1 Laboratory of Hepatobiology and Toxicology, Department of Pharmacology, 2 Curriculum in Toxicology, School of Medicine, and 3 School of Public Health, University of North Carolina, Chapel Hill, North Carolina 27599; and 4 Institut fur Krebsforschung, A-1090 Vienna, Austria
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ABSTRACT |
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Liver regeneration
after partial hepatectomy (PH) involves several signaling
mechanisms including activation of the small GTPases Ras and RhoA in
response to mitogens leading to DNA synthesis and cell proliferation.
Peroxisome proliferator-activated receptor- (PPAR
) regulates the
expression of several key enzymes in isoprenoid synthesis, which are
key events for membrane association of Ras and RhoA. Thus the role of
PPAR
in cell proliferation after PH was tested. After PH, an
increase in PPAR
DNA binding was observed in wild-type mice,
correlating with an increase in the PPAR
-regulated enzyme acyl-CoA
oxidase. In addition, the PPAR
-regulated genes farnesyl
pyrophosphate synthase and 3-hydroxy-3-methylglutaryl-coenzyme A
(HMG-CoA) synthase were significantly increased in wild-type mice. However, these increases were not observed in PPAR
knockout (PPAR
/
) mice. The peak in DNA synthesis observed 42 h
after PH was reduced by ~60% in PPAR
/
mice, despite
increases in TNF-
and IL-1. Also, under these conditions, membrane
association of Ras was high in wild-type mice after PH but was impaired
in PPAR
/
mice. Accordingly, Ras was significantly elevated in the cytosol in PPAR
/
mice. This observation correlated with lower levels of active GTP-bound Ras after PH in PPAR
/
mice compared with wild-type mice. Similar observations were made for RhoA.
Moreover, deletion of PPAR
blunted the activation of
cyclin-dependent kinase (cdk)2/cyclin E and cdk4/cyclin D complexes.
Collectively, these results support the hypothesis that PPAR
is
necessary for cell cycle progression in regenerating mouse liver via
mechanisms involving prenylation of small GTPases Ras and RhoA.
hepatocyte proliferation; cell cycle regulation; RhoA; peroxisome
proliferation-activated receptor-
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INTRODUCTION |
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THE ROLE OF LIPID
AND LIPID metabolism in cell cycle control is not clearly
understood (15, 29, 30). Peroxisome proliferator-activated receptor- (PPAR
) acts as a regulator of lipid metabolism and homeostasis through transcriptional regulation of key enzymes, such as
acyl-CoA oxidase, the initial rate-limiting enzyme in the conversion of
long-chain fatty acids to acyl-CoA thioesters, and
3-hydroxy-3-methylglutaryl-coenzyme A (HMG-CoA) synthase, a
rate-limiting enzyme in the biosynthesis of isoprenoids for protein
isoprenylation (30). This suggests that PPAR
may play a
role in the regulation of prenylation of small GTPases necessary for
cell cycle progression. Importantly, inhibition of the HMG-CoA reductase, an enzyme immediately downstream of HMG-CoA synthase, by
lovastatin completely prevents mesangial cell proliferation in culture
(14). Moreover, treatment of vascular smooth muscle cells
with the HMG-CoA reductase inhibitor simvastatin nearly completely
inhibited PDGF-induced DNA synthesis, retinoblastoma phosphorylation,
and cdk activation (24). PPAR
also regulates the
expression of farnesyl pyrophosphate synthase (FPPS), the terminal
enzyme in FPPS (44). Collectively, these observations support the hypothesis that isoprenoid synthesis via PPAR
-dependent mechanisms is an important factor in regulation of the cell cycle.
