Restoration of retinoic acid concentration supresses ethanol-enhanced c-Jun expression and hepatocyte proliferation in rat liver

Jayong Chung, Chun Liu, Donald E. Smith, Helmut K. Seitz1,, Robert M. Russell and Xiang-Dong Wang2,

Gastrointestinal Nutrition Laboratory, Jean Mayer USDA Human Nutrition Research Center on Aging at Tufts University, 711 Washington Street, Boston, MA 02111, USA and
1 Department of Medicine, Salem Medical Center, Heidelberg, Germany


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Chronic and excessive ethanol intake decreases hepatic retinoic acid (RA) concentrations, which may play a critical role in ethanol-induced hyperproliferation in hepatocytes. The present study was conducted to determine whether RA supplementation in chronic ethanol-fed rats could restore hepatic RA concentrations to normal levels and modulate hepatocyte hyperproliferation. Male Sprague–Dawley rats were divided into four groups: control, ethanol-fed, ethanol-fed + 50 µg all-trans-RA/kg body wt and ethanol-fed + 100 µg all-trans-RA/kg body wt. Ethanol was given to rats at 6.2% (v/v) in a liquid diet to provide 36% of total caloric intake. Control animals received the same amount of liquid diet with isocaloric maltodextrin in place of ethanol. Results show that the ethanol treatment in rats for a month significantly increased the mean number of proliferating cell nuclear antigen (PCNA)-positive hepatocytes [4.96 ± 1.36% (ethanol-fed) versus 0.29 ± 0.08% (control), P < 0.05]. This increase was associated with the induction of hepatic c-Jun protein (6.5-fold increase) and cyclin D1 protein (3-fold increase) in ethanol-fed animals as compared with controls. Furthermore, activator protein 1 (AP-1) DNA-binding activity was significantly higher in hepatic nuclear extracts from ethanol-fed rats than those from controls. In contrast, RA supplementation in ethanol-fed rats raised hepatic RA concentration to normal levels and almost completely abolished the ethanol-enhanced c-Jun, cyclin D and AP-1 DNA-binding activities. Moreover, RA supplementation at both doses markedly suppressed the ethanol-induced PCNA-positive hepatocytes by ~80%. These results demonstrate that the restoration of hepatic RA concentrations by dietary RA supplementation suppresses ethanol-induced hepatocyte proliferation via inhibiting c-Jun overexpression, and suggest that RA may play a role in preventing or reversing certain types of ethanol-induced liver injury.

Abbreviations: AP-1, activator protein 1; HPLC, high performance liquid chromatography; JNK, c-Jun N-terminal kinase; PCNA, proliferating cell nuclear antigen; RA, retinoic acid.


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Ethanol is the world's most widely abused addictive drug and is associated with an increased incidence of a variety of illnesses, including cancer (1). Although a number of epidemiological studies have indicated that chronic and excessive alcohol consumption is a significant risk factor for certain cancers (liver, esophagus, oropharynx, colorectal, breast, etc.), the mechanisms by which ethanol ingestion promotes carcinogenesis are not well understood. Several mechanisms have been proposed, such as induction of microsomal cytochrome P-450 enzymes that activate procarcinogens (2), generation of acetaldehyde (3) and reactive free radicals (4,5), formation of DNA strand breaks (6), impairment of the liver's ability to metabolize dietary nitrosamines (7) and impairment of nutritional status (8). Chronic ethanol feeding to experimental animals increases hepatocyte proliferation (911), which plays an integral role in both the promotion and progression stages of multistage hepatocarcinogenesis (1214). The proliferative response induced by ethanol intake promotes genomic instability and neoplasia; however, it is not known if specific dietary derived compounds would have protective effects against this type of ethanol-related liver response.

