 6B) and T6 cells (Figure 6C and 6D) into the wounded area. Inhibition of TLR7 signalling by TLR7 shRNA could restore the migration capacity of T6 cells, but this restoration was reversed in GFP-labelled shRNA T6 cells following CL075 treatment (Figures 6C and 6D).
Migration of primary activated HSCs and T6 cells was measured by wound healing assay after treatment with CL075
Furthermore, since degradation of the ECM by MMP-2 is required for the migration of activated HSCs , we investigated MMP-2 activity in rat T6 cells by gelatin zymography. We found significant down-regulation of MMP-2 activity in T6 cells treated with CL075 (Supplementary Figure S2 at http://www.BiochemJ.org/bj/447/bj4470025add.htm).
TLR7 regulates F-actin and collagen-α1 expression in T6 cells
We next examined the effects of TLR7 stimulation on the expression of F-actin. F-actin is known to regulate cell migration and collagen-α1 production. Using fluorescent immunostaining (Figures 7A–7C) and Western blotting (Figure 7D), we found that CL075 inhibited both F-actin and collagen-α1 expression in T6 cells. In contrast, shRNA targeting TLR7 markedly reversed CL075-induced inhibition of F-actin and collagen-α1 expression, further confirming the role of TLR7 in the modulation of cell migration and collagen production in HSCs.
Expression of the ECM protein F-actin and collagen-α1 in T6 cells after treatment with CL075
In the healthy liver, low levels of TLRs are widely expressed on parenchymal and non-parenchymal cells. TLRs play important roles in wound healing and regenerative processes, but are also involved in the pathogenesis and progression of various inflammatory and autoimmune liver diseases, chronic hepatitis infections and fibrogenesis [20,21]. Stimulation of TLRs directly or indirectly causes release of multiple cytokines, including type 1 and type 2 IFNs (interferons), induction of pathways that destroy intracellular pathogens and priming of the adaptive response by activation of immature dendritic cells and induction of their differentiation into professional antigen-presenting cells [22,23].
In the present study we demonstrated that obstructive jaundice in rats is associated with the down-regulation of TLR7 and up-regulation of profibrogenic gene expression in liver tissue and isolated primary HSCs. Selective activation of TLR7 in cultured, activated HSCs by treatment with the TLR7 agonist CL075 may modulate the profibrogenic phenotype by decreasing MCP-1, TGF-β1 and α-SMA expression and inhibiting MMP-2 activity and F-actin expression. These phenomena are reversed by TLR7 shRNA in T6 rat HSC cell lines.
The above findings provide a novel look at TLRs in the regulation of HSCs. One of the most important findings of the present study showed that there was differential expression of TLRs in activated HSCs isolated from BDL rats; TLR4, -7 and -9 mRNA were significantly down-regulated on BDL rats compared with sham-operated rats. Of these TLRs, TLR4 is known to directly stimulate HSCs inducing the pro-inflammatory phenotype and profibrogenic genes, whereas TLR9 signalling may enhance collagen production, but inhibits HSC migration . Previous studies that investigated the role of the TLR system in liver fibrosis largely focused on TLR4 and TLR9. In the present study we found that TLR7 plays a distinct role in HSCs. Although the abundance of several TLR mRNAs may have been affected by BDL, only TLR4, -7 and -9 displayed significant reductions (Figure 2). The present study has potential limitations in that TLR2, -3 and -6 did not achieve statistical significance, which may be due to additional variability in the sham groups. Although these differences in TLR signalling may be explained by the use of different models of liver injury and fibrogenesis, it is possible that distinct TLR-dependent mechanisms could also be involved.
The liver is constantly exposed to infectious pathogens, particularly the cholestatic or cirrhotic liver [8,24,25]. Cellular stress or liver damage, including that caused by alcohol consumption, may trigger the release of endogenous molecules into the extracellular space surrounding liver cells leading to activation of the host's innate immune system and TLR proteins . In previous studies, bacterial DNA has been detected in the serum of patients  and animals  with advanced cirrhosis, potentially increasing the amount of matrix deposition and inducing HSC activation via TLR9 signalling . Reducing the amount of portal bacterial DNA or inhibiting its interaction with TLR9 may attenuate the progression of fibrosis in patients with chronic liver disease . Chronic infection with HCV leads to development of liver fibrosis, causing morbidity and mortality. HCV-induced TLR7 mRNA instability results in low levels of TLR7 protein expression and function. A clinical study demonstrated that the selective TLR7 agonist isatoribine contributes to anti-HCV activity through stimulation of innate immunity . TLR7-mediated immunity against HCV involves type I IFN production by immune cells, such as plasmacytoid dendritic cells or leucocytes [30,31], and other endogenous antiviral immune mediators expressed by HCV-infected hepatocytes . Moreover, a TLR7 single nucleotide polymorphism was found to offer protection from the development of inflammation and fibrosis in men with chronic HCV infections . The results of the above studies are consistent with the present study in that the in vitro activation of TLR7 decreased profibrogenic protein production in activated HSCs, potentially providing another way to attenuate liver fibrosis in vivo.