Several hypolipidemic drugs, such as WY 14643 and clofibrate,
have gained considerable interest as hepatic peroxisome proliferators and as potent carcinogens in rodent liver. Moreover, PPAR has recently been demonstrated to be necessary for hepatocyte proliferation due to peroxisome proliferators (37). The mechanisms
underlying liver carcinogenesis due to peroxisome proliferators remain
uncertain; however, recent evidence suggests mitogenic cytokines, such
as TNF-
, play key roles (6, 42). Interestingly,
TNF-
-induced hepatocyte proliferation has been shown to involve
critical small GTPases Ras and RhoA (2). PPAR
is a
nuclear transcription factor responsible for the upregulation of a
number of genes involved in peroxisomal
-oxidation, cholesterol and
isoprenoid synthesis (26, 48), yet, it is unclear how
PPAR
plays a role in hepatocyte proliferation.
The two-thirds partial hepatectomy (PH)/liver regeneration model is an
in vivo model in which progression of hepatocyte proliferation occurs
in a relatively synchronous manner and is regulated by a number of
proinflammatory/mitogenic cytokines and growth factors. There is
compelling evidence that TNF- is necessary for liver regeneration in
this model (1, 46). Since PPAR
likely plays a role in
upregulation of isoprenoid synthesis and TNF-
-induced cell
proliferation involves Ras and RhoA, it was hypothesized that PPAR
mediates hepatocyte proliferation in vivo after PH.
Recently, it was reported that the rate of DNA synthesis in
PPAR-deficient mice was not different from controls 72 h after PH (40). Indeed, data presented here demonstrate that no
difference in the rate of proliferation is observed at 72 h.
However, since other studies (1, 47) report a peak DNA
synthesis at 36-44 h in this model, it is possible that
differences in DNA synthesis were overlooked. Thus our studies address
the role of PPAR
in the early stages of DNA synthesis (i.e.,
0-48 h after PH) and, in contrast, clearly demonstrate an
essential role for PPAR
in mechanisms of maximal Ras and RhoA
activation and cell cycle progression.
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MATERIALS AND METHODS |
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Materials. Antibodies used in these studies included the following: cyclin E (Santa Cruz Biotechnology, Santa Cruz, CA), ckd2 (sc-163; Santa Cruz Biotechnology), cdk4 (sc-260; Santa Cruz Biotechnology), phospho-RbT181 (BioSource, Camarillo, CA), p21Cip1 and p27Kip1 (BioSource), RhoA (kind gift of Dr. Channing Der, University of North Carolina), and pan-Ras (OP-40; Chemicon, Temecula, CA). The plasmid including glutathione-S-transferase Ras binding domain of Raf-1 (GST-RBD; binding substrate for activated GTP-bound Ras) was a kind gift of Dr. Adrienne Cox, University of North Carolina. Glutathione Sepharose 4B beads were obtained from Pharmacia (Piscataway, NJ), and protein A/G beads were purchased from Santa Cruz Biotechnology.
Animals.
Mice deficient in the peroxisome proliferator-activated receptor-
(PPAR
/
) and the appropriate SV129 wild-type mice originally characterized and described by Lee et al. (25) have since
been back-crossed more than five generations to C57Bl6 mice. Mice were anesthetized with pentobarbital sodium (60 mg/kg body wt). The left and median lobes of the liver, constituting ~70% of total liver
mass, were removed by the method of Higgins and Anderson (19) through a midabdominal incision. Experimental animals
were allowed to recover for 2-4 h on a 37°C warm plate. All
surgeries were performed between 1,000 and 1,400 h to control for
diurnal variation.
Histochemical analysis and 5-bromo-2-deoxyuridine incorporation. Animals were injected with 100 mg/kg ip 5-bromo-2-deoxyuridine (BrdU; Sigma, St. Louis, MO) 1 h before death. Livers were weighed, fixed in formalin, embedded in paraffin, and sectioned at 6 µm. A section of small intestine, a rapidly proliferating tissue, was collected as a positive control. Unstained slides were deparaffinized in xylene and hydrated in graded concentrations of alcohol and hydrolyzed with 4 N HCL. Sections were further digested with pepsin before incubation with anti-BrdU antibody (DAKO, Carpenteria, CA) diluted 1:200 in 1.0% BSA/PBS. Immunostaining was detected using secondary reagent and diaminobenzidine as recommended by the manufacturer (DAKO). Tissues were counterstained in hematoxylin, followed by dehydration and mounting. Parenchymal cells undergoing DNA synthesis were quantitated by determining the percentage of BrdU-positive nuclei in 10 random high-power fields per slide. BrdU-positive nonparechymal cells were excluded for calculations.
