Male Kunming mice (age, 6 weeks; weight, 22–25 g; total n=40) were obtained from the Shanghai SLAC Laboratory Animal Co., Ltd. (Shanghai, China) and 5 mice/cage were maintained under a 12-h light/dark schedule, with a temperature range of 20 to 24°C. Experiments were conducted according to protocols and guidelines approved by the Longyan University Institutional Animal Care and Use Committee (Longyan, China). Liquid diets were based on the modified Lieber-DeCarli formulation and provided 1 kcal/ml (prepared by Trophic Animal Feed High-tech Co., Ltd., Nantong, China). The ethanol diet consisted of 18% of the total energy as protein, 35% as fat, 19% as carbohydrate and 28% as ethanol. In the control diet, ethanol was replaced isocalorically with carbohydrate. Following a 1-week period of adaptation to the environment with free access to the liquid diet, mice were randomly divided into four experimental groups (n=10 mice/group): Control diet-fed mice; control diet-fed mice + zopolrestat (zopol); ethanol diet-fed mice and ethanol diet-fed mice + zopol. Mice were administered 50 mg/kg/day zopol as a single daily intraperitoneal injection for 4 weeks. Equivalent volumes of saline were administered to the control groups of mice. On the final day, mice were sacrificed under anesthesia. Tissues were snap-frozen in liquid nitrogen immediately following resection and stored at −80°C until further analysis.

Liver tissues were fixed in 10% formalin for 24 h at room temperature and were routinely embedded in paraffin. Briefly, tissues were rinsed in tap water for 5 min at room temperature prior to dehydration with a series of alcohol (70, 80 and 90%, respectively, for 5 min each, followed by 3 incubations with 100% alcohol for 5 min each at room temperature). The tissues were then treated twice with xylene for 5 min each at room temperature, prior to paraffin embedding. Sections (5-µm) were deparaffinized in xylene twice for 5 min each and treated with 100% alcohol twice for 3 min, then 95, 70 and 50% alcohol, respectively, for 3 min each at room temperature. Sections were stained with hematoxylin and eosin (H&E) for histological analysis. Hepatic steatosis was graded as previously described (10), according to the percentage of lipid-laden hepatocytes: 0, 0%; 1, 0–25%; 2, 26–50%; 3, 51–75% and 4, 76–100%.

AML12 mouse hepatocyte cells were obtained from the American Type Culture Collection (Manassas, VA, USA) and cultured in Dulbecco's modified Eagle's medium/Ham's F12 containing 100 U/ml penicillin, 100 µg/ml streptomycin and 10% fetal bovine serum (Hyclone; GE Healthcare Life Sciences, Logan, UT, USA), in a 37°C incubator with 5% CO₂.

AML12 cells were plated on 24-well plates at a density of ~1×10⁵ cells/well. Following 24 h of incubation, cells were exposed to 100 mM ethanol and co-treated with 50 µM zopol. A total of 36 h subsequent to ethanol and zopol treatment, ROS generation in cells was determined using ROS assay kit (cat no S0033; Beyotime Institute of Biotechnology, Haimen, China), according to the manufacturer's instructions.

Total lipoperoxides were measured as thiobarbituric acid reactive substances (TBARS) in liver homogenates or cell lysates, using a lipid peroxidation malondialdehyde (MDA) assay kit (cat no S0131; Beyotime Institute of Biotechnology), according to the manufacturer's instructions. TBARS were quantified using MDA as a standard and Microsoft Excel 2013 (Microsoft Corporation. Redmond, WA, USA).

