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Hepatic ethanol metabolism: Respective roles of alcohol dehydrogenase, the microsomal ethanol-oxidizing system, and catalase
ARCHIVES
OF
BIOCHEMISTRY
AND
BIOPHYSICS
175, 635-643 (1976)
Hepatic Ethanol Metabolism: Respective Roles of Alcohol Dehydrogenase, the Microsomal Ethanol-Oxidizing System, and Catalase’ ROLF TESCHKE,
YASUSHI
HASUMURA,
AND
CHARLES
S. LIEBER
Alcohol Research Laboratory of the Section of Liver Disease and Nutrition, Veterans Administration Hospital, Bronx, New York 10468, and Department of Medicine, Mount Sinai School of Medicine of the City University of New York, New York, New York 10029 Received
October 27, 1975
The respective role of alcohol dehydrogenase, of the microsomal ethanol-oxidizing system, and of catalase in ethanol metabolism was assessed quantitatively in liver slices using various inhibitors and ethanol at a final concentration of 50 mM. Pyrazole (2 rn& virtually abolished cytosolic alcohol dehydrogenase activity but inhibited ethanol metabolism in liver slices by only 50-60%. The residual pyrazole-insensitive ethanol oxidation in liver slices remained unaffected by in vitro addition of the catalase inhibitor sodium azide (1 mM). At this concentration, sodium azide completely abolished catalatic activity of catalase in liver homogenate as well as peroxidatic activity of catalase in liver slices in the presence of nn-alanine. Similarly, in vivo administration of 3-amino-1,2,4triazole, a compound which inhibits the activity of catalase but not that of the microsoma1 ethanol-oxidizing system, failed to decrease both the overall rates of ethanol oxidation and the activity of the pyrazole-insensitive pathway. Finally, butanol, a substrate and inhibitor of the microsomal ethanol-oxidizing system but not of catalaseHZ02, significantly decreased the pyrazole-insensitive ethanol metabolism in liver slices. These results indicate that alcohol dehydrogenase is responsible for half or more of ethanol metabolism by liver slices and that the microsomal ethanol-oxidizing system rather than catalase-H,O, accounts for most if not all of the alcohol dehydrogenaseindependent pathway.
It was generally assumed that ethanol metabolism proceeds exclusively via alcohol dehydrogenase (ADH),2 an enzyme of the cell sap of the hepatocyte. Indeed, this concept is satisfactory at low ethanol concentrations since the oxidation of ethanol is almost completely abolished under these conditions by pyrazole, a potent inhibitor of alcohol dehydrogenase activity (1). At ’ This work was supported in part by USPHS Grants AA 00224 and AM 12511 and VA project No. 5251-02. Portions of this study were presented at the Spring Meeting of the American Society for Pharmacology and Experimental Therapeutics, Atlantic City, April 1975. Z Abbreviations used: ADH, alcohol dehydrogenase; MEOS, microsomal ethanol-oxidizing system; DEAE-, diethyl aminoethyl.
intermediate and higher concentrations, however, ethanol metabolism becomes less sensitive to pyrazole, a finding which suggested the operation of a non-ADH-mediated pathway for ethanol metabolism (l6). Since this ADH-independent pathway was considered to account for up to 75% of the overall rates of ethanol metabolism at high ethanol concentrations (1) the question of the biochemical nature of this pathway became relevant. Recent studies have shown that in addition to ADH, ethanol can also be metabolized by the microsomal fraction of the hepatocyte which comprises the endoplasmic reticulum. This microsomal ethanol-oxidizing system (MEOS) was extensively studied (2, 3, 7, 8) and it was separated by 635
Copyright 0 1976 by Academic F’rcss, Inc. All rights of reproduction in any form reserved.
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TESCHKE,
HASUMURA
DEAE-cellulose column chromatography from both alcohol dehydrogenase and catalase (9-12). A variety of reports provided strong evidence for a significant role of the microsomal ethanol-oxidizing system in ethanol metabolism (1-6, 13). This concept, however, was challenged recently, and it has been claimed that the alcohol dehydrogenase-independent pathway of ethanol metabolism is due exclusively to catalase (14). In view of these conflicting interpretations, the present study was undertaken to clarify the respective roles of the microsoma1 ethanol-oxidizing system and catalase in ethanol metabolism. MATERIALS
AND
METHODS
Materials. The chemicals and enzymes were obtained from the following sources: NADP+ (yeast), P-NAD+ (grade III), m-isocitrate (type I), and isocitric dehydrogenase from Sigma Chemical Company, Na,-EDTA, sodium St. Louis, MO.; m.-alanine, azide, hydrogen peroxide (30%), semicarbazide hydrochloride, n-butanol, and n-glucose from Fisher Scientific Company, Fairlawn, N.J.; acetaldehyde and pyrazole from Eastman Kodak Company, Rochester, N.Y.; ethanol (dehydrated) from Publicker Industries Co., Linfeld, Pa.; 3-amino-1,2,4-triazole from Aldrich, Milwaukee, Wis.; glucose oxidase (type I) from Boehringer Mannheim Corp., New York, N.Y.; and the gas mixtures from Matheson Gas Products, East Rutherford, N.J. Animals. Male Sprague-Dawley rats (CD, Charles River Breeding Laboratories, North Wilmington, Mass.) of body weight 340-440 g were used; they were fed Purina laboratory chow ad Zibitum and had free access to tap water. When indicated, rats were pretreated with 3-amino-1,2,4-triazole (1 g/kg body weight ip in 0.9% saline) 1 h before sacrifice, whereas the corresponding littermate received 0.9% saline solution only. Procedure for the measurement of ethanol oxidation by liver slices. The animals were killed by decapitation, and liver slices of approximately 0.5-mm thickness with a wet weight of about 50-60 mg each were prepared by means of a Stadie-Riggs microtome (Arthur H. Thomas Company, Philadelphia, Pa.). Randomized liver slices of a total weight of about 500 mg were added to 50-ml Erlenmeyer flasks containing 4.5 ml of isotonic Krebs-Ringer bicarbonate buffer (pH 7.4) (151, and, when indicated, the following compounds: pyrazole, 2 mM, sodium azide, 1 mM, and nbalanine, 40 mM. Then 0.5 ml of ethanol in Krebs-Ringer bicarbonate buffer (pH 7.4) was added to achieve a final alcohol concentration of 50
AND
LIEBER
rnM in a final volume of 5 ml. With each incubation set, experiments were run in which boiled liver slices were incubated with 50 mM ethanol. The values thus obtained were used as evaporation controls and subtracted from the corresponding experimental values. The vessels were sealed with serum caps and flushed for 5 min with a gas mixture of 95% 0, and 5% CO,. The subsequent incubations were carried out at 37°C for a total of 150 min in a Dubnoff water bath shaking at 100 strokes/min. Aliquots of the incubation medium were harvested with a needle and syringe through the rubber top immediately before the start of the incubations and then at intervals of 30 min. One hundred microliters of the harvested incubation medium was added to 0.5 ml of 35% perchloric acid contained in a 25-ml glass flask designed for analysis by a Perkin-Elmer F-40 gasliquid chromatograph (16). The sample bottles were immediately closed and incubated for 20 min at 60°C in the water bath attached to the chromatograph. Aliquots of the head space gas of these flasks were then injected by an automatic electropneumatic dosing system (injection time 4 s) into the gas-liquid chromatograph, and a 2-m x 2-mm column packed with 15% polyethyleneglycol on 50-60 mesh Celite was used. Helium was employed as a carrier gas at a flow rate of 40 ml/min. The temperature was 75°C for the column and 145°C for the flash heater as well as for the hydrogen flame detector. Quantitative assessment of the ethanol remaining in the incubation medium following the incubation was achieved by the use of a Hewlett-Packard GC digital integrator (Model 3370A) connected to the gas-liquid chromatograph. Solutions with known amounts of ethanol served as standards. It was found that an excellent reproducibility could only be achieved with a gas-liquid chromatograph with an automatic electropneumatic dosing system which was equilibrated for at least 4 h before the start of the analysis. Subcellular fractionation. The rats were killed by decapitation and their livers were perfused with icecold 0.15 M KC1 through the portal vein, excised, chilled, and homogenized in three volumes of 0.15 M KC1 using a glass homogenizer with a Teflon pestle. The following steps were carried out at 0-4°C. The 25% homogenate was spun at 10,OOOgfor 30 min, and the supernatant was centrifuged at 105,OOOgfor another 30 min. The resulting supematant (cytosol) was used as enzyme source for alcohol dehydrogenase, whereas the pellet was resuspended in 0.15 M KCl, and washed microsomes were obtained by spinning this suspension at 105,OOOgfor another 30 min. Biochemical determinations. The activity of alcohol dehydrogenase was assayed in the hepatic cytosol following the method of Bonnichsen and Brink (17) with ethanol at a final concentration of 50 mM. Catalatic activity of catalase was determined by
