The Journal of Nutrition Vol. 128 No. 4 April 1998, pp. 758-763

Chronic Ethanol Consumption Affects Glutathione Status in Rat Liver^1,2,3

Se In Oh, Cho-Il Kim^*, Hui Jung Chun, and and Sang Chul Park⁴

Department of Biochemistry, College of Medicine, Seoul National University, Seoul 110-799, Korea; ^* Nutrition Research Department, Korea Institute of Food Hygiene, Seoul 156-050, Korea; and  Department of Food and Nutrition, Sookmyung Women's University, Seoul 140-742, Korea

	ABSTRACT

Abstract Introduction Methods Results Discussion References

There is no consensus yet on the role of oxidative stress in the nutritional outcome of chronic ethanol feeding and the status of cellular antioxidative defense systems against ethanol toxicity. In this study, chronic alcohol consumption in humans was reproduced in Sprague-Dawley rats to investigate the effect of ethanol ingestion on the regulation of oxidative stress in liver with a special focus on glutathione. Adult male rats were given 36% of total energy as alcohol in the Lieber-DeCarli liquid diet for 6 wk. The control group was pair-fed the diet containing isocaloric dextrin-maltose instead of ethanol. Chronic ethanol ingestion enhanced expression of cytochrome P450 II E1 in the liver, but did not significantly alter either the level of hepatic thiobarbituric acid reactive substances or the carbonyl group content of proteins. The hepatic concentrations of total and reduced glutathione and the activities of catalase, glutathione reductase and glutathione S-transferase were significantly higher in the ethanol group than in the control group. The activities of glutathione peroxidase and glucose-6-phosphate dehydrogenase were significantly lower in the ethanol group than in controls. Chronic ethanol consumption by well-nourished rats for 6 wk increased enzyme activities related to the recycling and utilization of glutathione in the liver. Such an enhancement in the activities of the hepatic antioxidative defense system may be one of the protective mechanisms of the body against oxidative tissue damage caused by ethanol-induced free radicals.

	INTRODUCTION

Abstract Introduction Methods Results Discussion References

Ethanol toxicity, including a decline in nutritional status, is directly due to ethanol per se and its metabolite, acetaldehyde, or is indirectly due to the metabolic sequelae of ethanol oxidation such as the decreased ratio of cytoplasmic NAD⁺/NADH and the involvement of reactive oxygen species (Lieber 1991). The microsomal cytochrome P450 system and xanthine oxidase pathway may be responsible for the generation of ethanol-induced oxygen radicals (Bondy and Naderi 1994, Hodgson and Levi 1994, Lieber and DeCarli 1970). The consequences of radical formation include the peroxidation of lipids, carbonylation of proteins and hydroxylation of nucleotides (Comporti 1993, Meneghini et al. 1993). However, the oxidative stress and resulting tissue pathology in the liver after chronic ethanol feeding have been disputed (Coudray 1993, Ekström and Ingelman-Sundberg 1989, Kawase et al. 1989, Shaw et al. 1983). The discrepancies among the reports seem to originate from attempts to explain the complex reactive phenomena in the body, especially the changes in the radical generation and radical scavenging systems, in response to chronic ethanol feeding.

On the basis of the assumption that the alcohol-associated pathologic changes resulted from the interaction between the alcohol-induced radical generation and the reactive radical scavenging capacity of the tissues, the cellular defense system against ethanol-induced oxidative stress has been studied intensively, but with discordant results (Fernández-Checa et al. 1987 and 1991, Garcia-Ruiz et al. 1994, Hassing et al. 1979, Hetu et al. 1982, Hirano et al. 1992, Iizuka et al. 1991, Martensson et al. 1990, Morton and Mitchell 1985, Shaw et al. 1983, Teare et al. 1994). Although the role of glutathione in the detoxification processes of ethanol-induced radicals has been illustrated through alterations in glutathione content and glutathione-utilizing enzyme activities (Hirano et al. 1992, Shaw et al. 1983), there are few studies of the effect of ethanol on the regulation of glutathione redox status. Because glutathione must be in reduced form to be utilized, it is necessary to maintain or enhance the level of glutathione reductase and its coenzyme, NADPH. Therefore it is important to study the effect of chronic ethanol feeding on the oxidative liver damage by determining the activities of glutathione reductase, the major enzyme for glutathione reduction, and glucose-6-phosphate dehydrogenase, the enzyme responsible for NADPH supply simultaneously with monitoring the activities of glutathione-utilizing enzymes and the amount of total and reduced glutathione.

