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Introduction
Hepatic fibrogenesis occurs as a wound-healing process after various chronic liver injuries, including virus infection,
autoimmune liver diseases, and sustained alcohol abuse. Hepatic fibrosis eventually results in end-stage liver cirrhosis if it
is not treated effectively. Hepatic stellate cells (HSC), previously known as fat- or
vitamin A-storing cells or Ito cells, are the most relevant cell
type for the development of hepatic
fibrosis[1-3]. During liver injury, regardless of etiology, HSC become
active and transdifferentiate into myofibroblast-like cells characterized by an increase in cell proliferation, loss
of vitamin A-storing capability, expression of
a-smooth muscle actin (a-SMA), and overproduction of
extracellular matrix (ECM). Much research carried out from a therapeutic perspective has focused on searching for novel agents with inhibitory effects on HSC
proliferation and activation to prevent hepatic
fibrogenesis[4,5].
Cytokines and growth factors such as transforming growth factor
beta (TGF-b) participate in these complicated processes,
as evidenced by TGF-b-induced overproduction of extracellular matrix proteins in activated
HSC[6].TGF-b acts as a potent stimulatory signal for connective tissue formation during wound repair and in fibrotic conditions, whereas the growth
stimulatory effect of TGF-b appears to be mediated via an indirect mechanism involving
connective tissue growth factor (CTGF, also known as hypertrophic chondrocyte-specific gene product 24 or Hcs24). CTGF, belonging to the CCN
(CTGF, Cystein rich protein-Cyr61, and Nephro-blastoma
overexpressed gene-Nov; CCN) family of immediate early proteins, is a 38
kD secretion polypeptide with abundant of cysteines in the C-terminal, which are involved in cell proliferation, migration and
matrix production[7,8]. CTGF is specifically induced by
TGF-b1 and is considered to be the downstream response element and
effective molecule of TGF-b1. It has been reported that
CTGF is the key regulator of extracellular matrix production and plays
an important role in hepatic and renal fibrosis, thus
the TGF-b1- CTGF signal pathway might be an effective target for reversal
of hepatic fibrosis[9,10].
Peroxisome proliferator-activated receptors (PPAR), including
a, b/d and g are a family of ligand-activated nuclear
transcriptional factors that are emerging as important determinants of cell growth and
differentiation[11]. PPARg is present in
various cells, including endothelial cells, vascular smooth muscle cells, monocytes/macrophages, and HSC.
PPARg forms a heterodimer with another nuclear receptor, retinoid X receptor
a (RXRa), and the complex subsequently binds to a specific
DNA sequence designated the peroxisome proliferating response element (PPRE), which is located in the promoter region of
PPARg target genes and modulates their
transcription[12]. It has been demonstrated that depletion of
PPARg accompanies myofibroblastic transdifferentiation of HSC, and
thetreatment of activated HSC in
vitro or in vivo with a variety of
endogenous and exogenous PPARg ligands suppresses the fibrogenic activity of HSC. However, it is uncertain whether
PPARg is indeed a molecular target of this effect, because the ligands are also known to have receptor-independent actions, and the
precise mechanisms of this inhibition have not been
determined[13,14].
Based on the aforementioned information, we postulated that
PPARg might function as a countervailing factor to
attenuate neocollagen formation during hepatic fibrosis, and that CTGF downregulation by
PPARg activation might be one of the mechanisms by which this occurs. To test this hypothesis, we examined the effects of activation of
PPARg on HSC growth and TGF-b1-induced CTGF expression in the present study.
Materials and methods
Reagents Recombinant TGF-b1 was purchased from Sigma (St Louis, MO, USA).
15-deoxy- D12,14-prostaglandin J2
(15-d-PGJ2) was the product of Cayman Chemicals
(Ann Arbor, Michigan, USA). GW7845 and GW9662 were obtained from
GlaxoSmithKline Pharmaceuticals (Brentford, UK). Goat anti-CTGF antibody,
b-actin polyclonal antibody, and horseradish peroxidase-conjugated rabbit anti-goat IgG secondary antibody were purchased from Santa Cruz Biotechnology (Santa Cruz,
CA, USA).
Culture of HSC Rat HSC cells, kindly given to us by
Prof Shi-gang XIONG (Hepatopathy Research Center of California
University, USA), were resuscitated in the routine manner, resuspended with
Dulbecco¡¯s modified Eagle¡¯s medium (DMEM; Gibco BRL, Grand Island, NY, USA)
supplemented with 10 % (v/v) fetal bovine serum (FBS;
Hyclone, USA), 100 kU/mL penicillin G and 100 g/L streptomycin, and then planted in a
25-cm2 culture bottle and incubated in a 5%
CO2 humidified atmosphere at 37
oC. The medium was changed every 3 d and the cells were trypsinized using trypsin/edetic acid when they
reached 80%-90% confluence. HSC aged at passages 4-8 were used for experiments.