HEPATIC
ETHANOL
measuring the disappearance at 240 nm of H,O, added to aliquots of the 25% liver homogenate and expressing the results in units according to Luck (18). The activity of the NADPH-dependent microsoma1 ethanol-oxidizing system (MEOS) was determined with washed microsomes (3 mg of protein/ flask) which were preincubated with ethanol (50 mM) and sodium azide (1 mM) for 5 min at 37°C. The reactions were initiated by addition of a NADPHgenerating system (0.4 mM NADP+, 8 mM sodium isocitrate, and 0.34 unit/ml of isocitric dehydrogenase). The incubation medium contained, in a final volume of 3.0 ml, 1.0 mM NaZ-EDTA and 5.0 mM MgCl, in 0.1 M phosphate buffer (pH 7.4). The incubations were performed in closed 50-ml Erlenmeyer flasks with center wells containing 0.6 ml of 0.015 M semicarbazide hydrochloride in 0.1 M phosphate buffer (pH 7.4), and the acetaldehyde bound to the semicarbazide after an overnight diffusion period was determined according to Lieber and DeCarli (2). When the effect of butanol was studied, this alcohol was preincubated at a final concentration of 10 mM with microsomes and ethanol (50 mM) for 5 min before the reaction was started by adding the cofactor. Under these conditions, acetaldehyde bound to the semicarbazide was determined by gas-liquid chromatography as described above for the determination of ethanol in the liver slice study (12). Aliquota (100 ~1) of the semicarbazone solution of the center wells were added to 0.5 ml of 35% perchloric acid contained in 25-ml sample bottles. The bottles were sealed and subsequently incubated for 20 min at 60°C before aliquots of the head space were injected into the gas-liquid chromatograph. Flasks to which known amounts of acetaldehyde were added were carried through the complete procedure and were used as standards. The amount of acetaldehyde produced upon incubation was measured by peak heights. Peroxidatic activity of catalase was determined in washed microsomes under assay conditions described for the NADPH-dependent microsomal ethanol-oxidizing system except that sodium azide was omitted from the incubation medium and the NADPH-generating system was replaced by a H202producing one consisting of glucose (10 mM) and glucose oxidase (1.0 wg/ml of incubation medium). Under these conditions, glucose was preincubated with microsomes and ethanol (50 mM), and the reaction was started by adding glucose oxidase. Protein concentration was determined according to the method of Lowry et al. (19). Statistical analysis. Each individual result was compared with the value of its corresponding control; the means (+SE) of individual differences were calculated and their significance was assessed by the paired Student’s t test.
637
METABOLISM RESULTS
Effect of Pyrazole, Sodium Azide, and DLAlanine on Ethanol Metabolism Ethanol was oxidized by liver slices at a rate of approximately 50-60 pmol h-’ (g of liver)-‘, when ethanol was employed at a final concentration of 50 mM. The amount of ethanol disappearance was measured 150 min following the start of the incubation at which time 2530% of the alcohol added to the incubation medium had been oxidized. To verify the linearity of the reaction, measurements were also performed during the incubation time at regular intervals of 30 min each. The rate of ethanol metabolism was linear for 150 min. To assess quantitatively the role of alcohol dehydrogenase in ethanol metabolism, liver slices were incubated with ethanol (50 mM) and, when indicated, with pyrazole added to the incubation medium at a final concentration 2 mM. At this concentration, pyrazole almost completely abolished the activity of alcohol dehydrogenase when measured in the cytosol of the hepatocyte with ethanol as substrate (Table I). Pyrazole also decreased catalatic activity in liver homogenate by 24% (Table I); under these conditions, MEOS activity was found before to be inhibited by 11% (2). In contrast to the almost complete abolition of the activity of cytosolic ADH (Table I), pyrazole decreased the rate of ethanol metabolism by only 60% (P < 0.001) in liver slices (Table II). This finding suggests that although ADH activity plays a major role in ethanol metabolism, a considerable fraction of ethanol metabolism in liver slices may be accounted for by a pathway not involving the activity of alcohol dehydrogenase. To determine to what extent the pyrazole-insensitive pathway of ethanol metabolism in liver slices is due to the activity of catalase, the catalase inhibitor sodium azide was included in the incubation medium to achieve a final concentration of 1 mM. At this concentration, sodium azide virtually abolished catalase activity in liver homogenates when assayed by its catalatic property to decompose added hy-
638
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HASUMURA
AND
TABLE EFFECT
OF PYRAZOLE
AND SODIUM
AZIDE
LIEBER
I
ON THE ACTIVITY
OF ALCOHOL
DEHYDROGENASE
ADH P
Activity (rim01 min-* liver)-‘) Control Pyrazole Azide Pyrazole
1821 60 1710 55
+ azide
+ + k -t
(g of 61 5 99 4
AND CATALASE”
Catalase
(% of control) 100 3.3 93.9 3.0
Catalatic (units (g of liver)-‘)
6065 4606 7.3 6.8
+ ? + 2
452 389 3.4 2.5
P
activity (% of control) 100 76.0 0.1 0.1
“Alcohol dehydrogenase activity was determined in the hepatic cytosol (17) with 50 mM ethanol as substrate. The data are expressed as nanomoles of NADH formed per minute per gram of liver. Catalatic activity was assessed in the liver homogenate and expressed in units (18). When indicated, pyrazole and sodium azide were used in final concentrations of 2 and 1 mM, respectively. The data for ADH and catalase activity represent means (&SE) of six experiments each. TABLE
II
EFFECT OF PYRAZOLE, SODIUM AZIDE, AND DLALANINE ON ETHANOL METABOLISM BY LIVER SLICE@
Incubation
medium
Ethanol oxidat$pcFoyl
P
liver)-‘) Control Pyrazole Pyrazole Pyrazole Pyrazole azide
+ azide + DL-alanine + nkalanine
+
55.3 22.3 26.8 58.3 23.8
+ k k + r
7.1 3.8 5.6 9.3 4.4
a Rat liver slices were incubated with ethanol (50 mM) and, when indicated, with the following compounds: pyrazole (2 mM1, sodium azide (1 mM), and m.-alanine (40 mM1. The incubations were carried out for a total of 150 min, and ethanol disappearance was measured by gas-liquid chromatography. The data represent means (&SE) of five experiments.
drogen peroxide (Table I). Sodium azide at a concentration of 1 mM had no significant effect on the activity of cytosolic ADH (Table I); it had also been shown before that it does not affect the purified microsomal ethanol oxidizing system (11). In rat liver slices, the combination of sodium azide and pyrazole failed to decrease the rates of ethanol oxidation below the level achieved by pyrazole alone (Table II). To test whether sodium azide actually entered the cells in liver slices and effectively blocked the peroxidatic activity of catalase, m-alanine at a final concentration of 40 mM was used to stimulate the oxidation of ethanol via catalase-H,O, by increasing the pro-
duction of hydrogen peroxide by peroxysoma1 n-amino acid oxidase. As expected, addition of nbalanine resulted in a stimulation of ethanol metabolism in liver slices (Table II). This in,-alanine-dependent ethanol oxidation was completely abolished by sodium azide in a concentration of 1 mM which left the pyrazole-insensitive ethanol oxidation unchanged (Table II). These findings indicate that catalase plays little if any role in ethanol metabolism under physiological experimental conditions in which no nL-alanine was added. Ethanol Oxidation following in Vivo Administration of 3-Amino-1,2,4-triazole Pretreatment with 3-amino-1,2,4-triazole (1 g/kg body weight ip) 1 h before sacrifice almost completely abolished the catalase activity of liver homogenate (tested by the catalatic property to decompose added hydrogen peroxide). By contrast, the activities of cytosolic alcohol dehydrogenase and of the microsomal ethanol-oxidizing system remained virtually unchanged (Table III). Despite an almost complete abolition of catalase activity (Table III), pretreatment with 3amino-1,2,4-triazole failed to inhibit ethanol metabolism in liver slices when compared to controls receiving saline alone (Table IV). Similarly, the rates .of the pyrazole-insensitive ethanol oxidation remained unchanged following administration of 3-amino-1,2,3-triazole (Table IV). As expected, addition of nn-alanine increased the rates of ethanol metabolism
HEPATIC
ETHANOL TABLE
EFFECT
639
METABOLISM III
OF 3-AMINO-1,2,4-TRIAZOLE TREATMENT ON THE ACTIVITIES OF ALCOHOL DEHYDROGENASE, CATALASE, AND THE MICROSOMAL ETHANOL-OXIDIZING SYSTEM’~
Enzyme activity
Treatment Saline
ADH (nmol min’ g-‘) Catalase (units g-l) MEOS (nmol mix--’ (mg of protein)-‘)
Aminotriazole
1840 ” 65 (61 6074 + 442 (6) 10.0 k 0.6 03)
1880 c 75 (6) 267 t 44 (6) 10.3 2 0.5 (8)
Ratio (aminotriatole/ saline)
P
1.04 0.04 1.03
N.S.