	MATERIALS AND METHODS

Abstract Introduction Methods Results Discussion References

Animals.  Weanling male Sprague-Dawley rats consumed a nonpurified laboratory rat diet (Samyang, Seoul, Korea) and water ad libitum until they weighed 120-180 g. Then the rats were divided into weight-matched pairs and allocated to the two experimental groups. They were housed in individual stainless steel wire-bottomed cages in a room kept at 22-25°C with a 12-h light:dark cycle. All animal procedures and handling were conducted in compliance with the Seoul National University guidelines.

Experimental design.  Rats in the ethanol group consumed a liquid diet containing 36% of energy as ethanol ad libitum, and the pair-fed daily control rats received an isocaloric amount of similar diet containing dextrin-maltose instead of ethanol on the following day. Ethanol was introduced into the diet gradually starting from 0% (w/v) and increasing to a final 5% (w/v) over a 1-wk period. Rats continued that feeding regimen for five more weeks. On d 41, food was given in two half-portions, one in the morning and the other half in the evening. The rats were killed the next morning by decapitation. Diet was supplied by Dyets (Bethlehem, PA) and compositions were as reported by Kim et al. (1988).

Sample preparation.  Blood was taken from the neck vessels and let stand for 30 min at room temperature. Serum was obtained by centrifugation at 1000 × g for 20 min and stored at 70°C in aliquots until the analyses. Livers were homogenized in 9 volumes of ice-cold 11.5 g/L KCl with 0.2 mmol/L phenylmethylsulfonylfluoride and 1 mmol/L dithiothreitol. Homogenates were centrifuged at 800 × g for 10 min to remove cell debris and nuclei; the supernatant was centrifuged at 13,200 × g for 10 min. The postmitochondrial supernatant (13,200 × g) was centrifuged again at 105,000 × g for 60 min to prepare microsomes and cytosol fractions. The pellet (microsomes), resuspended in the same buffer, and the supernatant fraction (cytosol) were centrifuged again at 105,000 × g for 30 min to reduce cross-contamination. Washed microsomes resuspended in the same buffer and the cleaner supernatant taken as the cytosol fraction were stored at 70°C in aliquots. The whole procedure was conducted at 0-4°C.

Liver histology.  A piece of liver tissue was fixed in formalin and embedded in paraffin; sections (6-µm thick) were cut, and each section was stained with hematoxylin and eosin.

Biochemical analysis.  Serum alkaline phosphatase (EC 3.1.3.1), lactate dehydrogenase (EC 1.1.1.27), uric acid, cholesterol, total protein, albumin, total bilirubin and direct bilirubin were measured on a Technicon autoanalyzer (Technicon, Tarrytown, NY). Activities of aspartate aminotransferase (EC 2.6.1.1) and alanine aminotransferase (EC 2.6.1.2) were determined using commercially available diagnostic kits (#58-20 and #59-20; Sigma Chemical, St. Louis, MO).

Xanthine oxidase (EC 1. 1. 3. 22) and xanthine dehydrogenase activities were measured by using a slight modification of the methods descibed previously (Waud and Rajagopalan 1976). The enzyme reaction mixture contained 0.2 mmol/L xanthine, 0.1 mol/L Tris-HCl buffer (pH 8.1), 0.1 mmol/L EDTA and 150 µL of cytosol fraction in a final volume of 1 mL. Activity was monitored by reading optical densities at 300 and 340 nm, in the absence or presence of 0.5 mmol/L NAD⁺ at 30°C. Specific activity was calculated as nanomoles per milligram protein.

The content of 2-thiobarbituric acid reactive substances (TBARS)⁵ in the liver homogenate was measured as a marker of lipid peroxidation by the method of Uchiyama and Mihara (1978) with the use of 1,1,3,3-tetraethoxypropane as a standard.