Lactate dehydrogenase release assay The cytotoxicity of
PPARg ligands for HSC was evaluated by using the lactate
dehydrogenase (LDH) release assay. The growing HSC were treated with increasing amounts of
15-d-PGJ2 (1, 5, 10, 15, and 20 µmol/L) or GW7845 (0.1, 0.5, 1.0, 1.5, and 2.0 µmol/L) for 48 h . The LDH concentration in conditioned media was measured
as medium LDH, and the LDH concentration in cell lysates was measured as cellular LDH. The concentration of LDH in
DMEM with 10% FBS was defined as contamination arising from FBS and subtracted from the medium and cellular LDH
concentrations. LDH activities were determined by using an LDH assay kit (Sigma). Results are presented as the percentage
of total LDH=medium LDH/(medium LDH+ cellular LDH)×100%.
Cell proliferation assay The status of HSC proliferation was determined by MTT assay (Amresco, USA). Exponentially
growing HSC were adjusted to
2.5×104 cells/mL with DMEM, plated in 96-well plates (Corning, USA) at 200 µL/well and then
incubated for 12 h according to routine procedure. After being treated with
15-d-PGJ2 or GW7845 at various concentrations
and incubated for 48 h (5 duplicate wells for each sample), 20 µL/well MTT (5 g/L) was added to each well. The medium was
then removed after 4 h incubation and 100 µL/well dimethylsulfoxide (M
e2SO) was added to dissolve the reduced
formazan product. Finally, the plate was read in an enzyme-linked immunosorbent microplate reader (Bio-Rad 2550, USA) at
490 nm. The cellular proliferation inhibition rate (CPIR) was calculated using the following formula: CPIR=(1-average A value
of experimental group/average A value of control group)×100%.
Apoptosis assay The effects of PPARg ligands on HSC cell cycle and apoptosis were examined by flow cytometry. In brief,
pretreated HSC were harvested and washed twice with phosphate-buffered saline (PBS) buffer, fixed with 70% ethanol at -20
oC for 30 min and stored at 4
oC overnight, then washed with PBS again, treated with 100 mL 100 mg/L RNase at 37
oC for 30 min and stained with 100 mL 50 mg/L Propidium Iodicle (PI) at 4
oC for 30 min in the dark. The multiplication cycle and apoptotic
rate were assayed using an EPICS XL Flow Cytometer (Coulter, USA) at 488 nm, and the data were analyzed using CellQuest
Software. The percentages of cells in the
G0/G1 phase and S phase, and the apoptotic rate were measured by calculating the
ratio of the number of corresponding cells to the number of total cells. For each sample,
10 000 cells were measured. Morphological changes resulting from apoptosis were determined by electron microscopy.
Measurement of TGF-b1-induced CTGF mRNA expression
RT-PCR was performed according to the protocols
recommended by TaKaRa Bio (Osaka, Japan) with some modifications. RT was performed in a total volume of 25 µL containing 3 µg
total RNA (isolated by TRIzol reagent; Invitrogen, Carlsbad, CA, USA), oligo dT-adaptor primer and Avian Myeloblastosis
Virus (AMV) reverse transcriptase. Primers used for amplification of CTGF cDNA were synthesized by Sangon Gene
Company (Shanghai, China) with reference to the sequence described by Murphy
et al[15]. The sense primer was 5¡¯-CTA AGA CCT
GTG GGA TGG GC-3¡¯ and the antisense primer was 5¡¯-CTC AAA GAT GTC ATT GTC CCC-3¡¯. PCR amplification yielded a PCR
product of 383 bp. Primers used for amplifying a 452 bp of glyceraldehyde phosphate dehydrogenase (GAPDH) cDNA as an
internal control were as follows: 5¡¯-ACC ACA GTC CAT GCC ATC AC-3¡¯ (sense) and 5¡¯-TCC ACC ACC CTG TTG CTG TA-3¡¯
(anti-sense). The PCR conditions were as follows: 30 cycles of denaturation at
94 oC for 30 s, annealing at 55
oC for 30 s, and extension at 72
oC for 60 s. Ten microliters of the PCR product was separated on 1.5% agarose gel, stained with ethidium
bromide (0.5 g/L) and quantitated by densitometry using the Image Master VDS system and associated software (Pharmacia,
USA).