Acyl-CoA oxidase activity.
Activity of the peroxisomal enzyme acyl-CoA oxidase, an accepted
indicator of peroxisome induction (21), was measured from the amount of formaldehyde formed by the peroxidation of methanol by
hydrogen peroxide, a product of peroxisomal -oxidation. Liver samples (~100 µg) were homogenized in 10 volumes of 0.25 M sucrose buffer. A reaction mixture containing (in mg) 13.9 palmitate 3.54 CoA,
68.9 ATP, 20.4 MgCl2, 66.7 NAD+, 45 fatty acid
free BSA, 36.8 semicarbazide, 370 Tris base, 307 Tris · HCl, and 201 niacinamide, and 200 µl
methanol, and 5 µl Triton X-100 (per 50 ml of buffer; pH 8.3) was
warmed to 37°C and 1.4 ml was mixed with 200 µl of liver
homogenate. The reaction was terminated after 10 min with 40% TCA. The
solution was centrifuged to pellet protein, and 1 ml of supernatant was
added to 400 µl of Nash reagent to measure formaldehyde
(34). After the reaction was incubated for 60 min at
37°C, the absorbance was read at 405 nm (
= 6.58). Protein
concentration was determined by the method of Bradford
(8).
Electrophoretic mobility shift assay.
For studies in whole liver, nuclear extracts were isolated as described
by Dignam et al. (12) with minor modifications
(43). The consensus DNA binding oligo for PPAR (Santa
Cruz Biotechnology) was end labeled with [32P-
]ATP
using T4 kinase. Binding conditions for PPAR
were characterized, and
EMSA was performed as described elsewhere (4, 49).
Briefly, nuclear extracts (20 µg) from liver tissue were preincubated
with 1 µg poly(dI-dC), 20 µg BSA (Pharmacia Biotech, Piscataway,
NJ) and 2 µl of a 32P-labeled DNA probe (10,000 counts · min
1 · µl
1,
Cerenkov) containing 1 ng of double-stranded oligonucleotide in a total
volume of 20 µl. Mixtures were incubated 20 min on ice and resolved
on 5% polyacrylamide (29:1 cross-linking) and 0.4×
Tris · HCl/boric acid/EDTA gels. After electrophoresis,
gels were dried and exposed to X-OMAT LS Kodak film. For supershift and
competition assay, anti-PPAR
antibody (4 µg) or unlabeled oligonucleotide was added to the nuclear extract 15 min before incubation with the labeled probe.
Transcription factor array. Biotin-labeled DNA binding oligonucleotides (TranSignal probe mix; Panomics) were incubated with 10 µg of nuclear extract for 1 h to allow the formation of protein/DNA (or transcription factor/DNA) complexes. The protein/DNA complexes were then separated from the free probes by 2% agorase gel electrophoresis. Probes were then extracted from the gel, and ethanol was precipitated, resuspended, and hybridized to the TranSignal Array membrane overnight at 42°C. Membranes were incubated with horseradish peroxidase-labeled streptavidin (DAKO). Detection of signals was obtained using enhanced chemiluminescence imaging system.
RNase protection assay. Total RNA was isolated from liver tissue using RNA STAT 60 (Tel-Test, Friendswood, TX). RNase protection assays were performed using the RiboQuant multiprobe assay system (Becton Dickenson Pharmingen). Briefly, 32P-labeled RNA probes were transcribed with T7 polymerase using the multiprobe template set mCK-3b. RNA (20 µg) was hybridized with 4 × 105 counts/min of probe overnight at 56°C. Samples were then digested with RNase followed by proteinase K treatment, phenol/chloroform extraction, and ethanol precipitation. Samples were resolved on a 5% acrylamide-bisacrylamide (19:1) urea gel. After drying, the gel was visualized by autoradiography.