Retinoic acid (RA), the most active form of vitamin A, plays an important role in controlling the progression to early carcinogenesis in a variety of cancers (15,16). Retinoic acid is currently used for treatment of several types of cancers, including acute promyelocytic leukemia, squamous cell carcinoma of the head and neck, oral and cervical premalignant lesions and skin cancer (15,16). However, chronic and excessive ethanol intake reduces vitamin A levels in liver tissue in alcoholics (17). Moreover, we observed that both plasma and hepatic RA concentrations are significantly decreased in ethanol-fed rats after 1 month of treatment (18). The observation that retinoid (retinol and retinyl esters) concentrations in hepatic cancerous tissues are significantly lower than those in the surrounding non-cancerous tissues of alcoholic patients suggests a role for retinoid depletion in hepatocarcinogenesis (19). One of the chemoprotective effects of retinoids is thought to be mediated through the control of cell proliferation by delaying the progression of damaged cells into S phase, thus enabling enhanced DNA repair with an associated reduction of the risk of carcinogenic initiation and promotion. It has been reported that after treatment with polyprenoic acid (an acyclic retinoid) there is a significant reduction in the incidence of second primary tumors in patients who have previously undergone hepatoma resection (20). In the recent follow-up analysis (62 months), a significant difference in survival (74 versus 46%, P = 0.04) was also found (21). Moreover, Baba et al. (22) showed that RA inhibited N-nitrosomorpholine induction of hepatic cell proliferation and carcinogenesis in rats; however, the mechanisms involved are uncertain.

We recently reported that chronic ethanol intake in rats increases hepatic c-Jun protein levels as compared with control animals (18). c-Jun, the cellular homologue of v-Jun oncogene in avian sarcoma virus 17 (23) and a component of activator protein 1 (AP-1), is known to play a critical role in cell cycle progression. A causal role of c-Jun in promoting cell proliferation was suggested by studies using microinjection of antibodies or antisense RNA directed against c-Jun, which showed a failure of progression from G1 into S phase (2426). Conversely, cell cycle distribution in cells overexpressing c-Jun is shifted toward S phase (27). It has been reported (28) that c-Jun is required for progression through the G1 phase of the cell cycle by a mechanism that involves direct transcriptional control of the cyclin D1 gene, which is a major player in cell proliferation. c-jun knock-out in mice is lethal and primary fibroblasts from the c-jun null embryos are completely defective in their proliferation (29). Recent studies have shown that all-trans-RA can suppress c-Jun expression in lung cancer cells (30) and in human skin after ultraviolet irradiation (31). However, it is not known whether the up-regulation of c-Jun by ethanol is an important mechanism for ethanol-induced hepatocyte proliferation, and whether RA supplementation can inhibit ethanol-induced c-Jun overexpression and the resultant hepatocyte proliferation.

In the present study, we demonstrate that dietary RA supplementation in ethanol-fed rats can restore normal hepatic RA concentrations and inhibit ethanol-induced c-Jun and cyclin D1 overexpression and hepatocyte hyperproliferation.


    Materials and methods
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Animals and diets
Twenty-four male Sprague–Dawley rats (130–150 g; Charles River Breeding Laboratory, Kingston, NY) were individually housed in stainless steel suspended rodent cages and provided free access to Teklad 7012 rodent meal (Harlan Teklad, Madison, WI) and water for a 1 week acclimation period. Subsequently, all animals were fed a Lieber–DeCarli liquid diet (Dyets, Bethlehem, PA) without ethanol for 5 days. Animals were then distributed, by weight-matching, into four groups: (i) control group (C); (ii) ethanol group (E); (iii) ethanol group with dietary all-trans-RA supplementation (50 µg/kg body wt) (ERA50); and (iv) ethanol group with dietary all-trans-RA supplementation (100 µg/kg body wt) (ERA100). Ethanol was fed with the Lieber–DeCarli liquid diet containing 36% of total calories as ethanol, yielding a concentration of 6.2% (v/v). Ethanol was gradually introduced into the experimental diets over a 10 day period before providing animals with the final concentration. Control animals were maintained on the Lieber–DeCarli liquid diet without ethanol. In the control diet, ethanol was replaced by an isocaloric amount of maltodextrin (Purina Test diets, Richmond, IN). Both diets contained 16.4% of total calories as protein and 35% as fat; 48.6% of total calories were provided from carbohydrates in the control diet, whereas 12.6% of total calories were from carbohydrates in the ethanol diet. The Lieber–DeCarli liquid diet contains vitamin A (0.9 mg/1000 kcal), which is a sufficient amount for both the control and experiment groups (32). There was no RA in the base liquid diet, as analyzed by HPLC. For dietary RA supplementation, doses of 50 or 100 µg/kg body weight all-trans RA (Sigma, St Louis, MO) were given. All-trans-RA was dissolved in 95% ethanol and added directly into the liquid diet. Since the liquid diet provides physiological amounts of fluid, extra water was not given. One animal from each group was matched by body weight at the beginning of the study. Rats were pair-fed using ERA100 as a leading group for 1 month. Body weights were recorded weekly. At the end of the experimental period, all rats were terminally exsanguinated under AErrane® anesthesia (Fort Dodge Animal Health, Fort Dodge, IO). Liver tissues were collected, frozen under liquid nitrogen and stored at –80°C for further analysis.

