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Abstract

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

We tested the hypothesis that the pathogenesis of alcoholic liver injury is mediated by epigenetic changes in regulatory genes that result from the induction of aberrant methionine metabolism by ethanol feeding. Five-month-old cystathionine beta synthase heterozygous and wild-type C57BL/6J littermate mice were fed liquid control or ethanol diets by intragastric infusion for 4 weeks. Both ethanol-fed groups showed typical histopathology of alcoholic steatohepatitis, with reduction in liver S-adenosylmethionine (SAM), elevation in liver S-adenosylhomocysteine (SAH), and reduction in the SAM/SAH ratio with interactions of ethanol and genotype effects. Hepatic endoplasmic reticulum stress signals including glucose-regulated protein-78 (GRP78), activating transcription factor 4, growth arrest and DNA damage-inducible gene 153 (GADD153), caspase 12, and transcription factor sterol response element binding protein-1c (SREBP-1c) were up-regulated in ethanol-fed mice with genotype interactions and negative correlations with the SAM/SAH ratio. Immunohistochemical staining showed reduction in trimethylated histone H3 lysine-9 (3meH3K9) protein levels in centrilobular regions in both ethanol groups, with no changes in trimethylated histone H3 lysine-4 levels. The chromatin immunoprecipitation assay revealed a decrease in levels of suppressor chromatin marker 3meH3K9 in the promoter regions of GRP78, SREBP-1c, and GADD153 in ethanol-treated heterozygous cystathionine beta synthase mice. The messenger RNA expression of the histone H3K9 methyltransferase EHMT2 (G9a) was selectively decreased in ethanol-fed mice. Conclusion: The pathogenesis of alcoholic steatohepatitis is mediated in part through the effects of altered methionine metabolism on epigenetic regulation of pathways of endoplasmic reticulum stress relating to apoptosis and lipogenesis. (HEPATOLOGY 2009.)

Previous studies established associations of abnormal hepatic methionine metabolism with the development and clinical expression of alcoholic steatohepatitis (ASH).1, 2 In transmethylation reactions, homocysteine is methylated to methionine and then S-adenosylmethionine (SAM), which is a substrate and principal methyl donor in methylation reactions, whereas S-adenosylhomocysteine (SAH) is both a product and potent inhibitor of methylation reactions.3 Therefore, the SAM/SAH ratio is considered a useful expression of methylation capacity.2 SAH is also the substrate for SAH hydrolase, a reversible reaction that generates homocysteine in the forward direction, but increases SAH when homocysteine is in excess. In transsulfuration reactions, homocysteine is a substrate for the cystathionine beta synthase (CβS) reaction, which is facilitated by SAM to generate cystathionine and ultimately glutathione (GSH), the principal antioxidant in the liver.4

Our prior studies in ethanol-fed micropigs linked elevated liver homocysteine and SAH levels to endoplasmic reticulum (ER) stress.5 In mice fed intragastric ethanol, betaine prevented hepatic lipid accumulation and hepatocellular apoptosis by lowering homocysteine and SAH levels.6 Feeding ethanol to micropigs with a folate-deficient diet accelerated the onset and severity of ASH while increasing liver homocysteine and SAH and reducing SAM and the SAM/SAH ratio.1 ER stress signals including nuclear sterol response element binding protein 1-c (SREBP-1c) and its target lipogenic enzymes acetyl-coenzyme A carboxylase and fatty acid synthase were increased in livers of pigs fed ethanol and folate-deficient diets, and their levels correlated positively with liver homocysteine, SAH, and the SAM/SAH ratio.5 Subsequent micropig studies showed that SAM supplementation of ethanol diets prevented the pathology of ASH by correcting the SAM/SAH ratio and inhibiting expressions of SREBP-1c and its target lipogenic genes7 and pathways of oxidative liver injury.8

In ER dysfunction, the accumulation of unfolded proteins triggers a series of events referred to as the unfolded protein response. Key components of this response in mammals involve activated ER membrane transducers including PKR-like ER kinase, activating transcription factor 6 (ATF6) and inositol-required enzyme 1.9 Activation of ATF6 leads to increased expression of ER chaperones, including glucose-regulated protein-78 (GRP78) that may be involved in repair.10 Up-regulation of PKR-like ER kinase also increases activating transcription factor 4 (ATF4) and growth arrest and DNA damage-inducible gene 153 (GADD153), a transcription factor for apoptosis. A different ER stress-induced apoptotic pathway involves procaspase 12, which is activated by its cleavage during ER stress.11 ER-resident transcription factor SREBP-1c plays an important role in lipogenesis during prolonged unfolded protein response.12