The extent of protein carbonylation was also determined in liver homogenates by using 2,4-dinitropheylhydrazine reaction according to the method of Levine et al. (1990). Insoluble materials were removed by centrifugation in the Eppendorf tubes and the absorbance of the supernatant was read. A molar absorption coefficient of 22 ×10⁶ cm² at 370 nm was used to calculate the carbonyl group content.

Two small pieces of liver were cut, rinsed in homogenate buffer and dried by blotting. One piece was immediately placed into liquid N₂, stored at 70°C and used within the next 24 h to measure oxidized glutathione (GSSG). The other piece was used while fresh to measure total glutathione (GSH+GSSG) with the method detailed by Sies and Akerboom (1984). For either GSSG or GSH+GSSG, calculation was based on the standard curves prepared with GSSG for each experiment. The content of reduced glutathione (GSH) was obtained from the difference between two values.

Glutathione peroxidase (EC 1.11.1.9) was assayed in 50 mmol/L Tris-HCl , 0.1 mmol/L EDTA buffer (pH 7.6) at 37°C by measuring the oxidation of NADPH at 340 nm with cumene hydroperoxide as a substrate (Tappel 1978). Glutathione reductase (EC 1.6.4.2) was assayed in 0.2 mol/L potassium phosphate buffer, pH 7.0, by monitoring the oxidation of NADPH at 340 nm with GSSG as a substrate (Carberg and Mannervick 1985). Assay of glutathione S-transferase (EC 2.5.1.18) activity was performed in phosphate buffer, pH 6.5, by using 1-chloro-2,4-dinitrobenzene as a substrate; the absorbance change was recorded at 340 nm (Harbig et al. 1974). The activity was calculated by using a molar extinction coefficient of 9.6 × 10⁶ cm². Liver catalase (EC 1.11.1.6) activity was assayed in the homogenate at 20°C according to the method of Aebi (1984). H₂O₂ disappearance was monitored kinetically at 240 nm and the activity was expressed as the rate constant (k/mg protein) as recommended by Aebi (1984). Glucose-6-phosphate dehydrogenase (EC 1.1.1.49) activity was measured at 25°C according to Löhr and Waller (1974).

Proteins.  Proteins in liver samples were assayed according to the Bradford method (1970) with bovine serum albumin as a standard.

Western blotting.  SDS polyacrylamide gel electrophoresis was conducted by using a 10% acrylamide gel with 30 µg of microsomal protein in each well. Gels were transferred to nitrocellulose paper (Laemmli 1970). The enhanced chemiluminescence method was adopted to monitor the intensity of P450 II E1 bands with mouse monoclonal antibody as the primary antibody against cytochrome P450 II E1 (kindly donated by Byung June Song from NIAAA) and peroxidase-conjugated antimouse immunoglobulin (Vector Laboratories, Burlingama, CA) as the secondary antibody.

Statistical analysis.  Data were expressed as means ± SEM. Statistical significance of the difference between group means was determined by Student's t test. Differences with P < 0.05 were considered to be significant.

	RESULTS

Abstract Introduction Methods Results Discussion References

Although the total food intake was similar between the two groups due to pair-feeding, ethanol-fed rats gained less weight than pair-fed controls during the study period (2.39 ± 0.19 vs. 2.93 ± 0.08 g/d, P < 0.05). As a result, there was a significantly lower energy efficiency ratio (P < 0.01) in the chronic ethanol group compared with that of the control group (11.69 ± 0.74 vs. 14.24 ± 0.33 g gain/kJ).

Liver weight per 100 g of body weight was significantly different (P < 0.001) between ethanol and pair-fed control groups (3.46 ± 0.12 vs. 2.79 ± 0.08 g liver/100 g body wt).

On hematoxylin and eosin stained sections, parenchymal accumulation of lipid was seen as clear intracytoplasmic vacuoles in the livers of the chronic ethanol group. The hepatocytes were uniformly filled with large and small fat droplets with nuclei eccentrically placed (Fig. 1).