All animals were maintained in an American Association for the Accreditation of Laboratory Animal Care accredited facility, in an environmentally controlled atmosphere (temperature, 23°C; relative humidity, 45%) with 15 air changes of 100% fresh filtered air per hour and a 12/12 h light/dark cycle. All animals were observed daily for clinical signs of illness. This project was approved by the Jean Mayer USDA Human Nutrition Research Center on Aging Animal Care and Use Committee.

HPLC analysis of retinoid concentration in liver tissue
Liver sample extractions were done as described by Wang et al. (18). Briefly, liver tissue was homogenized with an ice-cold 2:1 (v/v) HEPES buffer:methanol mixture. After adding the internal standard (100 µl retinyl acetate), retinoids were extracted twice without saponification using 6.0 ml 2:1 (v/v) chloroform:methanol mixture. The extracts were evaporated under N2 gas and resuspended in 50 µl ethanol for injection into the HPLC system. A gradient reverse phase HPLC system was used as follows: flow rate was 1 ml/min; 10% solvent A [50:20:30 (v/v/v) acetonitrile:tetrahydrofuran:water, with 0.35% acetic acid and 1% ammonium acetate in water] for 3 min, followed by a 6 min linear gradient to 40% solvent A and 60% solvent B [50:44:6 (v/v/v) acetonitrile:tetrahydrofuran:water, with 0.35% acetic acid and 1% ammonium acetate in water], a 12 min hold at 40% solvent A/60% solvent B, and then a 7 min gradient back to 100% solvent A. Individual retinoids were identified by co-elution with standards and absorption spectral analysis. Retinoids were quantified relative to the internal standard by determining peak areas calibrated against known amounts of standards.

Immunohistochemistry
Hepatocytes in S phase were quantified by immunohistochemical analysis of proliferating cell nuclear antigen (PCNA). Liver samples were fixed with 10% buffered formalin and embedded in paraffin wax. Five micrometer sections were cut using a microtome. After a standard deparaffinization–rehydration procedure, liver sections were incubated with 0.3% H2O2 in methanol for 30 min at 37°C to quench endogenous peroxidase activity. The sections were heated using microwaves for 5 min in 10 mM sodium citrate buffer (pH 6.0) to retrieve antigen. The routine biotin–streptavidin immunohistochemical method consisted of sequential incubations in horse serum blocking solution, monoclonal anti-PCNA (clone PC10; Dako, Carpinteria, CA), biotinylated horse anti-mouse immunoglobulin G and streptavidin conjugated to a horseradish peroxidase label. The slides were developed with diaminobenzidine substrate and counterstained with hematoxylin. Sections were examined under light microscopy by two independent investigators who were blinded as to treatment groups. Only hepatocytes with dark brown stained nuclei and cytoplasm were counted as S-phase cells. A total of 40 fields (about 250 hepatocytes in each field) were screened per liver sample. Positive S-phase cells were expressed per 100 hepatocytes.