o Rats were pretreated with 3-amino-1,2,4-triazole (1 g/kg body weight ip in physiological saline) or with physiological saline alone. Alcohol dehydrogenase (ADH) activity was determined in the hepatic cytosol(17) with 50 mM ethanol as substrate and expressed as nanomoles of NADH formed per minute per gram of liver. Catalatic activity was assessed in liver homogenate and expressed in units (18). The activity of the microsomal ethanol-oxidizing system (MEOS) was determined in washed microsomes with an NADPHgenerating system, 50 mM ethanol, 1 mM Na,-EDTA, 5 mM MgCl, and 1 mM sodium azide and expressed in nanomoles of acetaldehyde formed per minute per milligram of microsomal protein. The data represent means (*SE); the numbers of experiments are indicated in parentheses. TABLE EFFECT
IV
OF S-AMINO-1,2,4-TRIAZOLE METABOLISM BY LIVER
Incubation medium
Control Pyrazole Pyrazole + DLalanine
ON ETHANOL SLICE@
Ethanol oxidation (pm01 h-’ (g of liver)-‘) Aminotriazole
Saline
52.9 k 6.2 23.7 k 2.0 24.1 ? 2.7
49.8 2 5.9 24.5 k 2.1 39.5 c 4.1
P
N.S. N.S.
D Rats were pretreated with 3-amino-1,2,4-triazole at a dose of 1 g/kg body weight (in physiological saline ip) 1 h before sacrifice, whereas the corresponding littermate received saline solution alone. Rat liver slices were incubated for 150 min with ethanol (50 mM) and, when indicated, with pyrazole (2 mM) and nn-alanine (40 mM). Ethanol disappearance was measured by gas-liquid chromatography. The values represent means (*SE) derived from five experiments.
in liver slices of control rats (Table IV). By contrast, no such enhancement by m-alanine was observed in liver slices derived from rats pretreated with 3-amino-1,2,4triazole (Table IV), indicating that the catalase-mediated peroxidation of ethanol was effectively blocked. These results therefore provide strong evidence against a significant role of catalase in the ADHindependent ethanol oxidation. Effect of Butanol
on Ethanol
Metabolism
In view of our previous finding that higher aliphatic alcohols are substrates for the NADPH-dependent microsomal alcohol-oxidizing system but not for catalase
(7), we tested whether butanol might affect ethanol oxidation by the NADPH-dependent microsomal system as well as by liver slices. To that effect, liver microsomes were prepared and incubated with an NADPH-generating system, ethanol, and, when indicated, butanol. Since both acetaldehyde and butyraldehyde are formed during the oxidation of the respective alcohol and, when bound to the semicarbazide, both elicit changes of the absorbance at 224 nm (12), acetaldehyde formation had to be assessed by gas-liquid chromatography rather than by spectrophotometric assay. Upon analysis by gasliquid chromatography the retention time of acetaldehyde was sufficiently different not only from the one of butyraldehyde but also of ethanol and butanol employed as substrates in the assay system. In the presence of butanol (10 mM), the activity of the microsomal ethanol-oxidizing system was significantly inhibited by 46.9% (P < O.Ol), whereas this alcohol had no significant effect on the catalase-mediated ethanol oxidation (Table V). Similarly, the pyrazole-insensitive ethanol oxidation in liver slices was decreased by 43.2% (P < 0.01) in the presence of 10 mM butanol (Table VI), suggesting that the microsoma1 alcohol-oxidizing system is involved in the pyrazole-insensitive ethanol metabolism in liver slices. To evaluate whether the activity of MEOS could account quantitatively for the ADH-independent pathway, MEOS activity was compared with
640
TESCHKE,
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AND
TABLE
LIEBER
V
EFFECT OF BUTANOL ON THE ACTIVITY OF THE NADPH-DEPENDENT MICROSOMAL ETHANOL-OXIDIZING SYSTEM AND ON THE PEROXIDATIC ACTIVITY OF CATALAS@ Substrate Microsomal ethanol oxidizing system Catalase Activity
Ethanol Ethanol
+ butanol
(rim01 of acetaldehyde min-’ (mg of protein)-’
(% of controll
8.1 k 0.9 4.3 2 0.5
100 53.1
Peroxidatic P
activity
(nmol of acetaldehyde min-’ (mg of protein)-‘)
(% of controll
10.8 2 0.4 11.2 2 0.3
100 103.7
co.01
P
N.S.
a The activity of the microsomal ethanol oxidizing system (MEOS) was measured by incubating washed microsomes (3 mg of protein/flask) with ethanol (50 mM), sodium azide (1 mM) and the NADPH-generating system in a medium containing 1 mM Na,-EDTA, 5 mM MgCl,, and 0.1 M phosphate buffer (pH 7.4). When indicated, n-butanol was added at a final concentration of 10 mM. After an overnight diffusion period, acetaldehyde bound to the semicarbazide of the center well was assessed quantitatively by gas-liquid chromatography. Peroxidatic activity of catalase was determined in washed microsomes under similar assay conditions, except that sodium azide was omitted from the incubation medium and the NADPH-generating system was replaced by the H,Oz-producing one. The value of each enzyme activity represents the mean (&SE) of five experiments.
the rates of the pyrazole-insensitive ethanol oxidation by liver slices. Ethanol is oxidized by MEOS in vitro at a rate of 810 nmol min-’ (mg of microsomal protein)-’ (Table III and V), corresponding to 480-600 nmol h-’ (mg of protein)-*. When corrected for microsomal losses during the subcellular preparation, 1 g of liver contains approximately 40 mg of microsomal protein (21); consequently, about 24 pmol of ethanol h-’ (g of liver)-’ could be oxidized by MEOS in vitro, a value which is similar to the rates of the pyrazole-insensitive ethanol oxidation in liver slices of 2229 pmol h-l (g of liver)-’ (Tables II, IV, VI). DISCUSSION
The present study shows that in the presence of 2 mM pyrazole which virtually abolishes alcohol dehydrogenase activity in the hepatic cytosol (Table I) ethanol metabolism in rat liver slices was reduced by only 50-60%, using ethanol at a final concentration of 50 mM (Tables II, IV, and VI); the residual pyrazole-insensitive pathway remained unaffected by both in vitro addition of the catalase inhibitor sodium azide (Table II) or in vim administration of the catalase inhibitor 3-amino1,2,4-triazole (Table IV) but was significantly decreased by butanol (Table VI), an inhibitor of the activity of the microsomal
TABLE VI EFFECT OF n-BUTANOL ON ETHANOL METABOLISM BY LIVER SLICES Incubation medium
Control Pyrazole Pyrazole
+ butanol
Ethanol oxidation (pm01 h-’ (g of liver)-‘) 54.1 t 5.5 29.4 -c 3.6 18.1 2 1.9
P
-co.01
n Rat liver slices were incubated for 150 min with ethanol (50 mM) and, when indicated, with pyrazole (2 mM) and n-butanol (10 mru). Ethanol disappearance was assessed by gas-liquid chromatography. The data are derived from four experiments and are expressed as means (GE).