Epigenetic mechanisms of DNA methylation and histone modification affect gene transcription through chromatin remodeling. Histone modifications include histone H3 lysine acetylation in promoter regions of active genes and histone H3 lysine methylation, which is associated with gene activation or repression depending on the methylation site.13 Recent studies showed that lysine methylation is a key modulator for transcriptional activation or repression. For example, trimethylated histone H3 lysine-4 (3meH3K4) occurs mainly at the transcription start sites of active genes, whereas trimethylated histone H3 lysine-9 (3meH3K9) is associated with gene repression.14 Histone H3K9 methyltransferases that catalyze these modifications include Suv39h1 (KMT1A), which mediates the trimethylation of H3K9 to 3meH3K9, EHMT2 (G9a), which mediates the dimethylation of H3K9 to 2meH3K9, SUV39h2, and Setdb1 (ESET).15, 16 Becaue SAM, the principal methyl donor, and SAH, the principal inhibitor of methylation reactions closely regulate all methylation reactions,3 it seemed likely that ethanol-induced changes in SAM and SAH would result in altered histone methylation in this mouse model.

The goal of the present study was to define the mechanistic role of aberrant hepatic methionine metabolism in the pathogenesis of ASH in a genetically altered intragastric ethanol-fed mouse model and to determine the role of altered epigenetic regulation. We studied the effects of reduced liver SAM and elevated SAH on epigenetic regulation of ASH in the heterozygous CβS-deficient mouse, which is known to have ≈50% enzyme activity with elevated plasma homocysteine and normal growth, survival, and liver histology.17 While ethanol feeding predictably lowers SAM levels by inhibiting the transmethylation of homocysteine,4 CβS deficiency inhibits the transsulfuration of homocysteine,4, 17 and their combination would predictably elevate liver SAH through the reversible SAH hydrolase reaction. The net reduction in the SAM/SAH methylation ratio could affect the epigenetic regulation of genes relevant to liver injury. Establishing interactive effects of ethanol feeding and CβS heterozygosity on liver injury would further implicate aberrant methionine metabolism in the pathogenesis of alcoholic liver disease.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Animals and Diets.

Five-month-old CβS wild-type (+/+) and heterozygous (+/−) littermate mice on a C57BL/6J background (n = 22) were obtained from the breeding colony at the University of Iowa,18 and were grouped to receive a control or ethanol-containing diet by intragastric infusion for 4 weeks using an established method.19 The mouse intragastric ethanol infusion model was provided by the Animal Core of the University of Southern California Research Center for Alcoholic Liver and Pancreatic Diseases. All animals received human care according to criteria outlined in the Guide for the Care and Use of Laboratory Animals of the National Academy of Sciences. After 1 week of infusion of a control diet, ethanol infusion was initiated at 22.7 g/kg/day and incrementally increased to 33 g/kg/day (37.1% of kcal) over 4 weeks. At the initial ethanol dose, total calories derived from the diet was set at 568 kcal/kg/day, and the caloric percentages of ethanol, dextrose, protein, and fat (corn oil) were 29%, 11%, 25%, and 35%, respectively. Vitamins, salts, and trace minerals were included at the recommended amounts by the Committee on Animal Nutrition of the National Research Council (Dyets Inc, Bethlehem, PA). After 4 weeks of intragastric feeding, plasma was removed for measurements of ethanol, aspartate aminotransferase (AST), and alanine aminotransferase (ALT) levels. A portion of each liver specimen was fixed in 10% formalin for 2 hours, then placed in 80% ethanol and sent to S.W. French at Harbor/UCLA Medical Center for histological processing and evaluation. The remainder of each liver specimen was snap-frozen and sent to the University of California Davis for further studies.

Liver Methionine Metabolite Measurements.

Liver SAM, SAH, and GSH levels were measured by high-performance liquid chromatography coulometric electrochemical detection.20

Liver Chemistry and Histopathology.

AST and ALT were measured in terminal plasma as conventional markers of liver injury. Liver histopathology included quantitative scoring of appropriately stained slides, which were evaluated in blinded fashion using computerized software and scored according to published criteria for microscopic and macroscopic hepatocyte lipid accumulation, inflammation, necrosis, fibrosis, and mitochondrial alterations.21

Terminal Deoxynucleotidyl Transferase–Mediated dUTP Nick-End Labeling Assay.

Apoptotic bodies in liver specimens were detected by DNA fragmentation using terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL).22 Apoptotic nuclei in hepatocytes were counted in 10 fields in each liver sample to obtain average values for each sample as numbers of TUNEL-positive cells per mm2.