Western blot analysis
Nuclear extracts from rat livers were prepared as described by Wang et al. (18). Briefly, liver tissues were homogenized with ice-cold buffer A (10 mM Tris–HCl pH 7.5, 10% glycerol, 10 mM KCl, 10 mM monothioglycerol and a mixture of protease inhibitors) and centrifuged at 2000 g at 4°C. Nuclear pellets were solubilized in buffer B (10 mM Tris–HCl pH 7.5, 10% glycerol, 600 mM KCl, 10 mM monothioglycerol and 1 mM dithiothreitol) and vortexed for 15 min. The extracts were centrifuged at 100 000 g for 30 min at 4°C. Protein concentrations of the nuclear extracts (supernatants) were determined using a Coomassie Plus protein assay kit (Pierce, Rockford, IL). Nuclear protein (20 µg) in a sample buffer containing 50 mM Tris–HCl (pH 6.8), 200 mM 2-mercaptoethanol, 2% SDS, 0.1% bromophenol blue, 10% glycerol and the mixture of protease inhibitors were boiled for 6 min and run on 10% SDS–PAGE. The protein was transferred to a polyvinylidene difluoride membrane by using a semi-dry transfer system. Membranes were incubated overnight with 5% non-fat milk in Tris-buffered saline containing 0.05% Tween 20. Membranes were incubated with primary antibodies diluted in Tris-buffered solution for 1 h at room temperature. Primary antibodies (c-Jun and Cyclin D1) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The blots were washed and incubated with horseradish peroxidase-labeled secondary antibody (Bio-Rad, Hercules, CA) for 30 min and washed for 30 min. The blots were developed using the ECL Western Blotting system (Amersham, Piscataway, NJ) and analyzed using a densitometer (GS-710 Calibrated Imaging Densitometer; Bio-Rad).

RT–PCR
Total RNA was extracted from liver tissue using an RNA purification kit (Promega, Madison, WI). Total RNA (2 µg) from liver tissues was used for reverse transcription in a reaction mixture of 25 mM MgCl2, 5 mM KCl, 10 mM Tris–HCl (pH 7.5), 1 mM dNTP, 1 U/µl RNase inhibitor, 2.5 U/µl Moloney murine leukemia virus reverse transcriptase and 2.5 µM random hexamers. PCR was performed on the cDNA product in each tube using primers for c-jun from Clontech (Palo Alto, CA). PCR products from primer sets for GAPDH were used as controls. PCR was performed in a conventional PCR mixture (1.5 mM MgCl2, 5 mM KCl, 10 mM Tris–HCl, 2.5 U Taq DNA polymerase and 0.2 µM each primers) using a thermal cycler (Perkin-Elmer, Norwalk, CT). PCR consisted of 10 initial 5 min incubations at 95°C, followed by 30 amplification cycles (95°C for 1 min, 58°C for 1 min, and 72°C for 3 min) and a final extension step (72°C for 10 min). The cycle number of PCR within the range of linear amplification was determined for each gene (30 cycles for c-jun, 24 cycles for GAPDH). Equal volumes of the PCR product from each sample were subjected to electrophoresis on a 1.8% agarose gel stained with ethidium bromide and photographed. The sizes of PCR products (c-Jun and GAPDH were 432 and 316 bp, respectively) were compared with a {phi}X174/Hae III DNA size marker (Gibco, Gaithersburg, MD). c-Jun mRNA was determined after correcting for cDNA loading, based on GAPDH intensity.

Electrophoretic mobility-shift assays (EMSA)
Electrophoretic mobility-shift assays were performed on nuclear extracts from rat liver. Nuclear protein extract (15 µg) was incubated with a reaction mixture [20 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 100 mM KCl, 0.2 mM EDTA, 2 mM dithiothreitol, 20% glycerol, 200 µg/ml poly(dI–dC), pH 7.6] in the absence or presence of a 20-fold molar excess of a double-stranded oligonucleotide with the consensus sequence for AP-1. The sequence of the AP-1 oligonucleotide is 5'-GTAAAGCATGAGTCAGACACC-3' (33). After 10 min incubation, 2 ng 32P-labeled double-stranded AP-1 oligonucleotide was added, and incubation was continued for an additional 20 min. The DNA–protein complexes were resolved by electrophoresis through a 6% non-denaturing polyacrylamide gel in 0.5x TBE [45 mM tris(hydroxymethyl) aminomethane, 45 mM boric acid and 1 mM EDTA]. Gels were dried and subjected to autoradiography at –80°C. The specificity of the retarded AP-1 complexes was confirmed by using supershift assays incubating the reaction mixture with an antibody against c-Jun for 30 min at 25°C.