Cytokine ELISA.
Whole liver from wild-type and PPAR
/
mice was homogenized in
buffer containing 25 mM Tris, 150 mM NaCl, 5 mM EDTA, 0.1% NP-40 and a
cocktail of protease and phosphatase inhibitors. Extracts were used for
ELISA as described by the manufacturer's recommendations.
RT-PCR. cDNA was synthesized from 1.0 µg of total RNA from each sample in a 20-µl final volume of reaction buffer containing (in mM) 25 Tris · HCl (pH 8.3), 37.5 KCl, 10 dithiothereitol, 1.5 MgCl2, 10 of each 2-deoxynucleotide 5'-triphosphate (Perkin Elmer Cetus, Norwalk, CT), and 0.5 mg random hexamer primer (GIBCO-BRL). Samples were incubated for 45 min at 42°C, and the reaction was terminated by denaturing the enzyme at 95°C. The reaction mixture was diluted with distilled water to a final volume of 50 µl. Aliquots (5 µl) of synthesized cDNA were added to 45 µl of PCR mix containing 5 µl of 10 × PCR buffer, 1 µl of each deoxynucleotide (1 mM each), 0.5 µl of sense and antisense primers (0.15 mM), and 0.25 µl (1.25 U) of DNA polymerase (Boehringer-Mannheim). Primers used in these experiments contained the following sequences: FPPS: 5'-AAAATTGGCACTGACATCCAGG-3' (sense), 5'-GGGTGCTGCGTACTGTTCAATG-3' (antisense); HMG-CoA synthase: 5'-TGCCCTGGTAGTTGCAG-3' (sense), 5'-GCCTCTTTCTGCCACT-3' (antisense); glyceraldehyde-3-phosphate dehydrogenase (G3PDH): 5'-TGAAGGTCGGAGTCAACGGATTTGGT-3' (sense), 5'-CATGTGGGCCATGAGGTCCACCAC-3' (antisense). The size of amplified PCR products was 236 bp for FPPS, 291 bp for HMG-CoA synthase, and 983 bp for G3PDH.
The reaction mixture was covered with mineral oil, and amplification of G3PDH was initiated by 1 min of denaturation at 94°C for 1 cycle, followed by multiple cycles (20-35 cycles) at 94°C for 30 s, 58°C for 30 s, and 72°C for 1 min using a GeneAmp PCR system 9800 DNA Thermal Cycler (Perkin Elmer Cetus). After the last cycle of amplification, samples were incubated for 7 min at 72°C. Conditions for the amplification of FPPS and HMG-CoA synthase were identical except 60°C was used for the annealing temperature for FPPS and 55°C for HMG-CoA synthase. The amplified PCR products were subjected to electrophoresis at 75 volts through 2% agarose/ethidium bromide gel for 1 h.Preparation of mouse liver cytosolic and membrane extracts. Livers obtained from mice were homogenized in a buffer containing 10 mM Tris · HCl (pH 7.4), 1 mM EDTA, and 0.25 M sucrose and centrifuged at 10,000 g for 20 min at 4°C. The supernatant was then centrifuged at 100,000 g for 1 h at 4°C to prepare cleared cytosol. The resulting pellet was solubilized with 2% (vol/vol) Triton X-100 for 1 h at 4°C and then centrifuged at 100,000 g for 1 h at 4°C and was used as the membrane extract.
Western blot analysis. Nuclear extracts were separated by SDS-PAGE and transferred to immobilon-P membranes. For cdk2/4 expression, membranes were incubated overnight with anti-cdk2 (1:1,000 dilution, Santa Cruz Biotechnology) or anti-cdk4 (1:1,000 dilution; Santa Cruz Biotechnology) antibodies, followed by 2-h incubation with horseradish peroxidase-conjugated anti-mouse IgG secondary antibody (1:5,000 dilution; Amersham Pharmacia Biotech). Immunoblots were visualized by enhanced chemiluminescence autoradiography. For retinoblastoma (Rb) phosphorylation as a marker for cdk4/cyclin D1 activity, nuclear extracts were separated on a 10% SDS-PAGE gel. Immunoblots were performed using anti-phospho-specific Rb (RbT181; 1:500; BioSource, Camarillo, CA).