ethanol-oxidizing system (Table V). It is concluded that, whereas alcohol dehydrogenase is responsible for half or more of ethanol metabolism, the microsomal ethanol-oxidizing system rather than catalase may account for most if not all of the residual alcohol dehydrogenase-independent pathway. The demonstration that ethanol metabolism is partially inhibited by pyrazole (Tables II, IV, and VI) is in agreement with other studies (l-6) and confirms the concept of a significant role of alcohol dehydrogenase in ethanol metabolism. The residual pyrazole-insensitive pathway was ascribed to an enzymic process independent of the activity of alcohol dehydrogenase rather than to incomplete inhibition of
HEPATIC
ETHANOL
alcohol dehydrogenase activity (1, 3, 5, 22, 23). Indeed, in the presence of pyrazole, glucose labeling from [ 1R-3Hlethanol was nearly abolished, while H3H0 production was inhibited less than 50%. In view of the stereospecificity of ADH for [lR3Hlethanol, these findings suggest “the presence of a significant pathway not mediated by cytosolic ADH” (22). Additional evidence that this residual pyrazole-insensitive ethanol metabolism is not ADH mediated was derived from the fact that the cytosolic redox state was unaffected when ethanol was administered together with pyrazole compared to saline pretreated rats (23). Furthermore, compared to ethanol concentrations of 10 mM at which ADH is fully saturated the rates of alcohol metabolism are significantly higher at ethanol concentrations above 25 mM (24). Moreover, under experimental conditions comparable to those of this study (Tables II, IV, and VI), the K, of the pyrazoleinsensitive ethanol oxidation in isolated hepatocytes has been determined to be 13 mM for ethanol (24). This value is one order of magnitude higher than the corresponding value for alcohol dehydrogenase (25, 26). In addition, the pyrazole-insensitive ethanol metabolism was significantly greater at 25 mM ethanol than at 10 mM (24). The inhibition was also not significantly enhanced when the pyrazole concentration was increased from 2 to 4 mM (Matsuzaki and Lieber, unpublished observation). All these findings argue against the possibility that the pyrazoleinsensitive ethanol oxidation observed in the present study is due to incomplete inhibition of alcohol dehydrogenase. Of particular interest was the finding in the present study that butanol inhibits both the pyrazole-insensitive ethanol metabolism by liver slices (Table VI) as well as the activity of the NADPH-dependent microsomal ethanol-oxidizing system (Table V). The latter finding is in keeping with the earlier observation that butanol is a good substrate for the microsomal system (12). These observations strongly suggest the involvement of the microsomal ethanol-oxidizing system in the residual pyrazole-insensitive ethanol metabolism. Consistent with this hypothesis is the ob-
METABOLISM
641
servation that the pyrazole-insensitive ethanol metabolism in liver slices remained unaffected by in vitro addition of sodium azide (Table II) or in viva administration of 3-amino-1,2,4-triazole (Table IV) since the activity of the microsomal ethanol-oxidizing system was found to be insensitive to sodium azide (11) and to 3amino-1,2,4-triazole (Table III). Similarly, the striking increase in the non-ADH fraction of ethanol metabolism with increasing ethanol concentrations (1, 4) is consistent with the known K, of ADH and MEOS: whereas the former has a K, varying from 0.5 to 2 mM for ethanol (25, 261, the latter has a value of 8-9 mM (2). The in vitro K, of MEOS agrees well with the corresponding in uivo value of 8.8 mM for ethanol for the pyrazole-insensitive pathway (3). Furthermore, phenobarbital pretreatment as well as chronic ethanol consumption increased total MEOS activity in the liver, and this was accompanied by an acceleration of blood ethanol clearance, suggesting that the latter might be due, at least in part, to the increased activity of the microsomal system (3). Strong evidence for a microsomal component in ethanol metabolism has been provided by other studies (1, 4-6, 13, 23). In particular, it has been pointed out that the rate of ethanol utilization in perfused livers was much less in the fasted than in the fed state (13). This finding was considered to be consistent with a lack of NADPH generation due to depletion of glycogen in the fasted state, resulting in an insufficient supply of NADPH (13) the cofactor for the activity of the microsomal ethanol-oxidizing system (2). This discrepancy in metabolic rates of ethanol oxidation between the fasted and the fed state persisted even in the presence of mitochondrial inhibitors, supporting the view that an extramitochondrial mechanism was involved (13). Similar results were obtained with aminopyrine, another substrate for the NADPH-dependent microsomal drug-detoxifying enzyme system (13). It is also noteworthy that the rate of ethanol oxidation is strikingly enhanced in patients with glycogenosis type I (27). This finding was explained on the basis that in this particular disorder the content of hepatic glucose 6-phosphate is in-
642
TESCHKE,
HASUMURA
creased, which results in an enhanced rate of generation of NADPH (271, the cofactor of the microsomal ethanol-oxidizing system (2). Thus, the present study supports the thesis that the microsomal ethanoloxidizing system may play a significant role in ethanol metabolism in uivo (3) as well as in vitro in perfused liver (6, 131, isolated parenchymal liver cells (1, 51, and liver slices (1, 2, 4). The data presented in this study do rule out a significant role of catalase in ethanol metabolism, in agreement with other reports (5,6,28-32). This is supported by the observation that the pyrazole-insensitive ethanol metabolism in liver slices remained unaffected by in vitro addition of 1 mM sodium azide (Table II). At this concentration, this inhibitor abolished both the catalatic activity of catalase in liver homogenate (Table I) and the catalase-mediated peroxidation of ethanol in liver slices in the presence of 40 mM nn-alanine (Table II), a compound which stimulates peroxysomal H,O, generation via n-amino acid oxidase (33). Furthermore, in uiuo administration of 3-amino-1,2,4-triazole abolished the m-alanine-mediated peroxidation of ethanol via catalase in liver slices (Table IV) and inhibited the catalatic activity of catalase in liver homogenate by 96% (Table III), but this treatment failed to inhibit both the overall rate of ethanol oxidation in liver slices and the pyrazole-insensitive pathway (Table IV). Finally, the observation in the present study that butanol inhibits the pyrazoleinsensitive ethanol oxidation in liver slices (Table VI) without affecting ethanol peroxidation via catalase-H,O, (Table V) indicates that catalase cannot account for the pyrazole-insensitive ethanol metabolism. Supportive evidence against a significant role of catalase in ethanol metabolism was provided by others who reported an extremely low capacity of the liver to generate H,O, (341, which limits the rate of ethanol oxidation via catalase. From these as well as our studies we conclude that catalase plays little if any role in ethanol metabolism and that in addition to alcohol dehydrogenase the microsomal ethanol-
AND
LIEBER
oxidizing system accounts for a significant fraction of the overall ethanol metabolism. ACKNOWLEDGMENTS The authors are grateful to Miss Nancy Lowe for her excellent technical assistance. REFERENCES 1. GRUNNET, N., QUISTORFF, B., AND THIEDEN, H. I. D. (1973) Eur. J. Biochem. 40, 275-282. 2. LIEBER, C. S., AND DECARLI, L. M. (1970) J. Biol. Chem. 245, 25052512. 3. LIEBER, C. S., AND DECARLI, L. M. (1972) J. Pharmacol. Exp. Ther. 181, 279-287. 4. THIEDEN, H. I. D. (1971) Acta Chem. Sand. 25, 3421-3427. 5. ROGNSTAD, R. (1974) Arch. B&hem. Biophys. 163, 544-551. 6. PAPENBERG, J., VON WARTBURG, J. P., AND AEBI, H. (1970) Enzymol. Biol. Clin. 11, 237250. 7. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S. (1974) Biochem. Biophys. Res. Commun. 60, 851-857. 8. HILDEBRANDT, A. G., SPECK, M., AND ROOTS, I. (1974) Biochem. Biophys. Res. Commun. 60, 851-857. 9. TESCHKE, R., HASUMURA, Y., JOLY, J.-G., ISHII, H., AND LIEBER, C. S. (1972) Biochem. Biophys. Res. Commun. 49, 1187-1193. 10. MEZEY, E., POTTER, J. J., AND REED, W. D. (1973) J. Biol. Chem. 248, 1183-1187. 11. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S. (1974) Arch. Biochem. Biophys. 163, 404-415. 12. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S (1975) J. Biol. Chem. 250, 7397-7404. 13. SCHOLZ, R., HANSEN, W., AND THURMAN, R. G. (1971) in Metabolic Changes Induced by Alcohol (Martini, G. A., and Bode, Ch., eds.), pp. 101-107, Springer-Verlag, Berlin. 14. THURMAN, R. G., AND MCKENNA, W. (1974) Hoppe-Seyler’s Z. Physiol. Chem. 355, 336340. 15. DE LUCA, H. F., AND COHEN, P. P. (1964) in Manometric Techniques (Umbreit, W. W., Burris, R. H., and Stauffer, J. F., eds.), pp. 131-133, Burgess, Minneapolis. 16. KORSTEN, M. A., MATSUZAKI, S., FEINMAN, L., AND LIEBER, C. S. (1975) N. Engl. J. Med. 292, 386-389. 17. BONNICHSEN, R. K., AND BRINK, N. G. (1955) in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 1, pp. 495-500, Academic Press, New York. 18. LOCK, H. (1963) in Methods of Enzymatic Analysis (Bergmeyer, H. U., ed.). pp. 885-894, Academic Press, New York.