Statistical analysis
All group values are expressed as means ± SEM. Group means were compared using ANOVA followed by Tukey's honest significant difference test. Values of P < 0.05 were considered significant.


    Results
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
Animal body weights in the control group and the ethanol-fed groups with or without RA supplementation were the same at the start of the study. After 1 month of treatment, there were no significant differences in body weights due to supplementation with RA at doses of 50 and 100 µg/kg body wt, and body weight gains were similar in all ethanol-fed groups. Ethanol-fed rats that were supplemented with RA reached slightly higher body weights compared with ethanol-fed rats without RA supplementation [249 ± 5 (ethanol-fed), 255 ± 8 (ethanol-fed + 50 µg RA/kg body wt) and 260 ± 7 g (ethanol-fed + 100 µg RA/kg body wt)], but the differences were not significant. Despite pair-feeding, ethanol-fed rats gained less body weight than control rats (291 ± 7 g) during the study period.

Table IGo shows that, after 1 month of ethanol feeding, levels of hepatic RA were significantly reduced to half of that found in control rats. The concentrations of retinol and retinyl palmitate in the liver were also decreased (84 and 73%, respectively) in ethanol-fed rats as compared with control rats. However, ethanol-induced hepatic RA depletion was prevented by RA supplementation. Compared with rats that were fed with ethanol alone, RA supplementation at two different doses (50 and 100 µg/kg body wt) in ethanol-fed rats restored hepatic RA levels. RA supplementation at this range of doses did not elevate hepatic RA concentrations above the normal values that were found in control rats. There were no significant differences in hepatic RA concentrations among control versus ethanol-fed rats with RA supplementation (Table IGo).


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Table I. Effects of chronic ethanol feeding and retinoic acid supplementation for 1 month on retinoid concentrations in rat liver (n = 6, each group)
 
All ethanol-fed rats had significantly lower hepatic retinyl palmitate levels compared with those in the control group, and RA supplementation did not alter the hepatic concentration of retinyl palmitate. However, RA supplementation caused a significant increase in hepatic retinol levels (Table IGo). Both levels of RA supplementation in ethanol-fed rats significantly increased the hepatic retinol concentration by 2-fold compared with control rats (P < 0.05), although these changes were not dose-dependent.

The effect of ethanol feeding and RA supplementation on cell proliferation was assessed by immunohistochemical staining for PCNA in liver sections from each group (Figures 1 and 2GoGo). The PCNA-positive hepatocytes in liver sections from ethanol-fed rats increased dramatically after 1 month of treatment. The presence of fatty liver was also apparent in liver sections of the ethanol-fed rats (Figure 1Go). The mean number of PCNA positive cells was significantly increased by 17-fold in livers from ethanol-fed rats after 1 month of treatment (P < 0.05). In contrast, liver sections from rats given retinoic acid with ethanol had a significantly lower number of PCNA-labeled hepatocytes than those given only ethanol in their diet. Although the average number of PCNA-positive hepatocytes in ethanol-fed rats with RA supplementation was slightly higher than that in control rats (Figure 2Go), the differences were not significant (P > 0.05). The presence of fatty liver was also apparent microscopically in liver sections from the ethanol-fed rats with or without RA supplementation (Figure 1Go).



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Fig. 1. Immunohistochemical staining for proliferating cell nuclear antigen (PCNA) in representative liver sections. Only hepatocytes with uniformly dark brown stained nuclei were counted as S-phase cells (original magnification x400). (A) Control, (B) ethanol fed and (C) ethanol fed with retinoic acid supplementation (100 µg/kg body wt).

 


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Fig. 2. Effect of retinoic acid supplementation for 1 month on hepatocyte proliferation in ethanol-fed rats. Hepatocytes in S-phase were determined by immunohistochemistry of PCNA on formalin-fixed paraffin-embedded liver tissue. A total of 40 fields (about 10 000 hepatocytes) were screened per animal and examined under light microscopy. Means ± SEM (n = 6, each group). Treatments not sharing the same superscript are significantly different from each other (P < 0.05).