For Ras and RhoA localization, membrane, and cytosolic fractions prepared as described above were separated by 16% SDS-PAGE, transferred to immobilon-P membranes, and immunoblotted using antibodies against Ras (1:1,000 dilution) or RhoA (1:1,000 dilution). Gels were subsequently stained with Coomassie blue to control for equal loading.Immunoprecipitation and in vitro kinase reaction. Samples (300 µg protein) in a final volume of 550 µl were incubated in the presence of 1% (vol/vol) Triton X-100 and 0.5% (wt/vol) sodium deoxycholate at 4°C for 1 h. After centrifugation at 10,000 g for 2 min, the supernatant was treated for 30 min with 1 µg of nonimmune IgG. Twenty microliters of a 50% slurry of protein A/G-agarose beads were added, and after 30 min of incubation, the solution was centrifuged at 10,000 g for 2 min. Supernatants were incubated for 2 h with 20 µl of anti-cyclin E antibody at 4°C, after which 50 µl of a 50% slurry of protein A/G-agarose beads were added, and the solution was incubated for a further 1 h. After centrifugation at 10,000 g for 2 min, the pellet was washed three times with 1 ml of wash buffer (50 mM HEPES, pH 7.4, containing 1% Triton X-100, 0.1% SDS, 150 mM NaCl, 100 mM NaF, and 2 mM sodium orthovanadate).
Kinases from immunoprecipitated-associated complexes were then assayed by the transfer of phosphate from [Expression of GST fusion proteins and pull-down assay. pGEX plasmids, including GST-RBD (binding substrate for activated GTP-bound Ras), were a kind gift from Dr. Adrienne Cox, University of North Carolina. The expression of the GST-RBD (Ras binding domain of Raf-1) fusion protein was performed as described previously (39). Briefly, 10 ml of an overnight culture of bacteria transformed with expression plasmid was inoculated into 100 ml of LB medium containing 50 µg of ampicillin/ml. The culture was placed on a shaking incubator at 37°C until an absorbance of 1 at 600 nm was reached. Production of recombinant protein was induced with 1 mM (final concentration) isopropylthiol-d-galactoside (GIBCO-BRL), and the culture was incubated for a further 3-4 h at room temperature. Cells were harvested by centrifugation and resuspended in 5 ml of 1% Triton X-100-PBS and 1 mM PMSF. The suspension was incubated on ice for 30 min before cells were lysed by sonication. Bacterial lysate was centrifuged at 12,000 g for 20 min at 4°C to remove the insoluble fraction. The GST-RBD was purified using glutathione-beads (Amersham Pharmacia Biotech). Purified GST-RBD-glutathione complex was incubated overnight with 500 µg of whole liver extract prepared as described above. The resulting precipitate was washed and resuspended in 2× Laemlli sample buffer and separated by 16% SDS-PAGE. Immunoblot was performed using anti-pan Ras antibody (1:1,000; Chemicon).
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RESULTS |
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PPAR is involved in liver regeneration after PH.
Rodent hepatocyte proliferation induced by peroxisome proliferators has
been shown to require both TNF-
and PPAR
(6, 7). The
PH model is an established in vivo model of synchronized hepatocyte
proliferation that requires TNF-
and other mitogenic cytokines
(1, 46). Thus it was hypothesized that liver regeneration and hepatocyte proliferation after PH would also require PPAR
. To
test this hypothesis, two-thirds PH was performed on wild-type and
PPAR
/
mice. DNA synthesis was determined in livers from wild-type and PPAR
/
mice by measuring BrdU incorporation at several time points between 0 and 72 h after resection (Fig.