View larger version (147K): [in this window] [in a new window]   View larger version (170K): [in this window] [in a new window]   Fig 1. Representative micrographs of livers of rats that consumed ethanol-containing or control liquid diet for 6 wk. Liver histology of the pair-fed control group (A) is normal and that of the chronic ethanol group (B) shows alcoholic fatty liver. The hepatocytes (B) are uniformly filled with large fat droplets and nuclei are eccentrically placed. (H & E staining, ×200).

After ethanol had been administered for 6 wk, activities of serum alkaline phosphatase and alanine aminotransferase, and total bilirubin level in serum were significantly greater (P < 0.05) in the ethanol group than in the control group (7.6 ± 0.6 vs. 5.0 ± 1.0 µkat/L, 1.46 ± 0.13 vs. 0.93 ± 0.09 µkat/L and 9 ± 2 vs. 7 ± 0 µmol/L, respectively). However, chronic ethanol treatment had no effect on the other serum metabolites measured.

Chronic ethanol consumption induced cytochrome P450 II E1 (Fig. 2), suggesting a possible involvement of the activated oxygen radicals formed during ethanol metabolism in ethanol-induced liver pathology. Although the activity of xanthine oxidase in the liver was significantly lower after chronic ethanol treatment, that of xanthine dehydrogenase was not different between ethanol-fed and control rats (Table 1).

View larger version (47K): [in this window] [in a new window]   Fig 2. Western blot of liver microsomes of rats that consumed ethanol-containing or control liquid diet for 6 wk. Mouse monoclonal antibody reactive cytochrome P450 II E1 was detected by enhanced chemiluminescence; the band is visible only in the chronic ethanol group. (Lane 1, pair-fed control group; lane 2, chronic ethanol group).

The TBARS and carbonyl group levels in the liver homogenates did not differ between pair-fed control and ethanol-fed rats (0.442 ± 0.024 vs. 0.445 ± 0.057 nmol/mg protein and 2.65 ± 0.18 vs. 2.41 ± 0.22 nmol/mg protein, respectively).

The hepatic levels of total and reduced glutathione in ethanol-treated rats were significantly greater than in control rats (P < 0.0001), whereas the oxidized form of glutathione in liver was lower in the ethanol group (P < 0.05, Table 2). The activities of hepatic glutathione peroxidase and glucose 6-phosphate dehydrogenase in the ethanol group were significantly lower than those of controls (P < 0.05, Table 3). On the other hand, 6 wk of ethanol administration resulted in significantly higher hepatic activities of catalase, glutathione reductase and glutathione S-transferase (P < 0.01).

	DISCUSSION

Abstract Introduction Methods Results Discussion References

Isocaloric substitution of carbohydrates by ethanol resulted in lower weight gain in spite of similar energy intake, as reported previously (Kawase et al. 1989, Pirola and Lieber 1975). This lower body weight gain has been attributed to induction of the microsomal ethanol oxidizing system (a metabolic pathway that oxidizes ethanol without associated chemical energy production), increased sympathetic tone and associated thermogenesis and/or enhanced ATP breakdown (with increased purine catabolism) secondary to the acetate production from ethanol (Cunningham and Spach 1987, Lieber 1994).

The 6 wk of ethanol consumption by rats led to a variety of pathologic changes in liver, including hepatomegaly with fat accumulation in liver cells, and serological changes such as increased activity of alkaline phosphatase and alanine aminotransferase, the clinical spectrum suggesting alcoholic fatty liver (Fig. 1). These changes might have been due to enhanced mobilization of free fatty acids from the adipose tissue and the increased hepatic biosynthesis of lipid as suggested previously (Lieber et al. 1965, Lieber 1992).