 
Levels of both c-Jun and cyclin D1 protein expressions were examined by using western blot analysis of nuclear protein extracts from liver tissues in each group. The hepatic level of c-Jun protein was 6.5-fold greater in rats fed the ethanol diet than in rats fed the control diet (Figure 3Go). However, RA supplementation completely inhibited the overexpression of c-Jun in liver tissue caused by chronic ethanol exposure (Figure 3Go). Further, c-Jun mRNA levels were determined by RT–PCR analysis of total RNA in the liver of control, ethanol-fed and ethanol-fed + RA-treated rats. The result showed that ethanol feeding in rats increased c-Jun mRNA level, as compared with the non-ethanol-fed rats (Figure 4Go). However, this ethanol-induced elevation of c-Jun mRNA was inhibited by RA treatment in the ethanol-fed rats (Figure 4Go). Similar to c-Jun expression, the hepatic level of cyclin D1 proteins in ethanol-fed rats increased 3-fold, while RA supplementation inhibited cyclin D1 overexpression (Figure 5Go).



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Fig. 3. Effect of retinoic acid supplementation for 1 month on hepatic c-Jun protein levels in ethanol-fed rats. Western blotting analysis was followed by densitometric analysis. Densitometric units were normalized using control values in the respective pair-fed group. Means ± SEM (n = 6, each group). Inset, representative western blot. C, Control; E, ethanol-fed; E+RA (50), ethanol-fed + RA supplementation (50 µg/kg body wt); E+RA (100), ethanol-fed + RA supplementation (100 µg/kg body wt). Bottom, the membrane was stripped of the antibody against c-Jun and then reprobed with antibody against RAR{alpha} (18), as a control for equal amounts of protein loading.

 


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Fig. 4. Effect of retinoic acid supplementation on c-Jun mRNA level, as determined by RT–PCR analysis of total RNA, in the liver of ethanol-fed rats. Inset, ethidium bromide-stained agarose gel of the c-Jun RT–PCR product amplified from liver of control rats (C), ethanol-fed rats (E) and ethanol-fed, RA-treated rats (E+RA). RT–PCR products using primers for GAPDH were used as controls.

 


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Fig. 5. Effect of retinoic acid supplementation for 1 month on hepatic cyclin D1 protein levels in ethanol-fed rats. Western blotting analysis was followed by densitometric analysis. Densitometric units were normalized using control values in the respective pair-fed group. Means ± SEM (n = 6, each group). Inset, representative western blot. C, Control; E, ethanol-fed; E+RA (50), ethanol-fed + RA supplementation (50 µg/kg body wt); E+RA (100), ethanol-fed + RA supplementation (100 µg/kg body wt).

 
We further compared AP-1 DNA-binding activities of nuclear protein extracts from livers of different groups by using electrophoretic mobility shift assays (Figure 6Go). AP-1 binding activities in ethanol-fed rats were enhanced significantly, as indicated by a stronger intensity of the shifted band as compared with control rats (Figure 6Go). On the contrary, nuclear extracts from ethanol-fed rats that were supplemented with retinoic acid showed clearly reduced binding activity to AP-1. Both a competitive assay with the addition of unlabeled probe in excess and a supershift assay with c-Jun monoclonal antibody verified the specificity of the shifted band (Figure 6Go).



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Fig. 6. Effect of retinoic acid supplementation for 1 month on hepatic AP-1 binding activity in ethanol-fed rats. (Left) Electrophoretic mobility-shift analysis was performed on nuclear extracts from liver. C, Control; E, ethanol-fed; E+RA (50), ethanol-fed + RA supplementation (50 µg/kg body wt); E+RA (100), ethanol-fed + RA supplementation (100 µg/kg body wt). (Middle) Electrophoretic mobility-shift analysis was performed on hepatic nuclear extracts for competitive assay with the addition of unlabeled probe in excess. The specific protein–DNA complex was displaced by a 20-fold molar excess of unlabeled oligonucleotide with the consensus sequence for AP-1 (5'-GTAAAGCATGAGTCAGACACC-3') as competitor. (Right) The specificity of AP-1 binding activity was tested by a supershift assay with monoclonal antibody against c-Jun. The incubation with c-Jun antibody resulted in the appearance of a further shifted band, as indicated by an arrow.