1). Since it is likely that hepatocytes
and nonparenchymal cells, such as bile duct epithelia, are undergoing
mitosis after PH, only hepatocytes were included in the determination
of cell proliferation. Wild-type mice had a significant increase in
BrdU incorporation within 24 h after PH, which peaked around
42 h (Fig. 1A), consistent with other reports
(47). In contrast, DNA synthesis was significantly blunted
by 60-65% in PPAR
/
mice at 42 and 48 h after PH
(P < 0.05, repeated measures ANOVA) (Fig.
1B). However, 72 h after PH, the rate of DNA synthesis
was not significantly different between wild-type and PPAR
/
mice, confirming a previous report (40). To test whether
PPAR
was involved in production of TNF-
and other mitogenic
cytokines after PH, RNase protection assays were performed on mRNA
isolated from wild-type and PPAR
/
mice from 0 to 72 h
after resection (Fig. 2). TNF-
and
IL-6 mRNA levels peaked between 12 and 24 h after PH and were not
different between wild-type and PPAR
/
mice.
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PPAR is necessary for induction of key enzymes involved in
isoprenoid synthesis.
PPAR
is a transcription factor activated by a number of endogenous
ligands including fatty acids leading to upregulation of enzymes
involved in fat metabolism as well as sterol and isoprenoid synthesis
(16, 22, 23). To first test the hypothesis that PPAR
is
activated after PH, an electrophoretic mobility shift assay was
performed to evaluate PPAR
DNA binding activity after PH. With the
use of nuclear extracts from wild-type mice, a significant increase in
PPAR
DNA binding activity was observed 24 and 48 h after PH
(Fig. 4A). As expected, this
increase was not observed in nuclear extracts from PPAR
/
mice.
To validate the specificity of PPAR
DNA binding, nuclear
extracts from livers after PH were compared with nuclear extracts
isolated from livers of mice treated with WY 14643, classical agonist
of PPAR
(Fig. 4B). Twenty-four hours after 100 mg/kg WY
14643 treatment, a significant increase in PPAR
DNA binding was
observed compared with vehicle-treated mice. The PPAR
DNA binding
from animals after PH, although slightly less intense than after WY
14643 exposure, was observed. Competition assay using excess unlabeled
DNA oligo blunted PPAR
DNA binding and incubation with anti-PPAR
antibody caused a shift in the mobility, confirming the translocation
and activation of PPAR
after PH in mice. Importantly, a
time-dependent increase in the activity of the classical
PPAR
-regulated gene acyl-CoA oxidase was observed after PH; acyl-CoA
oxidase activity was significantly blunted in PPAR
/
mice (Fig.
5A). These data are consistent with the activation of PPAR
after PH and support a potential role
for PPAR
in liver regeneration.
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PPAR is necessary for activation and membrane localization of
Ras after PH.
Initiation of G1/S phase transition after PH is regulated
in part by small GTPases Ras and RhoA through their activation of cdk (32). The hypothesis is that PPAR
is involved in
the regulation of isoprenoid synthesis and therefore may influence
prenylation and function of small GTPases, such as Ras and RhoA.
Specifically, it is hypothesized that Ras and RhoA are concentrated in
the cytosol in livers from PPAR
/
mice rather than in the
membranes in which they are normally localized. To test this
hypothesis, membrane and cytosolic extracts from livers of both
wild-type and PPAR
/
mice were immunoblotted for Ras and RhoA at
several time points after PH (Fig.
6A). Prior to PH, the ratio of
membrane-bound to cytosolic Ras was nearly equal in wild-type mice,
whereas Ras was largely concentrated in the cytosol in PPAR
/
mice. After PH, the relative levels of Ras in the membrane vs. cytosol
were not significantly different in wild-type mice for
48 h. However, the amount of Ras in membrane extracts of PPAR
/
livers was decreased significantly compared with the amount of Ras in the cytosol
after PH. Relative changes of RhoA in the membrane compared with
cytosol were similar to Ras. The amount of membrane-bound RhoA was
higher than the levels of cytosolic RhoA before PH and was not
significantly different for
48 h after PH in wild-type animals. In
contrast, in PPAR
/
mice, membrane-bound RhoA was significantly
lower than RhoA in the cytosol initially and was barely detectable at
any time point studied after PH.