Immunohistochemistry of Methylated Histones.

Liver tissue was fixed in neutral buffered formalin, embedded in paraffin, cut into 4-μg-thick sections, stained with a rabbit polyclonal antibody to 3meH3K9 or 3meH3K4, each at 1/100 titer (Epitomics, Burlington, CA), followed by Donkey fluorescein isothiocyanate (FITC) labeled antibody 1/100 titer (Jackson ImmunoReaserch Labs Inc.,Westgrove, PA). The intensity of nuclear fluorescence was quantified and blinded to treatments and mice identity using a FITC filter and Nikon morphometrics software with a Nikon 400 fluorescent microscope 40× objective with the same sensitivity setting throughout.23 Centrilobular and periportal peripheral hepatocyte nuclei were analyzed separately.

RNA Isolation, Complementary DNA Synthesis, and Relative Quantitative Reverse-Transcription Polymerase Chain Reaction.

Total RNA was isolated from frozen liver specimens using the RNeasy total RNA kit (Qiagen, Valencia, CA). Reverse transcription was performed using 2 μg of DNase-treated RNA following the protocol provided in the first-strand complementary DNA (cDNA) synthesis kit (Invitrogen, Calsbad, CA). The primers for the mouse cDNA sequence were designed using the Primer Express program (Version 2, Applied Biosystems, Foster City, CA). β-Actin was used as an internal control, and each reaction was performed in triplicate using the ABI Prism 7900 sequence detection system (Applied Biosystems, Foster City, CA). Separate standard curve cDNA dilutions were included in each polymerase chain reaction (PCR) run. Liver transcripts were normalized to β-actin levels. The primer pairs for each gene are shown in Supporting Table 1S.

Tissue Fractionations and Western Blot Analysis.

Western blots of liver homogenate lysates were performed as described5 using mouse-specific primary antibodies to GRP78 (1:1,000) (Assay Designs), GADD153 (2 μg/mL) (Abcam), caspase-12 (2 μg/mL) (Sigma), ATF6 (2 μg/mL), ATF4 (1 μg/mL) (Imgenex), nuclear SREBP-1c (1:1,000) (Santa Cruz Biotechnology), and β-actin (1:10,000) (Sigma). Horseradish peroxidase–conjugated anti-rabbit immunoglobulin G (IgG) (Pierce, Rockford, IL) was used as the secondary antibody. Blots were developed with the HRP SuperSignal chemiluminescent detection system (Pierce). Band intensities were quantified using ImageQuant software (Molecular Dynamics) and standardized against β-actin.

Global DNA Methylation.

DNA was isolated from each liver sample (Qiagen, Valencia, CA) for assay of global DNA methylation by liquid chromatography tandem mass spectrometry,24 which measures the percentage of methylated dCyt in the DNA sample.

Chromatin Immunoprecipitation Assay.

Chromatin immunoprecipitation (ChIP) assays were performed following a tissue protocol.25 Briefly, 50 mg of liver tissues were cut in small pieces with a razor blade, cross-linked in 1.5% formaldehyde for 15 minutes, processed in a Medimachine (BD Biosciences) using a 50-μm medicon to produce a liver cell suspension. Nuclear extracts were prepared and sonicated using a Bioruptor Sonicator (Diagenode) and precleared using blocked Staphylococcus A cells. Ten percent of original precleared chromatin was removed for use as a control for total input DNA. In ChIP analyses, the antibody to the methylated histone immunoprecipitates and isolates the DNA/histone complex. Using selective and region-specific primers, subsequent PCR determines the extent of trimethylated histone binding to the promoter region of each relevant gene. Each ChIP assay was performed using 500 ng of chromatin and 2 μL of antibody. The primary antibody was rabbit polyclonal 3meH3K9 IgGs (Abcam, catalog # ab8898). Secondary rabbit anti-mouse IgG was purchased from MP Biomedicals (catalog # 55436). Nonspecific rabbit IgG was used as a negative control for the ChIP assays (Alpha Diagnostics, catalog # 20009-5). For PCR analysis of the ChIP samples, purified immunoprecipitates (QIAquick PCR purification kit, Qiagen) were dissolved in 20 μL of water. ChIP-enriched samples and inputs were analyzed in triplicate by way of PCR using primer sequences of promoter regions of GRP78, GADD153, SREBP-1c, and glyceraldehyde 3-phosphate dehydrogenase, as shown in Supporting Table 2S. PCR products were separated by electrophoresis through 1.5% agarose gels, visualized using ethidium bromide, and quantitated with ImageQuant Software (Molecular Dynamics). Data were normalized with input control.