Western blotting analysis Cell culture medium was removed and concentrated with a Biomax Column (Millipore, Bedford,
MA, USA) after TGF-b1 stimulation for 16 h. The adherent HSC were rinsed twice with cold PBS buffer, and were then
harvested and lysed in ice-cold lysis buffer
containing 150 mmol/L NaCl, 50 mmol/L Tris-HCl (pH 7.6),
0.1% sodium dodecylsulfate (SDS), 1% Nonidet P-40, and a protease inhibitor cocktail
(Boehringer Mannheim, Lewes, UK). The samples were cleared by
centrifugation at 13000×g for 10 min. Fifty micrograms of protein from cell lysate or concentrated cell medium was subjected
to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and electrotransferred to polyvinylidene fluoride
(PVDF) membranes (Immobilon, Bedford, MA, USA). After blocking in 20 mmol/L Tris-HCl (pH 7.6; containing 150 mmol/L
NaCl,
0.1% Tween-20, and 5% non-fat dry milk), membranes were incubated with primary antibodies against CTGF or
b-actin (used as a sample loading control) overnight at 4
oC and then incubated with horseradish peroxidase-conjugated secondary
antibody. The blot was developed using the ECL detection kit (Amersham Pharmacia Biotech) according to the manufacturer¡¯s
instructions.
Morphological observations HSC were pre-fixed with glutaraldehyde at the volume fraction of 2.5% and post-fixed with
0.1% osmic acid. Subsequently, the specimens were immersed in propylene oxide after dehydration in an ethanol gradient,
embedded in epoxy resin and made into ultrathin sections. Finally, the sections were stained with lead-uranium and the
changes in ultrastructural organization were observed under a Hitachi-600 (Tokyo, Japan) transmission electron
microscope(Figure 1).
Statistical analysis All data are expressed as mean±SD. Comparisons between groups were carried out using one-way
ANOVA and the Student-Newman-Keuls q test using SPSS 11.0 (SPSS, Chicago, IL, USA).
P values less than 0.05 were considered to be statistically significant.
Results
PPARg ligands inhibited HSC proliferation Both
15-d-PGJ2 and GW7845 significantly inhibited the proliferation of HSC
in a dose-dependent manner (Figure 2), for which the
half effective inhibitory concentrations
(IC50) were 7.38 µmol/L and 0.671 µmol/L, respectively. Correlation analysis indicated a marked positive correlation between the concentration of
15-d-PGJ2 or
GW7845 and CPIR (r=0.898 and r=0.904,
respectively, P<0.01). The cytotoxicity of
PPARg ligands for HSC was carefully studied by examining LDH release. As shown in Table 1,
15-d-PGJ2 and GW7845, compared with the controls, produced no
significant difference in LDH release even at their highest concentrations, respectively. Moreover, after
withdrawal of the PPARg ligands, cell proliferation recovered rapidly. These results indicate that
15-d-PGJ2 and GW7845 are not toxic to cultured HSC. Therefore,
15-d-PGJ2 at 20 µmol/L and GW7845 at 2.0 µmol/L were used for the subsequent experiments.
PPARg ligands caused arrest of cell cycle and induced apoptosis in cultured HSC
Cell cycle analysis of HSC was
carried out by flow cytometry after exposure to various concentrations of
15-d-PGJ2 and GW7845 for 48 h. As shown in
Figure 3A,3B, both 15-d-PGJ2 and GW7845 produced a
dose-dependent increase in the proportion of cells in the
G0/G1 phase and a decrease in the proportion of cells in the S phase. In addition, flow cytometry further demonstrated that
PPARg ligands significantly induced HSC apoptosis compared with the controls (Figure 3C-3E).
Activation of PPARg inhibited TGF-b1-induced
CTGF expression in HSC To understand the biological relevance of
CTGF regulation by PPARg, we investigated whether
PPARg activators regulate TGF-b1-induced CTGF expression in HSC.
The cultured HSC were pretreated with increasing amounts of
15-d-PGJ2 or GW7845 for 1 h and were subsequently stimulated
with TGF-b1 (4 ng/mL) for 4 h. The 4 ng/mL concentration and 4 h incubation time with
TGF-b1 were used because our preliminary
experiments indicated that these conditions yielded optimal activity (data not
shown). The concentration and purity of total RNA were determined by using a DU-800 Spectrophotometer (Beckman Coulter, USA), which determined that
the value of
A260/A280 was between 1.6 and
1.8. Semi-quantitative RT-PCR indicated that both
PPARg activators inhibited TGF-b1-induced CTGF mRNA expression in HSC in a dose-dependent manner (Figure 4).