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PPAR plays an essential role in cell cycle regulation.
The hypothesis that PPAR
is necessary for regulation of the cell
cycle was tested by evaluating critical parameters of early G1/S phase transition of the cell cycle. Cdk2 and -4 complex with cyclin E and cyclin D1, respectively, to trigger
G1 to S phase transition of the cell cycle
(31). The expression of both cdk2 and cdk4 was evaluated
by Western blot in nuclear extracts from wild-type and PPAR
/
animals 0-42 h after PH. Expression of cdk2 was minimal; however,
it was increased dramatically 24 and 42 h after PH in wild-type
mice (Fig. 7A). Importantly,
this increase in cdk2 expression did not occur in livers from PPAR
/
mice at any time after PH. Similarly, an increase in cdk4 was
also observed, which peaked 24 h after PH in wild-type mice;
increases in cdk4 were not detectable in livers from PPAR
/
mice. Expression of cyclin E and D1 were also measured by Western blot
and were not changed under these conditions (data not shown). These
findings are consistent with a recent report that overexpression of
constitutively active Ras enhances liver regeneration after PH without
a significant increase in cyclin D1 expression (28).
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DISCUSSION |
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Role of PPAR in isoprenoid synthesis.
It is well established that numerous genes encoding peroxisomal,
mitochondrial, and microsomal enzymes contain functional PPAR
-responsive elements in their promoter regions
(22). PPAR
-responsive genes are largely involved in
lipid metabolism and
-oxidation of fatty acids but also include
enzymes involved in isoprenoid synthesis, such as HMG-CoA synthase and
the terminal enzyme FPPS (5, 44). Although it is clear the
PPAR
participates in regulation, these genes are also controlled by
a variety of other transcription factors as well. For example, there
are PPAR, retinoid X receptor, Sp1, and CREB cis-element binding sites
within HMG-CoA synthase promoter, which may control basal and inducible
gene expression (18). Indeed, it is demonstrated in Fig.
5 that PPAR
activity required for maximal upregulation of
both FPPS and HMG-CoA synthase in vivo after PH. Basal expression and
the modest increase in gene expression in the PPAR
/
mice after
PH may be due to activation of other regulatory elements. It is also
well documented that inhibition of HMG-CoA reductase with statins
blunts G1/S phase progression in response to a number of
growth stimuli in cultured cells (17). Since HMG-CoA
reductase and synthase are in a linear pathway for sterol and
isoprenoid synthesis, these observations are consistent with the
hypothesis that these synthetic pathways are central to normal cell
cycle and proliferative function. As mentioned above, inhibition of
HMG-CoA reductase blunts cell proliferation in vitro and depletes
lipid-modified Ras and RhoA from membranes (14, 24). Ras
and RhoA have each been shown to play a pivotal role in early
progression of the cell cycle (27, 35). Furthermore, inhibition of cell growth and proliferation by farnesyl transferase inhibitors is dependent on the inhibition of prenylation and
membrane-association of small GTPases (11). Thus it was
hypothesized that activation of PPAR
plays a key role in the
synthesis of farnesyl and geranylgeranyl pyrophosphate, which are
required for the association of Ras and RhoA to membranes. Auer et al.
(2) reported that mitogen-induced cell proliferation in
hepatocytes required activation of Ras. It was shown here that membrane
association of both Ras and RhoA is diminished in PPAR
/
mice
compared with wild-type mice under normal conditions as well as at
several time points after PH (Fig. 6). Importantly, the impairment in
Ras membrane association is reflected as a decrease in Ras activity
after PH (Fig. 1B). These data indeed support the hypothesis
that PPAR
is required for lipid-modification and membrane
association of Ras and RhoA. These data are consistent with other
reports (24) showing that inhibition of HMG-CoA reductase
by mevinolin reduces the level of membrane-bound Ras and Ras-mediated
signal transduction in cell culture. However, this is the first report
linking the PPAR
to small G protein function in vivo.