HEPATIC
ETHANOL
19. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275. 20. ARSLANIAN, M. J., PASCOE, E., AND REINHOLD, J. G. (1971) &o&em. J. 125, 1039-1047. 21. JOLY, J.-G., FEINMAN, L., ISHII, H., AND LIEBER, C. S. (1973) J. L@pid Res. 14, 337-343. 22. ROGNSTAD, R., AND CLARK, D. G. (1974) Eur. J. Biochem. 42, 51-60. 23. GRUNNET, N., AND THIEDEN, H. I. D. (1972) Life sci. 11, 983-993. 24. MATSUZAKI, S., AND LIEBER, C. S. (1975) Gustroenterology 69, 845. 25. MAKAR, A. B., AND MANNERING, G. J. (1970) Biochem. Pharmucol. 19, 2017-2022. 26. REYXIER, M. (1969)Actu Chem. Stand. 23,11191129.
METABOLISM
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27. PAPENBERG, J. (1971) in Alcohol and the Liver (Gerok, W., Sickinger, K., and Hennekeuser, H. H., eds.), pp. 45-47, F. K. Schattauer Verlag, Stuttgart/New York. 28. BARTLETT, G. R. (1952) Quart. J. Stud. Ale. 13, 583-589. 29. KINARD, F. W., NELSON, G. H., AND HAY, M. G. (1956) Proc. Sot. Exp. Biol. Med. 92, 772-773. 30. SMITH, M. E. (1961) J. Pharmucol. 134,233-237. 31. LESTER, D., AND BENSON, G. D. (1970) Science 169, 282-284. 32. FEYTMANS, E., AND LEIGHTON, F. (1973) Biothem. Phurmucol. 22, 349-360. 33. DE DUVE, C., AND BAUDHUIN, P. (1966) Annu. Rev. Physiol. 46, 323-357. 34. BOVERIS, A., OSHINO, N., AND CHANCE, B. (1972) Biochem. J. 128, 617-630.
OF
BIOCHEMISTRY
AND
BIOPHYSICS
175, 635-643 (1976)
Hepatic Ethanol Metabolism: Respective Roles of Alcohol Dehydrogenase, the Microsomal Ethanol-Oxidizing System, and Catalase’ ROLF TESCHKE,
YASUSHI
HASUMURA,
AND
CHARLES
S. LIEBER
Alcohol Research Laboratory of the Section of Liver Disease and Nutrition, Veterans Administration Hospital, Bronx, New York 10468, and Department of Medicine, Mount Sinai School of Medicine of the City University of New York, New York, New York 10029 Received
October 27, 1975
The respective role of alcohol dehydrogenase, of the microsomal ethanol-oxidizing system, and of catalase in ethanol metabolism was assessed quantitatively in liver slices using various inhibitors and ethanol at a final concentration of 50 mM. Pyrazole (2 rn& virtually abolished cytosolic alcohol dehydrogenase activity but inhibited ethanol metabolism in liver slices by only 50-60%. The residual pyrazole-insensitive ethanol oxidation in liver slices remained unaffected by in vitro addition of the catalase inhibitor sodium azide (1 mM). At this concentration, sodium azide completely abolished catalatic activity of catalase in liver homogenate as well as peroxidatic activity of catalase in liver slices in the presence of nn-alanine. Similarly, in vivo administration of 3-amino-1,2,4triazole, a compound which inhibits the activity of catalase but not that of the microsoma1 ethanol-oxidizing system, failed to decrease both the overall rates of ethanol oxidation and the activity of the pyrazole-insensitive pathway. Finally, butanol, a substrate and inhibitor of the microsomal ethanol-oxidizing system but not of catalaseHZ02, significantly decreased the pyrazole-insensitive ethanol metabolism in liver slices. These results indicate that alcohol dehydrogenase is responsible for half or more of ethanol metabolism by liver slices and that the microsomal ethanol-oxidizing system rather than catalase-H,O, accounts for most if not all of the alcohol dehydrogenaseindependent pathway.
It was generally assumed that ethanol metabolism proceeds exclusively via alcohol dehydrogenase (ADH),2 an enzyme of the cell sap of the hepatocyte. Indeed, this concept is satisfactory at low ethanol concentrations since the oxidation of ethanol is almost completely abolished under these conditions by pyrazole, a potent inhibitor of alcohol dehydrogenase activity (1). At ’ This work was supported in part by USPHS Grants AA 00224 and AM 12511 and VA project No. 5251-02. Portions of this study were presented at the Spring Meeting of the American Society for Pharmacology and Experimental Therapeutics, Atlantic City, April 1975. Z Abbreviations used: ADH, alcohol dehydrogenase; MEOS, microsomal ethanol-oxidizing system; DEAE-, diethyl aminoethyl.
intermediate and higher concentrations, however, ethanol metabolism becomes less sensitive to pyrazole, a finding which suggested the operation of a non-ADH-mediated pathway for ethanol metabolism (l6). Since this ADH-independent pathway was considered to account for up to 75% of the overall rates of ethanol metabolism at high ethanol concentrations (1) the question of the biochemical nature of this pathway became relevant. Recent studies have shown that in addition to ADH, ethanol can also be metabolized by the microsomal fraction of the hepatocyte which comprises the endoplasmic reticulum. This microsomal ethanol-oxidizing system (MEOS) was extensively studied (2, 3, 7, 8) and it was separated by 635
Copyright 0 1976 by Academic F’rcss, Inc. All rights of reproduction in any form reserved.
636
TESCHKE,
HASUMURA
DEAE-cellulose column chromatography from both alcohol dehydrogenase and catalase (9-12). A variety of reports provided strong evidence for a significant role of the microsomal ethanol-oxidizing system in ethanol metabolism (1-6, 13). This concept, however, was challenged recently, and it has been claimed that the alcohol dehydrogenase-independent pathway of ethanol metabolism is due exclusively to catalase (14). In view of these conflicting interpretations, the present study was undertaken to clarify the respective roles of the microsoma1 ethanol-oxidizing system and catalase in ethanol metabolism. MATERIALS
AND
METHODS
Materials. The chemicals and enzymes were obtained from the following sources: NADP+ (yeast), P-NAD+ (grade III), m-isocitrate (type I), and isocitric dehydrogenase from Sigma Chemical Company, Na,-EDTA, sodium St. Louis, MO.; m.-alanine, azide, hydrogen peroxide (30%), semicarbazide hydrochloride, n-butanol, and n-glucose from Fisher Scientific Company, Fairlawn, N.J.; acetaldehyde and pyrazole from Eastman Kodak Company, Rochester, N.Y.; ethanol (dehydrated) from Publicker Industries Co., Linfeld, Pa.; 3-amino-1,2,4-triazole from Aldrich, Milwaukee, Wis.; glucose oxidase (type I) from Boehringer Mannheim Corp., New York, N.Y.; and the gas mixtures from Matheson Gas Products, East Rutherford, N.J. Animals. Male Sprague-Dawley rats (CD, Charles River Breeding Laboratories, North Wilmington, Mass.) of body weight 340-440 g were used; they were fed Purina laboratory chow ad Zibitum and had free access to tap water. When indicated, rats were pretreated with 3-amino-1,2,4-triazole (1 g/kg body weight ip in 0.9% saline) 1 h before sacrifice, whereas the corresponding littermate received 0.9% saline solution only. Procedure for the measurement of ethanol oxidation by liver slices. The animals were killed by decapitation, and liver slices of approximately 0.5-mm thickness with a wet weight of about 50-60 mg each were prepared by means of a Stadie-Riggs microtome (Arthur H. Thomas Company, Philadelphia, Pa.). Randomized liver slices of a total weight of about 500 mg were added to 50-ml Erlenmeyer flasks containing 4.5 ml of isotonic Krebs-Ringer bicarbonate buffer (pH 7.4) (151, and, when indicated, the following compounds: pyrazole, 2 mM, sodium azide, 1 mM, and nbalanine, 40 mM. Then 0.5 ml of ethanol in Krebs-Ringer bicarbonate buffer (pH 7.4) was added to achieve a final alcohol concentration of 50
AND
LIEBER