 

    Discussion
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 Abstract
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 Materials and methods
 Results
 Discussion
 References
 
Increased hepatocyte proliferation in the rats after long-term and excessive ethanol feeding has been demonstrated by several investigators using either immunohistochemistry for PCNA and/or 5-bromo-2-deoxyuridine as a marker of the proliferation of hepatocytes (911). In the present study, chronic ethanol feeding of rats for 1 month increased hepatocyte proliferation by 17-fold (evaluated by PCNA labeling) as compared with a control group (Figures 1 and 2GoGo), which is similar to previous reports (911). However, the possible mechanism(s) involved in ethanol-induced hepatocyte proliferation are not well defined. We hypothesized that the overexpression of c-Jun and cyclin D1 by chronic and excessive ethanol intake may cause abnormal cell cycle regulation and drive cells into premature S phase. This hypothesis is supported by our findings that ethanol feeding for 1 month increased both hepatic c-Jun (6.5-fold; Figure 3Go) and cyclin D1 proteins (3-fold; Figure 5Go), which were associated with an increased mean number of PCNA-positive hepatocytes (Figures 1 and 2GoGo) as compared with the control group. Since the cyclin D1 gene contains an AP-1 response element in its promoter region (28), increased cyclin D1 could be a consequence of enhanced c-Jun protein levels in ethanol-fed rat livers. This is supported by the results from our gel shift assay showing that the DNA binding activity of AP-1 was significantly increased in nuclear extracts from liver tissues of ethanol-fed rats (Figure 5Go). The levels of c-Jun and cyclin D1 proteins in proliferated hepatocytes could be even higher, since whole hepatic tissues were used for nuclear protein extraction in this study. In order to rule out a possibility that these changes were due to ethanol-induced activation of hepatic stellate cells, we isolated hepatic stellate cells from rat liver after 1 month of ethanol treatment and did not observe activation of the isolated stellate cells (assessed by collagen type I expression, PCNA and c-Jun gene expressions; J.Chung and X.D.Wang, unpublished data). In contrast to our finding, it has been reported that ethanol impairs the early proliferative response of hepatocytes induced by partial (2/3) hepatotectomy (3436). The explanation for these discrepant findings may be due to different models used in these studies. For example, major surgery accompanies vastly different hormonal milieu as well as altered metabolism.

In this experiment, high-dose ethanol treatment of rats for 1 month resulted in a significant reduction of retinyl ester content of the liver, as compared with control animals which were pair-fed an isocaloric control diet containing the same amount of vitamin A (Table IGo). Similar to our previous report (18), the hepatic RA concentration was significantly lower in the ethanol-treated animals as compared with the pair-fed control animals. There could be several mechanisms underlying the reduction in hepatic RA as a result of ethanol feeding (37), including (i) ethanol acts as a competitive inhibitor of retinol oxidation in liver; (ii) ethanol increases mobilization of vitamin A from hepatic tissue to peripheral tissue and (iii) ethanol enhances degradation of RA to more polar metabolites. Recently, we have shown that enhancement of catabolism of RA to 18-hydroxy-RA and 4-oxo-RA in ethanol-fed rats can be inhibited by both specific inhibitors and antibody against cytochrome P4502E1 (38), thus offering a biochemical mechanism for the reduction of hepatic retinoic acid concentrations observed with chronic ethanol consumption in vivo (Table IGo).