Regulation of cell proliferation by small GTPases.
Mitogens, through activation of Ras and RhoA, stimulate the progression
of hepatocytes from G1 into S phase by activating of both
cdk4/cyclin D and cdk2/cyclin E activities (3). Recently, it was demonstrated that activation of RhoA or overexpression of
geranylgeranylated RhoA facilitates cell cycle progression by
stimulating p27Kip1 degradation (20, 35, 36).
Cyclin/cdk complexes are inhibited by specific inhibitor proteins
p21Cip and p27Kip1, which are highly expressed
in most cells and serve as negative regulators of mitogen-induced,
cell-cycle progression (9, 41). Previously, it was shown
that activation of the Ras/Rho but not the Ras/Erk pathway is required
for p27Kip degradation and G1 progression
(45). Loss of p21 and p27 is difficult to accurately
determine in vivo since their expression rapidly changes within 24 h of PH. It is demonstrated here that a very transient decline in p21
and p27 levels occurs after PH in wild-type mice but not in PPAR
/
mice (Fig. 7). Additionally, the increase in cdk2/cyclin E and
cdk4/cyclin D activities in wild-type mice (Fig. 7) are correlated with
the loss of p21 and p27. Ras and RhoA are required for full activation
of cdk2/cyclin E and cdk4/cyclin D and degradation of p21 and p27. Very
recently it was demonstrated that the overexpression of
dominant-negative Ras using adenovirus blunted cell proliferation after
PH in rats (28), clearly supporting a role for Ras in
regulation hepatic regeneration. Since PPAR
/
mice have
significantly less membrane-associated Ras and RhoA as well as lower
levels of activated Ras after PH (Fig. 6), it is concluded that PPAR
is necessary for maximal Ras and RhoA-mediated cell cycle progression.
Role of PPAR in cell proliferation.
PPAR
was recently shown to be required for hepatocyte proliferation
and tumorigenesis due to a variety of lipophilic peroxisome proliferators, such as di(2-ethylhexyl) phthalate, and WY 14643 (33). It was also shown that peroxisome
proliferator-induced cell proliferation was dependent on activation of
Kupffer cells and production of mitogenic cytokines (42).
Interestingly, PPAR
does not seem to play a direct role in cytokine
production (Figs. 2 and 3), consistent with the finding that Kupffer
cells, which are the major source of TNF-
in the liver
(10), do not express PPAR
(38). Therefore,
it is possible that there is a requirement for both Kupffer cell
mitogen production and PPAR
for peroxisome proliferator-induced
hepatocyte proliferation. The two-thirds PH/liver regeneration model is
an in vivo model in which progression of hepatocyte proliferation
occurs in a relatively synchronous manner and is regulated by a number
of proinflammatory, mitogenic cytokines and growth factors
(13). Indeed, there is compelling evidence that TNF-
is
necessary for liver regeneration (1). It has been shown
that livers from mice deficient in TNF-
receptors do not regenerate
after PH compared with wild-type mice (47). The data are
consistent with other reports that TNF-
and other mitogenic
cytokines are rapidly induced after PH (Figs. 2 and 3). Despite
increases in the mitogenic signals, liver regeneration after PH was
blunted in mice deficient in PPAR
(Fig. 1). Loss of PPAR
does not
seem to impact liver size or hepatocyte cell number under normal
conditions based on liver morphology in these studies and other reports
(25).
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FOOTNOTES |
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Address for reprint requests and other correspondence: M. D. Wheeler, Univ. of North Carolina at Chapel Hill, CB# 7365, 3013 Thurston-Bowles Bldg, Chapel Hill, NC 27599 (E-mail: wheelmi{at}med.unc.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
First published October 16, 2002;10.1152/ajpgi.00175.2002
Received 14 May 2002; accepted in final form 7 October 2002.
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