Chronic ethanol consumption resulted in the induction of cytochrome P450 II E1 (Fig. 2). This very likely plays an important role in the enhanced generation of reactive oxygen species by hepatic microsomes during ethanol oxidation, because it reduces dioxygen to oxygen-derived radicals and H₂O₂ (Bondy and Naderi 1994, Hodgson and Levi 1994). A correlation between the production of these oxygen derivatives (O₂, H₂O₂) and the amount of cytochrome P450 II E1 was observed in the hepatic microsomal samples of variously treated rats by Ekström and Ingelman-Sundberg (1989). However, xanthine oxidase, another enzyme of the ethanol-induced radical generating system, was lower in livers of rats chronically fed ethanol (Table 1). This reduction of xanthine oxidase activity was somewhat unexpected because the conversion of xanthine dehydrogenase to xanthine oxidase that occurred in ischemic tissues, possibly by partial proteolysis (Engerson et al. 1987), is known to occur from ethanol feeding also. Perhaps chronic ethanol feeding rendered the tissues unable to generate excess free radicals by blocking the conversion of xanthine dehydrogenase to xanthine oxidase via an unknown mechanism.

There was no significant difference between the group chronically fed ethanol and the pair-fed control group in the indices of oxidative liver damage, namely, TBARS and carbonyl group content. This absence of oxidative liver damage in the ethanol group led us to postulate that chronic ethanol feeding prompted the strong radical scavenging activities to counteract the oxidative stress generated probably by the cytochrome P450 system.

Glutathione is a tripeptide present in all mammalian cells; it participates in many metabolic processes for the protection of cells against free radicals and toxic metabolites. In this study, there was a noticeably higher hepatic glutathione level after chronic ethanol feeding in contrast to the lower values observed in previous reports (Fernández-Checa et al. 1991, Garcia-Ruiz et al. 1994, Hirano et al. 1992, Martensson et al. 1990). However, our values were comparable to the results in several other studies (Hassing et al. 1979, Hetu et al. 1982, Iizuka et al. 1991, Morton and Mitchell 1985, Teare et al. 1994). The discrepancies in the total GSH levels in livers of rats chronically fed ethanol might have originated from the differences in the strain of rats used and the dose, duration and route of ethanol administration among different studies. Nevertheless, the greater hepatic GSH content after chronic ethanol consumption (Table 2) is beneficial not only in protecting liver against toxic effects of activated oxygen radicals and/or lipid peroxides but also as a source of its constituent amino acids. This might be one of the reasons why there are no striking changes in plasma and tissue concentrations of those amino acids after chronic ethanol ingestion (Adibi et al. 1992).

Although hepatic mitochondrial GSH is decreased by ethanol in rats and baboons, the decrease is attenuated by supplementation of the precursor S-adenosyl methionine (Lieber et al. 1990). However, we found increased GSH due to chronic ethanol feeding without any other supplementation in rats. Perhaps this was due to the specific metabolic effects of ethanol as suggested by Porta (1997). This enhanced level of GSH could have resulted from increased net GSH synthesis as suggested by Fernández-Checa et al. (1980) or, more specifically, from the increased activities of the glutathione-synthesizing enzyme system (-glutamylcysteine synthetase and glutathione synthetase) as described by Guerri and Grisolia (1980).

As Reeds and Beckett (1996) explained, protein mass and rates of protein gain or loss in a cell are entirely dependent on the balance of synthesis and degradation. Furthermore, the nutritional status and the response of protein turnover to endocrinological changes interact in a complex way. Yet, the effects of ethanol on hepatic protein synthesis per se are not clear.

However, there are some important effects of ethanol on amino acid metabolism. Branched-chain amino acids and -amino-n-butyric acid (AANB) are increased in the plasma of alcoholics and rats chronically fed ethanol (Shaw and Lieber 1980). Increased transsulfuration of methionine to cysteine, which produces -ketobutyrate, could account for the increase in AANB. This was hypothesized to occur because of the increased hepatic requirement for glutathione. Because cysteine is the rate-limiting amino acid in glutathione synthesis, the increase in AANB may reflect an increased hepatic requirement for glutathione synthesis (Mitchell 1990).

Jahoor et al. (1995) proposed that the biosynthetic supply of GSH in liver is sufficient to withstand an inflammatory challenge in well-nourished piglets. Our rats were similarly well nourished and stressed by ethanol feeding. Accordingly, it is tempting to explain the lack of a decrease in hepatic GSH after ethanol consumption, which was seen in malnourished alcoholics or protein-deficient piglets, in the same manner. That is, the biosynthetic supply of GSH in our well-nourished rats was sufficient to cope with the stress induced by chronic ethanol consumption.