rnM in a final volume of 5 ml. With each incubation set, experiments were run in which boiled liver slices were incubated with 50 mM ethanol. The values thus obtained were used as evaporation controls and subtracted from the corresponding experimental values. The vessels were sealed with serum caps and flushed for 5 min with a gas mixture of 95% 0, and 5% CO,. The subsequent incubations were carried out at 37°C for a total of 150 min in a Dubnoff water bath shaking at 100 strokes/min. Aliquots of the incubation medium were harvested with a needle and syringe through the rubber top immediately before the start of the incubations and then at intervals of 30 min. One hundred microliters of the harvested incubation medium was added to 0.5 ml of 35% perchloric acid contained in a 25-ml glass flask designed for analysis by a Perkin-Elmer F-40 gasliquid chromatograph (16). The sample bottles were immediately closed and incubated for 20 min at 60°C in the water bath attached to the chromatograph. Aliquots of the head space gas of these flasks were then injected by an automatic electropneumatic dosing system (injection time 4 s) into the gas-liquid chromatograph, and a 2-m x 2-mm column packed with 15% polyethyleneglycol on 50-60 mesh Celite was used. Helium was employed as a carrier gas at a flow rate of 40 ml/min. The temperature was 75°C for the column and 145°C for the flash heater as well as for the hydrogen flame detector. Quantitative assessment of the ethanol remaining in the incubation medium following the incubation was achieved by the use of a Hewlett-Packard GC digital integrator (Model 3370A) connected to the gas-liquid chromatograph. Solutions with known amounts of ethanol served as standards. It was found that an excellent reproducibility could only be achieved with a gas-liquid chromatograph with an automatic electropneumatic dosing system which was equilibrated for at least 4 h before the start of the analysis. Subcellular fractionation. The rats were killed by decapitation and their livers were perfused with icecold 0.15 M KC1 through the portal vein, excised, chilled, and homogenized in three volumes of 0.15 M KC1 using a glass homogenizer with a Teflon pestle. The following steps were carried out at 0-4°C. The 25% homogenate was spun at 10,OOOgfor 30 min, and the supernatant was centrifuged at 105,OOOgfor another 30 min. The resulting supematant (cytosol) was used as enzyme source for alcohol dehydrogenase, whereas the pellet was resuspended in 0.15 M KCl, and washed microsomes were obtained by spinning this suspension at 105,OOOgfor another 30 min. Biochemical determinations. The activity of alcohol dehydrogenase was assayed in the hepatic cytosol following the method of Bonnichsen and Brink (17) with ethanol at a final concentration of 50 mM. Catalatic activity of catalase was determined by
HEPATIC
ETHANOL
measuring the disappearance at 240 nm of H,O, added to aliquots of the 25% liver homogenate and expressing the results in units according to Luck (18). The activity of the NADPH-dependent microsoma1 ethanol-oxidizing system (MEOS) was determined with washed microsomes (3 mg of protein/ flask) which were preincubated with ethanol (50 mM) and sodium azide (1 mM) for 5 min at 37°C. The reactions were initiated by addition of a NADPHgenerating system (0.4 mM NADP+, 8 mM sodium isocitrate, and 0.34 unit/ml of isocitric dehydrogenase). The incubation medium contained, in a final volume of 3.0 ml, 1.0 mM NaZ-EDTA and 5.0 mM MgCl, in 0.1 M phosphate buffer (pH 7.4). The incubations were performed in closed 50-ml Erlenmeyer flasks with center wells containing 0.6 ml of 0.015 M semicarbazide hydrochloride in 0.1 M phosphate buffer (pH 7.4), and the acetaldehyde bound to the semicarbazide after an overnight diffusion period was determined according to Lieber and DeCarli (2). When the effect of butanol was studied, this alcohol was preincubated at a final concentration of 10 mM with microsomes and ethanol (50 mM) for 5 min before the reaction was started by adding the cofactor. Under these conditions, acetaldehyde bound to the semicarbazide was determined by gas-liquid chromatography as described above for the determination of ethanol in the liver slice study (12). Aliquota (100 ~1) of the semicarbazone solution of the center wells were added to 0.5 ml of 35% perchloric acid contained in 25-ml sample bottles. The bottles were sealed and subsequently incubated for 20 min at 60°C before aliquots of the head space were injected into the gas-liquid chromatograph. Flasks to which known amounts of acetaldehyde were added were carried through the complete procedure and were used as standards. The amount of acetaldehyde produced upon incubation was measured by peak heights. Peroxidatic activity of catalase was determined in washed microsomes under assay conditions described for the NADPH-dependent microsomal ethanol-oxidizing system except that sodium azide was omitted from the incubation medium and the NADPH-generating system was replaced by a H202producing one consisting of glucose (10 mM) and glucose oxidase (1.0 wg/ml of incubation medium). Under these conditions, glucose was preincubated with microsomes and ethanol (50 mM), and the reaction was started by adding glucose oxidase. Protein concentration was determined according to the method of Lowry et al. (19). Statistical analysis. Each individual result was compared with the value of its corresponding control; the means (+SE) of individual differences were calculated and their significance was assessed by the paired Student’s t test.
637
METABOLISM RESULTS
Effect of Pyrazole, Sodium Azide, and DLAlanine on Ethanol Metabolism Ethanol was oxidized by liver slices at a rate of approximately 50-60 pmol h-’ (g of liver)-‘, when ethanol was employed at a final concentration of 50 mM. The amount of ethanol disappearance was measured 150 min following the start of the incubation at which time 2530% of the alcohol added to the incubation medium had been oxidized. To verify the linearity of the reaction, measurements were also performed during the incubation time at regular intervals of 30 min each. The rate of ethanol metabolism was linear for 150 min. To assess quantitatively the role of alcohol dehydrogenase in ethanol metabolism, liver slices were incubated with ethanol (50 mM) and, when indicated, with pyrazole added to the incubation medium at a final concentration 2 mM. At this concentration, pyrazole almost completely abolished the activity of alcohol dehydrogenase when measured in the cytosol of the hepatocyte with ethanol as substrate (Table I). Pyrazole also decreased catalatic activity in liver homogenate by 24% (Table I); under these conditions, MEOS activity was found before to be inhibited by 11% (2). In contrast to the almost complete abolition of the activity of cytosolic ADH (Table I), pyrazole decreased the rate of ethanol metabolism by only 60% (P < 0.001) in liver slices (Table II). This finding suggests that although ADH activity plays a major role in ethanol metabolism, a considerable fraction of ethanol metabolism in liver slices may be accounted for by a pathway not involving the activity of alcohol dehydrogenase. To determine to what extent the pyrazole-insensitive pathway of ethanol metabolism in liver slices is due to the activity of catalase, the catalase inhibitor sodium azide was included in the incubation medium to achieve a final concentration of 1 mM. At this concentration, sodium azide virtually abolished catalase activity in liver homogenates when assayed by its catalatic property to decompose added hy-
638
TESCHKE,
HASUMURA
AND
TABLE EFFECT
OF PYRAZOLE
AND SODIUM
AZIDE
LIEBER
I
ON THE ACTIVITY
OF ALCOHOL
DEHYDROGENASE
ADH P
Activity (rim01 min-* liver)-‘) Control Pyrazole Azide Pyrazole
1821 60 1710 55
+ azide
+ + k -t
(g of 61 5 99 4
AND CATALASE”
Catalase
(% of control) 100 3.3 93.9 3.0
Catalatic (units (g of liver)-‘)
6065 4606 7.3 6.8
+ ? + 2
452 389 3.4 2.5
P
activity (% of control) 100 76.0 0.1 0.1
“Alcohol dehydrogenase activity was determined in the hepatic cytosol (17) with 50 mM ethanol as substrate. The data are expressed as nanomoles of NADH formed per minute per gram of liver. Catalatic activity was assessed in the liver homogenate and expressed in units (18). When indicated, pyrazole and sodium azide were used in final concentrations of 2 and 1 mM, respectively. The data for ADH and catalase activity represent means (&SE) of six experiments each. TABLE
II
EFFECT OF PYRAZOLE, SODIUM AZIDE, AND DLALANINE ON ETHANOL METABOLISM BY LIVER SLICE@
Incubation
medium
Ethanol oxidat$pcFoyl
P
liver)-‘) Control Pyrazole Pyrazole Pyrazole Pyrazole azide
+ azide + DL-alanine + nkalanine
+
55.3 22.3 26.8 58.3 23.8
+ k k + r
7.1 3.8 5.6 9.3 4.4
a Rat liver slices were incubated with ethanol (50 mM) and, when indicated, with the following compounds: pyrazole (2 mM1, sodium azide (1 mM), and m.-alanine (40 mM1. The incubations were carried out for a total of 150 min, and ethanol disappearance was measured by gas-liquid chromatography. The data represent means (&SE) of five experiments.