Total RNA was isolated from tissues and cells using TRIpure reagent (Roche Applied Science, Mannheim, Germany), according to the manufacturer's protocol. cDNA was synthesized from mRNA using RevertAid First Strand cDNA Synthesis kits (Thermo Fisher Scientific, Inc., Waltham, MA, USA). Expression levels of hepatic TNF-α, TGF-β1, interleukin (IL)-6 and sterol regulatory element binding protein (SREBP)-1c were analyzed. The specific primers used were: Forward, 5′-CGTGCTCCTCACCCACAC-3′ and reverse, 5′-GGGTTCATACCAGGGTTTGA-3′ for TNF-α; forward, 5′-ACAACCACGGCCTTCCCTACTT-3′ and reverse, 5′-GTGTAATTAAGCCTCCGACT-3′ for IL-6; forward, 5′-CAACTTCTGTCTGGGACCCT-3′ and reverse, 5′-TAGTAGACGATGGGCAGTGG-3′ for TGF-β1; forward, 5′-ATCGGCGCGGAAGCTGTCGGGGTAGCGTC-3′ and reverse, 5′-ACTGTCTTGGTTGTTGATGAGCTGGAGCAT-3′ for SREBP-1c and forward, 5′-CTATTGGCAACGAGCGGTTCC-3′ and reverse, 5′-GCACTGTGTTGGCATAGAGGTC-3′ for β-actin. qPCR was performed using the FastStart Universal SYBR-Green Master Mix (Roche Applied Science). The reaction was performed with the following thermocycling conditions: 95°C for 10 min, followed by 35 cycles of 95°C for 15 sec, 53–59°C for 30 sec and 72°C for 30 sec, then a final extension of 72°C for 10 min. The ^2-ΔΔCq method (11) was used for quantification and the data were normalized to β-actin.

Tissues and cells were homogenized in ice-cold radioimmunoprecipitation assay buffer (Beyotime Institute of Biotechnology). Protein concentrations of the extracts were measured using a bicinchoninic acid protein assay kit (Beyotime Institute of Biotechnology), according to the manufacturer's protocol. A total of 40 µg of each protein sample was loaded and separated using 10–12% SDS-PAGE and transferred to polyvinylidene difluoride membranes (EMD Millipore, Billerica, MA, USA). Membranes were blotted with 5% non-fat milk in TBS containing 0.1% Tween-20 for 1 h at room temperature. Membranes were then incubated with anti-AR (cat no. sc-17735; 1:500; Santa Cruz Biotechnology, Inc., Dallas, TX, USA), anti-cytochrome P450 2E1 (cat no. ab28146; 1:1,000; CYP2E1; Abcam, Cambridge, UK), anti-phosphorylated AMPK (cat no. 2535; 1:1,000), anti-AMPK (cat no. 2532; 1:1,000) (both from Cell Signaling Technology), anti-GAPDH (cat no. G8795; 1:2,000) or anti-β-actin (cat no. A2228; 1:2,000) (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) at 4°C overnight, and subsequently with horseradish peroxidase-conjugated anti-goat or anti-rabbit immunoglobulin G (cat nos. AP307P and AB324P, respectively; 1:2,000; Sigma-Aldrich; Merck KGaA) for 2 h at room temperature. Detection was achieved using the SuperSignal Chemiluminescent Substrate kit (Beyotime Institute of Biotechnology).

All data were processed and analyzed using GraphPad Prism software, version 5.0 (GraphPad Software, Inc., La Jolla, CA, USA) and were expressed as the mean ± standard error of the mean. One-way analysis of variance and the post hoc Newman-Keuls multiple comparison test was used for multi-group analyses. P<0.05 was considered to indicate a statistically significant difference.

Previous studies demonstrated that feeding C57BL/6 mice with Lieber-DeCarli ethanol diets induced fatty liver within 4–8 weeks (12). In order to investigate whether AR was involved in the development of ethanol diet-induced hepatosteatosis, the protein expression levels of hepatic AR were analyzed in mice fed with the ethanol diet. As presented in Fig. 1A, hepatic AR protein expression in C57BL/6 mice fed with the ethanol diet for 4 weeks was increased, compared with mice fed with the control diet. In addition to the induction of AR expression in mouse liver tissue, an increase in AR protein expression was observed in hepatocytes exposed to 100 mM ethanol for 36 h (Fig. 1B).