The effects of 15-d-PGJ2 and GW7845 on CTGF protein levels in
TGF-b1-stimulated HSC were next examined. Consistent
with previous work, we found that CTGF protein was nearly undetectable in untreated HSC by Western blotting analysis,
whereas TGF-b1 significantly induced CTGF
expression. Interestingly, both
15-d-PGJ2 and GW7845 dramatically inhibited
TGF-b1-induced CTGF production and secretion (Figure 5). Taken together, the results indicate that
PPARg activation inhibits TGF-b1-induced CTGF expression at both the mRNA and protein levels. Moreover, the inhibitory effect is more
profound and notable at the protein level than at the mRNA level, suggesting that
PPARg ligands might be exerting some translational or post-translational effects on CTGF expression. In addition, inhibition of CTGF expression was not due to cell
death, because our study had proved that neither
15-d-PGJ2 nor GW7845 was toxic to HSC.
Suppression of CTGF expression was mediated by
PPARg If the suppression of TGF-b1-induced CTGF expression
wasindeed mediated by PPARg, we would expect that the
PPARg-specific and irreversible antagonist GW9662 would abolish this
effect. To test this hypothesis, HSC were pretreated with or without GW9662 (1 µmol/L) for 30 min prior to the addition of
15-d-PGJ2 (20 µmol/L) or GW7845 (2.0 µmol/L) and were subsequently stimulated with
TGF-b1 (4 ng/mL) for 4 h. Semi-quantitative RT-PCR analysis showed that the suppression of
TGF-b1-induced CTGF expression by GW7845 was almost completely
abrogated by GW9662 (relative mRNA level changed from 1.8 to 6.0), whereas the inhibitory effect of
15-d-PGJ2 on CTGF mRNA expression was only partially reversed by GW9662 (relative mRNA level changed from 1.9 to 2.2; Figure 6). These data
indicate that the inhibitory effect of GW7845 on
TGF-b1-induced CTGF expression is largely mediated by
PPARg; however, the inhibitory effect of
15-d-PGJ2 on TGF-b1-induced CTGF expression was only mediated in part through
PPARg, suggesting that 15-d-PGJ2 could also activate some
PPARg-independent signaling pathway to repress CTGF expression in addition to the
activation of PPARg.
PPARg induced a phenotypic switch from activated to quiescent HSC
Unactivated HSC of the control group had typically
shuttle-like or stellate forms and there were no evidently abnormal changes in the nucleus or organelles. The
TGF-b1-activated HSC had enlarged cell volumes, increased amounts and dilation of the rough endoplasmic reticulum, obviously
swollen mitochondria, and no intracellular lipid droplets, which were consistent with the morphological changes of HSC
transdifferentiation. However, after pretreatment with
PPARg ligands, transmission electron microscopy showed that the
rough endoplasmic reticulum and mitochondria were arranged in an orderly fashion, the cell membrane was complete, and
intracellular lipid droplets were basically normal, which indicate that the HSC had changed from the activated to the quiescent
phenotype due to PPARg activation. Moreover, there was obvious chromatin margination and apoptotic bodies in some
HSC, indicating that PPARg ligands could also induce cell apoptosis in activated HSC (Figure 7).
Discussion
The results of the present study demonstrate that
PPARg activation inhibits HSC growth and TGF-b1-induced CTGF
expression. Because activation and proliferation of HSC are critical events in the occurrence and development of hepatic
fibrosis, and CTGF is a key factor in the regulation of extracellular matrix production, these repressive effects by
PPARg activation may be one of the mechanisms by which
PPARg agonists inhibit neocollagen formation during
hepatic fibrosis.
It has been reported that pathological changes in hepatic fibrosis were induced in part by transcription factors that
govern HSC proliferation, death, differentiation, and matrix
production[16]. PPARg is a nuclear receptor transcriptional factor
and plays an important role in many biological processes, including adipogenesis, cell growth regulation, and cell differentiation.