MCP-1 is one of the most significant chemokines regulating the signalling of monocytes and macrophages, and it plays an important role in the recruitment and maintenance of inflammatory infiltrates during liver injury . Non-parenchymal cells, predominantly activated HSCs and biliary epithelial cells are responsible for MCP-1 production and HSC recruitment and activation in chronic liver disease [17,28,33,34]. The results of the present study revealed an increase in MCP-1, TGF-β1 and collagen-α1 mRNA transcripts in the cholestatic liver and in isolated HSCs (Figures 1 and 3). Treatment of HSCs with the TLR7 agonist CL075 significantly decreased the profibrogenic gene expression, as indicated by the reduction in mRNA transcripts of the above targets (Figure 3).
TLR3, TLR7, TLR8 and TLR9 are localized in the intracellular compartment and specialize in the detection of viral nucleic acids. In the normal host, nucleic acids cannot trigger intracellular TLRs. Endogenous and exogenous single-stranded RNAs are ligands for TLR7. TLR7 is transported to endosomes within the cell during viral infection or during activation of autoimmune diseases [35,36]. TLR7 agonists can increase pathogen clearance and antiviral effects during systemic infections, but have not been reported to modulate liver injury or fibrogenesis through TLR7-mediated regulation of the activated HSCs. CL075 (thiazoloquinoline) is an agonist for both TLR7 and TLR8 . We demonstrated that TLR7 shRNA significantly blocked CL075-induced down-regulation of profibrogenic gene expression and restored the migration capacity of activated HSCs, indicating a role for TLR7 signalling in modulating the HSC phenotype. Although TLR8 has been shown to be functional in humans, several studies using TLR7-deficient mice have indicated a controversial role for TLR8 in mice [38,39]. We also found no significant change in TLR8 protein in T6 cells upon treatment with CL075. Furthermore, unlike TLR7, down-regulation of TLR8 with TLR8 shRNA was not associated with a significant increase in both α-SMA and collagen-α1 expression. Therefore the role of TLR8 in the modulation of liver fibrosis through regulation of HSCs still requires clarification.
Inhibition of the fibrogenic, proliferative and migratory effects of HSCs is an emerging experimental therapy for the prevention and regression of hepatic fibrosis [1,2,12]. The most comprehensively characterized target genes described in HSCs include α-SMA, collagen-α1, MCP-1, TGF-β1, TGF-β1 receptors and MMP-2 [1,40,41]. In the present study, we showed that the TLR7 agonist CL075 could inhibit these genes and proteins and that this modulation was associated with reduced proliferation and migration in HSCs. Moreover, in the present study we observed no significant changes in cell proliferation by WST-1 assay or immunoblotting with antibodies against caspase 3 (results not shown) for either activated HSCs or for T6 cells, suggesting that induction of the antifibrogenic phenotype is not necessarily associated with inhibition of cell growth in HSCs.
In summary, the results of the present study indicate that TLR7 signalling in activated HSCs may modulate the profibrogenic phenotype, thus supporting the use of TLR7 agonists as potential therapies to treat liver fibrosis associated with cholestatic liver injury in the future.
bile duct ligation
fetal bovine serum
glial fibrillary acidic protein
green fluorescent protein
hepatitis C virus
hepatic stellate cell
monocyte chemotactic protein-1
quantitative real-time PCR
short hairpin RNA
α-smooth muscle actin
transforming growth factor-β1
water-soluble tetrazolium salt 1
Ming-Huei Chou performed experiments and drafted the paper; Ying-Hsien Huang participated in the discussion and helped analyse the data; Tsun-Mei Lin participated in the discussion and conceiving part of the experiments; Yung-Ying Du performed experiments; Po-Chin Tsai and Chih-Sung Hsieh co-ordinated the study and provided technical support; and Jiin-Haur Chuang participated in the discussion, provided scientific advice and revised the paper. All of the authors read and approved the final paper.
We thank Professor S.H.H. Chan (Center for Translational Research in Biomedical Sciences, Kaohsiung Chang Gung Memorial Hospital, Kaohsiung, Taiwan) for confocal microscope instrumental support, Dr Y.H. Kao (E-DA Hospital, Kaohsiung, Taiwan) for providing the HSC-T6 cell line, I-Ya Chen for excellent help with the data analysis, and C.Y. Lin for providing the animal samples.
This work was supported by the National Science Council of the Republic of China, Taiwan [grant numbers HSC 97-2314-B-182-010-MY3 and NSC 99-2314-B-182A-032 -MY2].
These authors contributed equally to this work.