Statistical Analyses.

Significant differences between groups were determined by two-way analysis of variance. Statistical significance was assessed at P < 0.05 to determine the effects of ethanol feeding and genotype. Relationships among variables were determined by linear regression analyses of individual values using SPSS data editor 14.0 for Windows (SPSS, Inc., Chicago, IL).

Results

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Effects of Ethanol Feeding on Liver/Body Weight, Plasma Ethanol, and ALT Levels.

Four weeks of intragastric ethanol feeding increased liver/body weight ratios in both ethanol-fed groups with an interaction of ethanol and genotype in the heterozygous (Het-E) group (Table 1). Terminal plasma ethanol levels were elevated more than 40-fold, and ALT levels were elevated more than 10-fold in both ethanol-fed groups, consistent with previous studies.6, 26

Table 1. Effects of Intragastric Feeding of Control and Ethanol Diets on CβS Wild-Type and Heterozygous Mice
 Wild-Type ControlHeterozygous ControlWild-Type Ethanol-FedHeterozygous Ethanol-Fed
  • Values are expressed as the mean ± standard deviation.

  • *

    Ethanol effect (P < 0.001, P < 0.001, P < 0.001, P < 0.001, P < 0.005, P < 0.05, P < 0.003, P < 0.004, P < 0.001, and P < 0.001.)

  • Interaction of ethanol and genotype effects (P < 0.05, P < 0.05, and P < 0.01).

  • Genotype effect (P < 0.03, P < 0.04, P < 0.001, and P < 0.001).

Liver/body weight0.05 ± 0.0030.04 ± 0.0030.09 ± 0.002*0.101 ± 0.019*
Plasma ethanol (mg/mL)8.40 ± 5.535.60 ± 4.67352 ± 64.3*259 ± 79.2*
Plasma ALT (U/L)25.20 ± 7.35620.5 ± 7.33247 ± 63.1*214 ± 77.3*
Histology total score1.40 ± 1.140.5 ± 0.584.6 ± 1.52*7.0 ± 2.45*
Fat score0.60 ± 0.500.0 ± 0.003.0 ± 0.30*3.6 ± 0.20*
TUNEL score3.40 ± 4.009.0 ± 4.008.0 ± 2.00*17.0 ± 4.00*
GSH (nmol/mg)60.9 ± 3.4152.2 ± 3.4749.3 ± 4.64*47.9 ± 6.79*
Homocysteine (nmol/mg)0.33 ± 0.020.31 ± 0.020.30 ± 0.030.30 ± 0.09
SAM (nmol/mg)0.22 ± 0.050.19 ± 0.040.15 ± 0.04*0.14 ± 0.04*
SAH (nmol/mg)0.08 ± 0.020.17 ± 0.080.14 ± 0.02*0.20 ± 0.04*
SAM/SAH2.90 ± 0.801.40 ± 0.681.05 ± 0.33*0.75 ± 0.21*

Effects of Ethanol Feeding and Genotype on Liver Histopathology and Apoptosis.

The total liver histopathology score increased in both ethanol groups with an interactive effect of ethanol and genotype in the Het-E group (Fig. 1, Table 1). Intracellular macroscopic lipid according to fat score increased in both ethanol-fed groups (Table 1). There were nonsignificant increases in inflammatory cells and necrosis in the heterozygote ethanol-fed group and no fibrosis in any mice. TUNEL assay revealed increased hepatocellular apoptosis in both genotype and ethanol feeding, with additive effects in the Het-E group (Table 1).

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Figure 1. Effects of diets and CβS genotype on liver histopathology. Representative histology is shown for liver specimens from (A) wild-type control, (B) heterozygous control, (C) wild-type ethanol-fed, and (D) Het-E mice. Livers from both control-fed mice are normal. The liver from the wild-type ethanol-fed control mouse shows macrovesicular fat formation and an apoptotic cell formation; the liver from the Het-E mouse shows a necrotic focus and macrovesicular fat formation (magnification ×160).

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Effects of Ethanol and Genotype on Liver GSH, Homocysteine, SAM, and SAH.

Liver GSH, a measure of antioxidant defense capacity, was reduced in the heterozygous control and in both ethanol-fed groups, with additive effects of ethanol feeding and genotype in the Het-E group (Table 1). There were no differences among the groups in liver homocysteine levels. Liver SAM was reduced and SAH was elevated in both ethanol-fed groups, with an additive effect of genotype on SAH in the Het-E group. The SAM/SAH ratio of methylation capacity decreased in both ethanol fed groups, with interactive effects of genotype and ethanol in the Het-E group. The SAM/SAH ratio correlated negatively with the total pathology score (r = −0.57, P < 0.006) and TUNEL score (r = −0.52, P < 0.01). Scatter plots of these and subsequent regression analyses are shown in Supporting Figs. 1–5.