drogen peroxide (Table I). Sodium azide at a concentration of 1 mM had no significant effect on the activity of cytosolic ADH (Table I); it had also been shown before that it does not affect the purified microsomal ethanol oxidizing system (11). In rat liver slices, the combination of sodium azide and pyrazole failed to decrease the rates of ethanol oxidation below the level achieved by pyrazole alone (Table II). To test whether sodium azide actually entered the cells in liver slices and effectively blocked the peroxidatic activity of catalase, m-alanine at a final concentration of 40 mM was used to stimulate the oxidation of ethanol via catalase-H,O, by increasing the pro-
duction of hydrogen peroxide by peroxysoma1 n-amino acid oxidase. As expected, addition of nbalanine resulted in a stimulation of ethanol metabolism in liver slices (Table II). This in,-alanine-dependent ethanol oxidation was completely abolished by sodium azide in a concentration of 1 mM which left the pyrazole-insensitive ethanol oxidation unchanged (Table II). These findings indicate that catalase plays little if any role in ethanol metabolism under physiological experimental conditions in which no nL-alanine was added. Ethanol Oxidation following in Vivo Administration of 3-Amino-1,2,4-triazole Pretreatment with 3-amino-1,2,4-triazole (1 g/kg body weight ip) 1 h before sacrifice almost completely abolished the catalase activity of liver homogenate (tested by the catalatic property to decompose added hydrogen peroxide). By contrast, the activities of cytosolic alcohol dehydrogenase and of the microsomal ethanol-oxidizing system remained virtually unchanged (Table III). Despite an almost complete abolition of catalase activity (Table III), pretreatment with 3amino-1,2,4-triazole failed to inhibit ethanol metabolism in liver slices when compared to controls receiving saline alone (Table IV). Similarly, the rates .of the pyrazole-insensitive ethanol oxidation remained unchanged following administration of 3-amino-1,2,3-triazole (Table IV). As expected, addition of nn-alanine increased the rates of ethanol metabolism
HEPATIC
ETHANOL TABLE
EFFECT
639
METABOLISM III
OF 3-AMINO-1,2,4-TRIAZOLE TREATMENT ON THE ACTIVITIES OF ALCOHOL DEHYDROGENASE, CATALASE, AND THE MICROSOMAL ETHANOL-OXIDIZING SYSTEM’~
Enzyme activity
Treatment Saline
ADH (nmol min’ g-‘) Catalase (units g-l) MEOS (nmol mix--’ (mg of protein)-‘)
Aminotriazole
1840 ” 65 (61 6074 + 442 (6) 10.0 k 0.6 03)
1880 c 75 (6) 267 t 44 (6) 10.3 2 0.5 (8)
Ratio (aminotriatole/ saline)
P
1.04 0.04 1.03
N.S.
o Rats were pretreated with 3-amino-1,2,4-triazole (1 g/kg body weight ip in physiological saline) or with physiological saline alone. Alcohol dehydrogenase (ADH) activity was determined in the hepatic cytosol(17) with 50 mM ethanol as substrate and expressed as nanomoles of NADH formed per minute per gram of liver. Catalatic activity was assessed in liver homogenate and expressed in units (18). The activity of the microsomal ethanol-oxidizing system (MEOS) was determined in washed microsomes with an NADPHgenerating system, 50 mM ethanol, 1 mM Na,-EDTA, 5 mM MgCl, and 1 mM sodium azide and expressed in nanomoles of acetaldehyde formed per minute per milligram of microsomal protein. The data represent means (*SE); the numbers of experiments are indicated in parentheses. TABLE EFFECT
IV
OF S-AMINO-1,2,4-TRIAZOLE METABOLISM BY LIVER
Incubation medium
Control Pyrazole Pyrazole + DLalanine
ON ETHANOL SLICE@
Ethanol oxidation (pm01 h-’ (g of liver)-‘) Aminotriazole
Saline
52.9 k 6.2 23.7 k 2.0 24.1 ? 2.7
49.8 2 5.9 24.5 k 2.1 39.5 c 4.1
P
N.S. N.S.
D Rats were pretreated with 3-amino-1,2,4-triazole at a dose of 1 g/kg body weight (in physiological saline ip) 1 h before sacrifice, whereas the corresponding littermate received saline solution alone. Rat liver slices were incubated for 150 min with ethanol (50 mM) and, when indicated, with pyrazole (2 mM) and nn-alanine (40 mM). Ethanol disappearance was measured by gas-liquid chromatography. The values represent means (*SE) derived from five experiments.
in liver slices of control rats (Table IV). By contrast, no such enhancement by m-alanine was observed in liver slices derived from rats pretreated with 3-amino-1,2,4triazole (Table IV), indicating that the catalase-mediated peroxidation of ethanol was effectively blocked. These results therefore provide strong evidence against a significant role of catalase in the ADHindependent ethanol oxidation. Effect of Butanol
on Ethanol
Metabolism
In view of our previous finding that higher aliphatic alcohols are substrates for the NADPH-dependent microsomal alcohol-oxidizing system but not for catalase
(7), we tested whether butanol might affect ethanol oxidation by the NADPH-dependent microsomal system as well as by liver slices. To that effect, liver microsomes were prepared and incubated with an NADPH-generating system, ethanol, and, when indicated, butanol. Since both acetaldehyde and butyraldehyde are formed during the oxidation of the respective alcohol and, when bound to the semicarbazide, both elicit changes of the absorbance at 224 nm (12), acetaldehyde formation had to be assessed by gas-liquid chromatography rather than by spectrophotometric assay. Upon analysis by gasliquid chromatography the retention time of acetaldehyde was sufficiently different not only from the one of butyraldehyde but also of ethanol and butanol employed as substrates in the assay system. In the presence of butanol (10 mM), the activity of the microsomal ethanol-oxidizing system was significantly inhibited by 46.9% (P < O.Ol), whereas this alcohol had no significant effect on the catalase-mediated ethanol oxidation (Table V). Similarly, the pyrazole-insensitive ethanol oxidation in liver slices was decreased by 43.2% (P < 0.01) in the presence of 10 mM butanol (Table VI), suggesting that the microsoma1 alcohol-oxidizing system is involved in the pyrazole-insensitive ethanol metabolism in liver slices. To evaluate whether the activity of MEOS could account quantitatively for the ADH-independent pathway, MEOS activity was compared with
640
TESCHKE,
HASUMURA
AND
TABLE
LIEBER
V
EFFECT OF BUTANOL ON THE ACTIVITY OF THE NADPH-DEPENDENT MICROSOMAL ETHANOL-OXIDIZING SYSTEM AND ON THE PEROXIDATIC ACTIVITY OF CATALAS@ Substrate Microsomal ethanol oxidizing system Catalase Activity
Ethanol Ethanol
+ butanol
(rim01 of acetaldehyde min-’ (mg of protein)-’
(% of controll
8.1 k 0.9 4.3 2 0.5
100 53.1
Peroxidatic P
activity
(nmol of acetaldehyde min-’ (mg of protein)-‘)
(% of controll
10.8 2 0.4 11.2 2 0.3
100 103.7
co.01
P
N.S.
a The activity of the microsomal ethanol oxidizing system (MEOS) was measured by incubating washed microsomes (3 mg of protein/flask) with ethanol (50 mM), sodium azide (1 mM) and the NADPH-generating system in a medium containing 1 mM Na,-EDTA, 5 mM MgCl,, and 0.1 M phosphate buffer (pH 7.4). When indicated, n-butanol was added at a final concentration of 10 mM. After an overnight diffusion period, acetaldehyde bound to the semicarbazide of the center well was assessed quantitatively by gas-liquid chromatography. Peroxidatic activity of catalase was determined in washed microsomes under similar assay conditions, except that sodium azide was omitted from the incubation medium and the NADPH-generating system was replaced by the H,Oz-producing one. The value of each enzyme activity represents the mean (&SE) of five experiments.