Recent studies have found that thiazolidinediones (TZD) such as pioglitazone and troglitazone, a class of anti-diabetic drugs
that function as synthetic ligands of PPARg with high
affinity, inhibit neointima formation in vascular smooth muscle cells in
association with decreased DNA synthesis, suggesting that
PPARg may be a potential therapeutic target for the treatment of
fibrotic diseases[17-19]. In the present study,
we extended these observations by determining that
PPARg activation results in the inhibition of HSC growth in a dose-dependent manner. Cell cycle status and cell apoptosis are usually closely associated;
cells failing to progress to the mitosis phase are destined for apoptosis. In the past decade, emerging evidence has
suggested that PPARg ligands can induce cell apoptosis in many human malignant tumors, including breast cancer, pituitary
adenomas, colon cancer, pancreatic carcinoma, and esophageal
cancer[20-22]. In addition to cell cycle arrest, the inhibition of
cell growth observed in HSC with PPARg ligands may also be a result of the increase in apoptosis. In the present study,
treatment with PPARg agonists for 48 h caused
G0/G1 phase arrest and blocked cells from entering the S phase. Interestingly,
as seen in other tumor cells, we clearly demonstrated that
PPARg ligands induce significant apoptosis in HSC as evidenced
by flow cytometry and transmission electron microscopy data. However, whether there are differences between normal cells
(ie HSC) and tumor cells with respect to PPARg quantity and intracellular
distribution still remains unclear.
CTGF is a cysteine-rich mitogenic peptide that binds heparin and is secreted by fibroblasts after activation with
TGF-b. CTGF is considered to function as a downstream mediator of
TGF-b action on fibroblastic cells and shares some of the
biological actions of TGF-b, such as the stimulation of cell proliferation and extracellular matrix protein synthesis by
fibroblasts[23]. This concept is strongly supported by the fact that
TGF-b over-production has been documented in nearly every
fibrotic disorder, and CTGF is co-expressed at every site of fibrotic tissue
formation[24]. However, CTGF does not share the
growth inhibitory effect of TGF-b on epithelial cells, or appear to modulate immune or inflammatory
cells[25]. These important distinctions from
TGF-b suggest that CTGF could be a more desirable pharmacological target for the blockade of neocollagen
formation in fibrotic disorders where TGF-b acts as an initiator. In the present study, we demonstrated that CTGF was only
slightly expressed in quiescent HSC, but was dramatically upregulated by
TGF-b1-stimulation, which was in accordance with the results of previous
studies[26]. PPARg activation reduced
TGF-b1-induced CTGF expression at both transcriptional and
post-transcriptional levels, and activated HSC had significant morphological differences in the
activated and quiescent phenotypes, thus the PPARg-CTGF pathway might be a target for a novel antifibrotic strategy.
Recently, it has been reported that there exist
PPARg-independent effects of PPARg ligands at high
doses[27]. In addition, the PPARg natural ligand
15-d-PGJ2 has many functions other than that of a
PPARg activator[28]. In the present study, we
showed that the high affinity PPARg ligand GW7845 could inhibit HSC growth and
TGF-b1-induced CTGF expression. By using the
PPARg-specific antagonist GW9662, we demonstrated that the inhibitory effect of GW7845 on
TGF-b1-induced CTGF expression was mediated by
PPARg. However, the inhibitory effect of
15-d-PGJ2 on TGF-b1-induced CTGF expression
was partly due to PPARg activation, suggesting that
15-d-PGJ2 can activate other PPARg-independent signaling pathways to
repress CTGF expression. Sequence analysis of the CTGF promoter revealed that there are two putative
NF-kB sites, two putative AP-1 sites, and a putative Smads binding element
(SBE). The activated-PPARg-RXRa complex could bind to SBE,
also known as a PPRE, to inhibit CTGF gene
transcription[29,30]. Interestingly, it has been reported that
15-d-PGJ2 inhibits target gene expression not only by interference with Smads, but also by directly inhibiting
NF-kB, from which it could be inferred that the inhibition of
15-d-PGJ2 on TGF-b1-induced CTGF expression was only partly abrogated by
GW9662[31].
In conclusion, our studies demonstrated that
PPARg activation can inhibit HSC proliferation and induce cell apoptosis.
Furthermore, PPARg ligands markedly inhibited
TGF-b1-induced CTGF expression in HSC. Because the biological actions of
TGF-b are complicated and affect many different cell types, whereas CTGF is mainly produced and secreted by activated
HSC, CTGF may serve as a more specific target for selective intervention in processes involving connective tissue formation
during hepatic fibrosis. A better understanding of
PPARg could lead to the development of a new therapeutic approach for
the control and reversion of hepatic fibrosis in humans. However, it should be emphasized that the results in the present
study were generated from cultured HSC and that they might not necessarily and comprehensively reflect the behavior of
quiescent HSC in vivo. Further experiments, beyond the scope of the present study, are required to elucidate the underlying
mechanisms of PPARg in the inhibition of HSC proliferation.
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