Effects of Ethanol and Genotype on ER Stress Pathways.

ER chaperone GRP78 messenger RNA (mRNA) (Table 2) and its protein levels (Fig. 2A) increased in both ethanol-fed groups, with an interaction of genotype and ethanol on protein levels in the Het-E group. Protein levels of the ER stress transducer ATF4 increased in both ethanol groups with greatest and interactive effects in the Het-E group (Fig. 2B). Activated ER stress transducer ATF6 increased by genotype and maximally in the Het-E group (Fig. 3C). Liver transcript levels of the pro-apoptotic gene GADD153 increased in both ethanol-fed groups (Table 2), while protein expression rose with both genotype and ethanol feeding, with interactive effects in the Het-E group (Fig. 2D). Cleaved caspase 12, a protease that plays a central role in initiating ER stress-induced apoptosis, increased in both groups of ethanol-fed mice (Fig. 2E). Transcript and protein levels of SREBP-1c increased in both ethanol-fed groups, with additive and interactive effects of both treatments on mRNA expression in the Het-E group (Table 2, Fig. 2F). Ethanol feeding increased SREBP-1c targeted transcripts of acetyl-coenzyme A carboxylase, with interactive effects in the Het-E group, while fatty acid synthase expression rose by genotype only (Table 2). The SAM/SAH ratio of methylation capacity correlated negatively with protein levels of GRP78 (r = −0.43, P < 0.04), GADD153 (r = −0.62, P < 0.002), and cleaved caspase-12 (r = −0.73, P < 0.002).

Table 2. Effects of Ethanol and Genotype on Liver Transcripts of ER Stress Genes
 Wild-Type ControlHeterozygous ControlWild-Type Ethanol-FedHeterozygous Ethanol-Fed
  • All values are normalized to β-actin transcripts and are expressed as the mean ± standard error.

  • *

    Effects of ethanol (P < 0.05, P < 0.05, P < 0.03, and P < 0.05).

  • Effects of genotype (P < 0.02 and P < 0.05).

  • Interaction of ethanol and genotype (P < 0.05 and P < 0.05).

GRP784.2 ± 0.85.5 ± 1.87.1 ± 0.6*8.6 ± 1.0*
GADD1532.0 ± 0.32.3 ± 0.52.7 ± 0.1*3.5 ± 0.2*
SREBP-1c0.6 ± 0.11.2 ± 0.21.8 ± 0.5*4.0 ± 0.9*
Acetyl-coenzyme A carboxylase1.2 ± 0.11.1 ± 0.52.9 ± 0.5*7.3 ± 2.0*
Fatty acid synthase1.7 ± 0.22.3 ± 0.11.6 ± 0.33.0 ± 0.6
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Figure 2. Effect of diets and genotype on hepatic protein expression of ER stress genes. Representative western blots are shown above each bar graph of image intensities in each group, expressed as ratios with actin ± standard error. (A) GRP78. Ethanol effect (P < 0.001). Interaction of ethanol and genotype (P < 0.03). (B) Activated ATF4. Ethanol effect (P < 0.05). Interaction of ethanol and genotype (P < 0.05). (C) Activated ATF6. *Genotype effect (P < 0.05). (D) GADD153. *Genotype effect (P < 0.04). Ethanol effect (P < 0.001). Interaction of ethanol and genotype (P < 0.01). (E) Cleaved caspase-12. Ethanol effect (P < 0.002). (F) nSREBP-1c. Ethanol effect (P < 0.004).

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Figure 3. Immunohistochemistry. Representative mouse liver sections were stained with antibody to H3K9me3 and a secondary antibody labeled with FITC, then viewed with fluorescent microscopy. The liver sections from (A) wild-type control and (B) heterozygote control mice show uniform bright staining of hepatocyte nuclei of both central and peripheral hepatocytes. In contrast, the centrilobular hepatocytic nuclei of the (C) wild-type ethanol-fed mice and (D) Het-E mice show reduced brightness of the centrilobular hepatocytic nuclei compared with those in peripheral regions and with findings in both control groups. C, centrilobular region; P, peripheral region (magnification ×160).

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Global DNA Methylation.

The percentages of methylated cytosine were similar among all groups: 4.01% ± 0.03 in wild-type controls, 4.0% ± 0.1 in heterozygous controls, 3.8% ± 0.01 in wild-type ethanol-fed, and 3.9% ± 0.2 in Het-E mice.