We cannot extrapolate our results to alcoholics in general because our study period lasted only 6 wk and there are innate differences between the two species. Nevertheless, severe hepatic glutathione depletion is common in alcoholics, and potential danger exists if other drugs such as acetaminophen are also taken. The enhanced metabolism of acetaminophen that results from the increased microsomal drug-metabolizing activity secondary to chronic ethanol consumption would result in fulminant hepatic failure as seen in clinical cases because of the low glutathione status. Thus it seems reasonable to think of an increased requirement of amino acids in alcoholics especially of those used to synthesize glutathione.

The maintenance of a high GSH/GSSG ratio in livers of rats chronically fed ethanol (Table 2) implies a relative ease of glutathione utilization. In other words, the higher amount of reduced glutathione in the livers of rats chronically fed ethanol appears to result in efficient glutathione recycling. Therefore, the activities of glutathione peroxidase and reductase were monitored to examine the changes in glutathione utilization efficiency caused by chronic ethanol feeding. Chronic ethanol feeding resulted in the lower activity of glutathione peroxidase with significantly higher activity of glutathione reductase. Although the increase in the activity of glutathione reductase can promote the recycling of glutathione for the active detoxification of xenobiotics, the decrease in glutathione peroxidase activity may attenuate the radical scavenging function. Nonetheless, the increase in catalase activity in the livers of ethanol-fed rats may compensate, at least in part, for the curtailed radical scavenging capacity due to lower glutathione peroxidase activity (Table 3).

For optimal glutathione reductase activity, NADPH must be supplied in adequate concentration. Thus, we have looked for alterations in glucose-6-phosphate dehydrogenase activity and found that it is decreased after chronic ethanol administration (Table 3). This decrease could be explained by the findings of Szweda et al. (1993), who described the inactivation of purified bacterial glucose-6-phosphate dehydrogenase by trans-4-hydroxy-2-nonenal, a toxic product of membrane lipid peroxidation. It was shown to interact with a lysine residue in the enzyme to give a stable secondary amine derivative. Considering the physiologic role of this enzyme in supplying NADPH, the coupling of glutathione peroxidase and reductase for glutathione recycling in the livers of rats chronically fed ethanol could be hampered by the lower glucose-6-phosphate dehydrogenase activity, regardless of the change in glutathione reductase activity (Table 3).

In this study, chronic ethanol consumption resulted in a significantly higher activity of hepatic glutathione S-transferase. Iizuka et al. (1991) reported that glutathione S-transferase activity in alcoholic fatty liver was significantly higher and that in cirrhosis, it was significantly lower than that of normal liver. In fatty liver, immunohistochemical staining for the enzyme was strongly positive in hepatocytes around intensive fatty metamorphosis. The relatively easy inducibility of this enzyme by xenobiotics and ethanol as well indicates its role in detoxification of ethanol (Friedberg et al. 1979). Furthermore, it may affect the reduction of lipid hydroperoxides to stable lipid alcohols. Therefore, the enhancement of glutathione S-transferase activity in livers of ethanol-fed rats could be considered an adaptive response protecting the tissue against ethanol-induced oxidative damage.

In conclusion, although chronic ethanol administration induces cytochrome P450 II E1 and favors microsomal free-radical generation, the net outcome of chronic ethanol feeding of well-nourished rats for 6 wk indicated minimal oxidative stress in the liver. There was a compensatory hepatic response lessening ethanol-induced oxidative stress, which seemed to result from the lower activity of xanthine oxidase and the higher activities of catalase and other antioxidative defense enzymes, especially those related to recycling and utilization of glutathione. Therefore, it appears that this response can be considered to be one of the adaptive protection mechanisms that the body adopts against the toxic effects of chronic ethanol consumption, given the condition of a nutritionally balanced diet.

	FOOTNOTES

Manuscript received 5 March 1997. Initial reviews completed 16 May 1997. Revision accepted 8 December 1997.

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