In order to test our hypothesis that RA supplementation can modulate ethanol-induced hepatocellular proliferation by inhibition of ethanol-induced c-Jun expression, we first determined what dosage of RA supplementation in ethanol-fed rats would restore normal tissue RA concentrations. In our preliminary study, RA supplementation at the dose of 300 µg/ kg body weight raised hepatic RA concentration above that of normal levels and doubled the plasma RA concentration compared with control animals, while an RA dose of 100 µg/kg body wt restored hepatic and plasma RA concentrations similar to those of controls (data not shown). To avoid potential RA toxicity, we selected supplementation of RA (50 and 100 µg/kg body wt) (Table IGo). These RA intakes did not induce any changes in animal weight, although ethanol-fed rats gained less body weight than control rats at the end of the study period, which is consistent with earlier reports (32,39).

It is interesting that RA supplementation increased hepatic concentrations of retinol but not retinyl ester in ethanol-fed rats (Table IGo). Previous studies (40,41) in non-ethanol fed rats have shown that the addition of RA to the vitamin A deficient diet resulted in an increase in the hepatic retinyl ester concentration. Although the mechanism(s) of this `sparing effect' of RA is not well defined, it has been shown that RA inhibits retinol oxidation and stimulates retinol esterification (42,43). Recently, Zolfaghari and Ross (44) have demonstrated that RA rapidly induces lecithin:retinol acyltransferase mRNA in the livers of vitamin A-deficient mice and rats, indicating that RA serves as an important signal for regulating hepatic vitamin A metabolism. However, in the present study, the increase of retinol in the liver of ethanol-fed rats after being given RA may be due to both a feedback inhibition of retinol oxidation by RA (42), and an impairment of the esterification of retinol in the liver of ethanol-fed rats, as reported in a previous study (45). The exact mechanism(s) by which the levels of retinol in the livers of ethanol-fed rats with RA treatment become elevated needs further study.

The most important observations of this study are identification of (i) c-Jun and cyclin D1 as two potential targets of retinoid action in ethanol-fed rats and (ii) the role of RA in regulation of ethanol-induced hepatocyte proliferation. We demonstrate that the overexpression of c-Jun in chronic ethanol-fed rats can be inhibited by RA supplementation at both the mRNA and protein level in vivo (Figures 3 and 4GoGo). The level of c-Jun protein in liver tissue was inversely associated with hepatic RA concentrations (Table IGo), implying a causal relationship between diminished retinoid action and hepatocyte hyperproliferation by ethanol. This hypothesis is further supported by the fact that the RA supplementation at both doses inhibited ethanol-enhanced expression of cyclin D1 (Figure 5Go), AP-1 DNA-binding activity (Figure 6Go), as well as hepatocyte proliferation (Figures 1 and 2GoGo). The suppression of c-Jun by RA in ethanol-fed rats could be through inhibition of c-Jun N-terminal kinase phosphorylation (30), post-transcriptional regulation of c-Jun (31) or competition for a limited pool of cAMP response element-binding protein (CBP; a co-activator for both nuclear retinoid receptors and c-Jun transcription factor) (46). Recently, we demonstrated that the inhibition of ethanol-enhanced hepatic c-Jun expression by RA is through a mechanism of suppressing c-Jun N-terminal kinase-dependent signaling pathways (J.Chung, R.M.Russell and X.D.Wang, in preparation).

A number of epidemiological and experimental studies have indicated that chronic and excessive ethanol consumption is a significant risk factor for cancer. While ethanol by itself is not carcinogenic, chronic ethanol feeding to experimental animals does result in impaired nutritional status of retinoids and increased hepatic cell proliferation, which may provide a promoting environment for carcinogenesis. Our results demonstrate that the maintenance of normal RA levels by RA supplementation in ethanol-fed rats inhibits hepatocyte hyperproliferation. This study provides important information on the possible role of RA in preventing or reversing certain types of ethanol-induced liver injury.


    Notes
 
2 To whom correspondence should be addressed Email: xwang{at}hnrc.tufts.edu Back


    Acknowledgments
 
We thank Drs H.J.Palmer and Y.J.Suzuki for their helpful advice. This material is based upon work supported by grant R01CA49195 from the National Institutes of Health (Bethesda, MD) and by the US Department of Agriculture, under agreement No. 1950-51000-048-01A. Any opinions, findings, conclusions or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the views of the US Department of Agriculture.


    References
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

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Received November 8, 2000; revised February 28, 2001; accepted March 7, 2001.