Immunofluorescent Analysis of Nuclear Chromatin Histone.

Immunofluorescent staining with antibody to gene suppressor 3meH3K9 showed greatest intensity in the centrilobular regions of the liver of a control wild-type mouse (Fig. 3A) and least intensity in the centrilobular and peripheral regions of the liver from an ethanol-fed heterozygote mouse (Fig. 3D), with intermediate and predominately centrilobular staining in a heterozygote control (Fig. 3B) and wild-type ethanol fed mouse (Fig. 3C). Quantitative values for each group showed significant ethanol effect on the centrilobular distributions of fluorescent hepatocyte nuclei (P < 0.02), without differences in peripheral distributions. Antibodies to 3meH3K4 showed no differences among the groups (data not shown).

ChIP Analysis of Repressive 3meH3K9 in Promoter Regions of GRP78, GADD153, and SREBP-1c.

We examined quantitative binding of the repressive epigenetic marker 3meHeK9 to selective gene promoters using the ChIP assay and semiquantitative PCR analyses. We preferentially selected three liver specimens from each group according to highest histopathology score and lowest SAM/SAH ratios. Each sample was measured three times, using mean values for subsequent statistics. As shown in Fig. 4 and Table 3, 3meH3K9 binding to the promoter regions of GRP78, GADD153, and SREBP-1c decreased in response to ethanol feeding, with an interaction of ethanol and genotype for GRP78 binding in Het-C mice. Binding of 3meH3K9 to promoters of GRP78 and GADD153 correlated positively with the liver SAM/SAH ratio (r = 0.61, P < 0.03; r = 0.69, P < 0.01) and negatively with liver SAH levels (r = −0.52, P < 0.05; r = −0.62, P < 0.05).

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Figure 4. Chromatin immunoprecipitation. Representative ChIP results from each group, using antibodies to 3meH3K9 and promoter primers from selected ER stress genes known to be activated by ethanol feeding with interaction with genotype (GRP78 and GADD153) or alone (SREBP-1c). 3MeH3K9 ChIP DNA samples were prepared from selected mouse livers followed by PCR analysis to amplify the promoter regions of candidate genes. Quantification of PCR products in each group normalized to input control is shown in Table 3.

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Table 3. Effects of Ethanol and Genotypes on Binding of Trimethylated H3K9 to Selected ER Stress Gene Promoters According to ChIP Analysis
 Wild-Type ControlHeterozygous ControlWild-Type Ethanol-FedHeterozygous Ethanol-Fed
  • PCR products of trimethylated histone H3K9 and input DNA (Fig. 3) were fractionated on agarose gels, photographed, and quantified with ImageQuant software. All values are normalized to input control and are expressed as the mean ± standard error.

  • *

    Interaction of ethanol and genotype (P< 0.02).

  • Effects of ethanol (P < 0.05, P < 0.05, and P < 0.03).

GRP781.05 ± 0.080.8 ± 0.020.6 ± 0.3*0.06 ± 0.01*
GADD1533.9 ± 0.22.5 ± 0.51.3 ± 0.7*1.3 ± 0.3*
SREBP-1c3.8 ± 0.24.0 ± 0.22.3 ± 0.8*1.3 ± 0.7*

Effects of Diets on mRNA Expression of Histone H3K9 Methyltransferases.

The liver transcripts of EHMT2 (G9a), which dimethylates H3K9, were down-regulated in heterozygote control mice and in ethanol-fed mice of each genotype, while expressions of other methyltransferases were similar among the groups (Table 4). However, the expressions of EHMT2 (G9a) and Setdb1 correlated positively with liver SAM/SAH ratio (r = 0.66, P < 0.006; r = 0.64, P < 0.01) and negatively with liver SAH levels (r = −0.58, P < 0.01; r = −0.48, P < 0.05), consistent with a regulatory role of methylation in expression of these enzymes.

Table 4. Effects of Ethanol and Genotypes on Liver K9 Histone Methyltransferase Transcripts
 Wild-Type ControlHeterozygous ControlWild-Type Ethanol-FedHeterozygous Ethanol-Fed
  • All values are normalized to β-actin transcripts and are expressed as the mean ± standard error.

  • *

    Effects of genotype (P < 0.01).

  • Effect of ethanol (P < 0.02).