In order to further determine the role of AR in the development of ethanol-induced hepatosteatosis, AR activity was inhibited by treating mice with an AR-specific inhibitor, zopol. As presented in Fig. 2A and B, examination of H&E-stained sections demonstrated marked hepatic steatosis in mice fed with the ethanol diet for 4 weeks, while mice fed with the control diet exhibited low-grade steatosis. Following inhibition of AR activity, hepatic steatosis in mice fed with the ethanol diet was significantly attenuated (P<0.05).

In order to clarify the mechanism through which AR is associated with the development of ethanol-induced steatosis, the effect of AR inhibition on hepatic lipoperoxide production was investigated. As presented in Fig. 3A, consumption of the ethanol diet resulted in an increase in hepatic TBARS levels, compared with the control diet, and the increase was attenuated significantly by treating the ethanol diet-fed mice with zopol (P<0.001). The protein expression of CYP2E1, a principal mediator of lipid peroxidation (13,14), was induced by ethanol, and AR inhibition significantly prevented the induction of CYP2E1 (Fig. 3B). In addition, the protective effect of AR inhibition on ethanol-induced oxidative stress was analyzed in vitro. As presented in Fig. 3C and D, inhibition of AR resulted in a 33.0% decrease in MDA levels (P<0.001) and a 33.9% decrease in ROS levels (P<0.05) in ethanol-treated AML12 cells. However, CYP2E1 protein expression was not detected in AML12 cells. The results of the present study demonstrated that the induction of AR by ethanol may promote hepatic oxidative stress in mice with alcoholic liver disease, and that the increased hepatic oxidative stress is due in part to the AR-mediated induction of CYP2E1.

Previous studies demonstrated that AMPK inactivation was associated with the development of ethanol-induced hepatic steatosis (15,16). In order to clarify the mechanisms whereby AR affects lipid accumulation in hepatocytes, the effect of AR inhibition on AMPK activity was investigated. As presented in Fig. 4A, phosphorylated AMPK protein expression levels in the livers of ethanol-fed mice treated with zopol were increased compared with ethanol-fed mice, indicating that AR inhibition activated hepatic AMPK in ethanol-fed mice. The activation on AMPK by the AR inhibitor was additionally confirmed in ethanol-treated AML12 cells (Fig. 4B). The effect of AR inhibition on the mRNA expression of SREBP-1c, a transcriptional factor regulated by AMPK which controls the synthesis of fatty acids, was assessed in the livers of ethanol-fed mice. As presented in Fig. 4C, AR inhibition did not significantly decrease the mRNA expression levels of SREBP-1c in ethanol-fed mice when compared with the ethanol-fed group. However, AR inhibition significantly decreased the mRNA expression levels of SREBP-1c in ethanol-treated AML12 cells (Fig. 4D).

In order to evaluate the effect of AR inhibition on the development of ethanol-induced liver injuries, the expression levels of certain proinflammatory cytokines were assessed, including TNF-α, IL-6 and TGF-β1. Mice fed on the ethanol diet exhibited a marked elevation of hepatic mRNA expression of TNF-α compared with mice fed on the control diet (Fig. 5A). However, the ethanol-induced TNF-α mRNA elevation was significantly attenuated by 51.4% (P<0.01) in mice treated with zopol. Compared with TNF-α, the effect of AR inhibition on IL-6 and TGF-β1 was less significant. Consistent with the in vivo results, AR inhibition in ethanol-treated AML12 cells resulted in a significant decrease in the mRNA expression levels of TNF-α (P<0.01; Fig. 5B). AR inhibition in ethanol-treated AML12 cells resulted in a significant decrease in the mRNA expression levels of IL-6 (P<0.05) and TGF-β1 (P<0.05). The results of the present study demonstrated that hepatic AR elevation in mice fed on the ethanol diet may directly affect the production of pro-inflammatory cytokines, particularly TNF-α, in the liver.