G9a16.0 ± 2.011.0 ± 0.3*11.0 ± 0.26.0 ± 2.2*
SUV39h120.0 ± 1.514.0 ± 0.425.0 ± 3.019.0 ± 4.0
SUV39h252.0 ± 10.040.0 ± 2.441.0 ± 3.533.0 ± 9.6
Setdb142.0 ± 11.023.0 ± 9.023.5 ± 4.026.9 ± 6.0

Discussion

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

While others studied hepatic ER stress in CβS-deficient26 and in intragastric ethanol-fed mice,6, 27 ours is the first study to combine CβS deficiency with high ethanol exposure through intragastric feeding in order to test the hypothesis that ethanol-induced aberrant methionine metabolism regulates the pathogenesis of ASH. The model showed that altered methylation, as evidenced by changes in the liver SAM/SAH ratio by interactions of genotype and ethanol feeding, affected epigenetic regulation of genes involved in the ER stress pathways of lipogenesis and apoptosis. Histological evidence of advanced liver injury and apoptosis resulting from the interaction of the two treatments (Table 1, Fig. 1) was paralleled by additive or interactive effects of these treatments on liver SAH and the SAM/SAH ratio, as well as by decreases in the transsulfuration product and principal antioxidant GSH (Table 1). These findings are consistent with our prior observations of worsening liver injury in relation to abnormal SAM/SAH ratio in micropigs fed ethanol with folate-deficient diets.1, 5 The finding of unchanged hepatic homocysteine concentrations among groups is most likely due to its conversion to SAH through the reverse SAH hydrolase reaction. Others who used the same wild-type C56Bl6J mouse showed marked elevation of plasma homocysteine after intragastric ethanol feeding but did not measure liver levels,6, 27 whereas we previously found four-fold elevation of plasma homocysteine but only modest increase in liver levels in chronic ethanol fed micropigs.1 The concentration disparity is likely due to the fact that homocysteine undergoes continuous rapid metabolism in the liver, whereas plasma homocysteine is not metabolized and represents the cumulative export of homocysteine from liver and other tissues.28 The metabolic regulation of homocysteine in the liver would predictably cause elevated liver SAH in the Het-E group as a result of the dual inhibitory effects of ethanol on transmethylation of homocysteine to methionine and of CβS deficiency on reducing homocysteine excretion through the transsulfuration pathway.4 The correlation between the decreased SAM/SAH ratio of methylation capacity and the worsening histopathology and apoptosis in the present model strengthens evidence that aberrant methionine metabolism contributes to the pathogenesis of ASH.

In evaluating mechanisms for development of ASH through altered methionine metabolism in our model, we found that ethanol, genotype, and their interaction increased the induction of ER stress pathways of lipogenesis and apoptosis. These pathways included enhanced expression of ER chaperone GRP78 and lipogenic transcription factor SREBP 1-c, as well as apoptosis mediators ATF4, ATF6, GADD153, and caspase-12 (Table 2, Fig. 2). These findings extend other observations on ER stress from the intragastric ethanol-fed mouse.6, 27 Furthermore, the findings on the relationships of altered SAM/SAH ratio and ER stress-induced lipogenesis and apoptosis can explain the effects of the different diets on the histopathology and TUNEL scores shown in Table 1 and Fig. 1.

In addition to ER stress, the increased response of SREBP-1c mRNA expression to ethanol feeding (Table 2) may also reflect the additional contribution of the adiponectin signaling pathway of lipogenesis, as described in ethanol-fed micropigs7 and in C57BL6 mice fed oral ethanol diets.29 However, the effect of intragastric infusion of a high ethanol diet on the adiponectin signaling pathway of steatosis is not known. The enhanced SREBP-1c expression in the Het-E group (Table 2) is consistent with our prior finding of its correlation with elevated SAH levels in the ethanol-fed micropig.5 The observed discordance of mRNA and protein levels of SREBP-1c in the Het-E group (Table 2, Fig. 2F) may reflect instability and enhanced protein degradation of SREBP-1c. It is unlikely that this observation reflects a defect of in vivo translation and nuclear processing of SREBP-1c, because we observed a similar approximately two-fold increase in the transcript levels of its targeted lipogenesis genes acetyl-coenzyme A carboxylase and fatty acid synthase in the same Het-E group (Table 2). The interactive effect of CβS heterozygosity and ethanol feeding on ATF4 expression (Fig 2B) is a novel finding with no obvious mechanism. It is known that ER stress induces phosphorylation of eukaryotic initiation factor 2 concomitant with increased production of ATF4. The potential effects of altered methionine metabolism through CβS deficiency and its interaction with ethanol on eukaryotic initiation factor 2 phosphorylation and hence ATF4 are not known.

Increasing evidence suggests that ethanol-induced epigenetic changes contribute to the development of ASH.30 Studies in primary hepatocytes from ethanol-treated rats found associations of dimethylated or trimethylated H3K4 with promoter regions of up-regulated genes, including alcohol dehydrogenase and glutathione S-transferase, whereas dimethylated or trimethylated H3K9 was associated with genes down-regulated by ethanol, including L-serine dehydrase and CYP450 2c11.31 SAM treatment blocked LPS-induced tumor necrosis factor expression in a murine macrophage cell line by inhibiting trimethylated H3K4 binding to its promoter region.32 In an intragastric ethanol-fed rat model, ethanol-induced proteosome inhibition was associated with reduced levels of H3K9 dimethylation,33 whereas increased hepatic levels of dimethylated H3K4 indicated increased gene activation in chronic ethanol-fed rats.34 Therefore, methylation at different lysine residues of histone H3 have opposite effects on gene expression. Another study showed that dietary methyl deficiency in rats and mice leads to changes in methyltransferase expression and levels of methylated histones.35 The fact that we found no changes in global DNA methylation among the groups underscores the importance of evaluating effects of diet and genotype on specific methylated histone-regulated genes in our study.

Immunohistochemical analysis of the mouse livers showed antibody binding sites for 3meH3K9 but not for 3meH3K4, and diminished binding to gene suppressor sites for 3meH3K9 in centrilobular but not peripheral regions of lobules of ethanol-fed mice (Fig. 3). 3meH3K9 covers broad regions of the genome as a chromatin-repressive marker that binds to gene promoters, followed by assembly of repressive complexes and transcriptional gene silencing.36 Therefore, reduced 3meH3K9 binding predicts gene activations consistent with enhanced mechanisms for centrilobular apoptosis and steatosis, which explains the present observations (Fig. 3) and is consistent with our prior findings of early centrilobular steatosis and hepatocellular apoptosis in the ethanol-fed micropigs.37 Based on these findings, we used an antibody to 3meH3K9 in the ChIP assay to study the effects of histone H3 lysine methylation on relevant ER stress genes. The selection of liver samples for ChIP assays was based on the demonstrated changes induced by diet and genotype on liver methionine metabolites and histopathology, in accordance with our hypothesis that epigenetic effects would be determined by these changes. Results from the preselected liver samples were not likely to have been artifactual, because they were repeated in triplicate with excellent fidelity. However, since the experiments used preselected specimens, the data cannot be generalized to the entire population of mice in each group. Our findings of reduced 3meH3K9 binding to promoters of GRP78, GADD153, and SREBP-1c (Fig. 4, Table 3) support the hypothesis that increased gene activation through altered histone modifications may be a key mechanism underlying ethanol-induced gene activation in ASH, in particular those involved in pathways of ER stress-related apoptosis and lipogenesis. Additional studies are required to determine the effects of ethanol and CβS genotype on the levels of transcription factors such as C/EBPβ, XBP-1 and YY1 to the promoter of GRP78, on expression of other genes relevant to alcoholic liver injury, and on the binding of other epigenetic markers such as methylated histone H3K4, H3K27, and acetylated histone H4.

To study additional mechanisms for the observed findings from ChIP with 3meH3K9, we then measured the expression of four recognized H3K9 methyltransferases. The reduced expression of EHMT2 (G9a) (Table 4) suggests that ethanol-induced changes in expression of this enzyme play a role in differential effects on H3K9 methylation among the groups. Further, the correlations among SAM/SAH methylation ratio and SAH levels with the expression of both EHMT2 and Setdb1suggest that altered methylation capacity plays a role in the expression of these methyltransferases. Additional studies are required to determine the full extent of the effects of other histone methylation modifications and their mediation by their histone methyltransferases on the expression of ER stress and other genes relevant to alcoholic liver injury.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

We acknowledge the support of S. Jill James and Stephan Melnyk at the Arkansas Children's Hospital Research Institute (Little Rock, AR) for assays of liver levels of GSH, homocysteine, SAM, and SAH. We are also thankful to members of Peggy Farnham's laboratory at the University of California, Davis, for assistance in developing the ChIP assay.

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  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and Methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  8. Supporting Information

Additional Supporting Information may be found in the online version of this article.

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HEP_23382_sm_suppfig1.tif692KSupplemental Figure 1
HEP_23382_sm_suppfig2.tif1721KSupplemental Figure 2
HEP_23382_sm_suppfig3.tif694KSupplemental Figure 3
HEP_23382_sm_suppfig4.tif687KSupplemental Figure 4
HEP_23382_sm_suppfig5.tif695KSupplemental Figure 5

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