the rates of the pyrazole-insensitive ethanol oxidation by liver slices. Ethanol is oxidized by MEOS in vitro at a rate of 810 nmol min-’ (mg of microsomal protein)-’ (Table III and V), corresponding to 480-600 nmol h-’ (mg of protein)-*. When corrected for microsomal losses during the subcellular preparation, 1 g of liver contains approximately 40 mg of microsomal protein (21); consequently, about 24 pmol of ethanol h-’ (g of liver)-’ could be oxidized by MEOS in vitro, a value which is similar to the rates of the pyrazole-insensitive ethanol oxidation in liver slices of 2229 pmol h-l (g of liver)-’ (Tables II, IV, VI). DISCUSSION
The present study shows that in the presence of 2 mM pyrazole which virtually abolishes alcohol dehydrogenase activity in the hepatic cytosol (Table I) ethanol metabolism in rat liver slices was reduced by only 50-60%, using ethanol at a final concentration of 50 mM (Tables II, IV, and VI); the residual pyrazole-insensitive pathway remained unaffected by both in vitro addition of the catalase inhibitor sodium azide (Table II) or in vim administration of the catalase inhibitor 3-amino1,2,4-triazole (Table IV) but was significantly decreased by butanol (Table VI), an inhibitor of the activity of the microsomal
TABLE VI EFFECT OF n-BUTANOL ON ETHANOL METABOLISM BY LIVER SLICES Incubation medium
Control Pyrazole Pyrazole
+ butanol
Ethanol oxidation (pm01 h-’ (g of liver)-‘) 54.1 t 5.5 29.4 -c 3.6 18.1 2 1.9
P
-co.01
n Rat liver slices were incubated for 150 min with ethanol (50 mM) and, when indicated, with pyrazole (2 mM) and n-butanol (10 mru). Ethanol disappearance was assessed by gas-liquid chromatography. The data are derived from four experiments and are expressed as means (GE).
ethanol-oxidizing system (Table V). It is concluded that, whereas alcohol dehydrogenase is responsible for half or more of ethanol metabolism, the microsomal ethanol-oxidizing system rather than catalase may account for most if not all of the residual alcohol dehydrogenase-independent pathway. The demonstration that ethanol metabolism is partially inhibited by pyrazole (Tables II, IV, and VI) is in agreement with other studies (l-6) and confirms the concept of a significant role of alcohol dehydrogenase in ethanol metabolism. The residual pyrazole-insensitive pathway was ascribed to an enzymic process independent of the activity of alcohol dehydrogenase rather than to incomplete inhibition of
HEPATIC
ETHANOL
alcohol dehydrogenase activity (1, 3, 5, 22, 23). Indeed, in the presence of pyrazole, glucose labeling from [ 1R-3Hlethanol was nearly abolished, while H3H0 production was inhibited less than 50%. In view of the stereospecificity of ADH for [lR3Hlethanol, these findings suggest “the presence of a significant pathway not mediated by cytosolic ADH” (22). Additional evidence that this residual pyrazole-insensitive ethanol metabolism is not ADH mediated was derived from the fact that the cytosolic redox state was unaffected when ethanol was administered together with pyrazole compared to saline pretreated rats (23). Furthermore, compared to ethanol concentrations of 10 mM at which ADH is fully saturated the rates of alcohol metabolism are significantly higher at ethanol concentrations above 25 mM (24). Moreover, under experimental conditions comparable to those of this study (Tables II, IV, and VI), the K, of the pyrazoleinsensitive ethanol oxidation in isolated hepatocytes has been determined to be 13 mM for ethanol (24). This value is one order of magnitude higher than the corresponding value for alcohol dehydrogenase (25, 26). In addition, the pyrazole-insensitive ethanol metabolism was significantly greater at 25 mM ethanol than at 10 mM (24). The inhibition was also not significantly enhanced when the pyrazole concentration was increased from 2 to 4 mM (Matsuzaki and Lieber, unpublished observation). All these findings argue against the possibility that the pyrazoleinsensitive ethanol oxidation observed in the present study is due to incomplete inhibition of alcohol dehydrogenase. Of particular interest was the finding in the present study that butanol inhibits both the pyrazole-insensitive ethanol metabolism by liver slices (Table VI) as well as the activity of the NADPH-dependent microsomal ethanol-oxidizing system (Table V). The latter finding is in keeping with the earlier observation that butanol is a good substrate for the microsomal system (12). These observations strongly suggest the involvement of the microsomal ethanol-oxidizing system in the residual pyrazole-insensitive ethanol metabolism. Consistent with this hypothesis is the ob-
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641
servation that the pyrazole-insensitive ethanol metabolism in liver slices remained unaffected by in vitro addition of sodium azide (Table II) or in viva administration of 3-amino-1,2,4-triazole (Table IV) since the activity of the microsomal ethanol-oxidizing system was found to be insensitive to sodium azide (11) and to 3amino-1,2,4-triazole (Table III). Similarly, the striking increase in the non-ADH fraction of ethanol metabolism with increasing ethanol concentrations (1, 4) is consistent with the known K, of ADH and MEOS: whereas the former has a K, varying from 0.5 to 2 mM for ethanol (25, 261, the latter has a value of 8-9 mM (2). The in vitro K, of MEOS agrees well with the corresponding in uivo value of 8.8 mM for ethanol for the pyrazole-insensitive pathway (3). Furthermore, phenobarbital pretreatment as well as chronic ethanol consumption increased total MEOS activity in the liver, and this was accompanied by an acceleration of blood ethanol clearance, suggesting that the latter might be due, at least in part, to the increased activity of the microsomal system (3). Strong evidence for a microsomal component in ethanol metabolism has been provided by other studies (1, 4-6, 13, 23). In particular, it has been pointed out that the rate of ethanol utilization in perfused livers was much less in the fasted than in the fed state (13). This finding was considered to be consistent with a lack of NADPH generation due to depletion of glycogen in the fasted state, resulting in an insufficient supply of NADPH (13) the cofactor for the activity of the microsomal ethanol-oxidizing system (2). This discrepancy in metabolic rates of ethanol oxidation between the fasted and the fed state persisted even in the presence of mitochondrial inhibitors, supporting the view that an extramitochondrial mechanism was involved (13). Similar results were obtained with aminopyrine, another substrate for the NADPH-dependent microsomal drug-detoxifying enzyme system (13). It is also noteworthy that the rate of ethanol oxidation is strikingly enhanced in patients with glycogenosis type I (27). This finding was explained on the basis that in this particular disorder the content of hepatic glucose 6-phosphate is in-
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creased, which results in an enhanced rate of generation of NADPH (271, the cofactor of the microsomal ethanol-oxidizing system (2). Thus, the present study supports the thesis that the microsomal ethanoloxidizing system may play a significant role in ethanol metabolism in uivo (3) as well as in vitro in perfused liver (6, 131, isolated parenchymal liver cells (1, 51, and liver slices (1, 2, 4). The data presented in this study do rule out a significant role of catalase in ethanol metabolism, in agreement with other reports (5,6,28-32). This is supported by the observation that the pyrazole-insensitive ethanol metabolism in liver slices remained unaffected by in vitro addition of 1 mM sodium azide (Table II). At this concentration, this inhibitor abolished both the catalatic activity of catalase in liver homogenate (Table I) and the catalase-mediated peroxidation of ethanol in liver slices in the presence of 40 mM nn-alanine (Table II), a compound which stimulates peroxysomal H,O, generation via n-amino acid oxidase (33). Furthermore, in uiuo administration of 3-amino-1,2,4-triazole abolished the m-alanine-mediated peroxidation of ethanol via catalase in liver slices (Table IV) and inhibited the catalatic activity of catalase in liver homogenate by 96% (Table III), but this treatment failed to inhibit both the overall rate of ethanol oxidation in liver slices and the pyrazole-insensitive pathway (Table IV). Finally, the observation in the present study that butanol inhibits the pyrazoleinsensitive ethanol oxidation in liver slices (Table VI) without affecting ethanol peroxidation via catalase-H,O, (Table V) indicates that catalase cannot account for the pyrazole-insensitive ethanol metabolism. Supportive evidence against a significant role of catalase in ethanol metabolism was provided by others who reported an extremely low capacity of the liver to generate H,O, (341, which limits the rate of ethanol oxidation via catalase. From these as well as our studies we conclude that catalase plays little if any role in ethanol metabolism and that in addition to alcohol dehydrogenase the microsomal ethanol-
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oxidizing system accounts for a significant fraction of the overall ethanol metabolism. ACKNOWLEDGMENTS The authors are grateful to Miss Nancy Lowe for her excellent technical assistance. REFERENCES 1. GRUNNET, N., QUISTORFF, B., AND THIEDEN, H. I. D. (1973) Eur. J. Biochem. 40, 275-282. 2. LIEBER, C. S., AND DECARLI, L. M. (1970) J. Biol. Chem. 245, 25052512. 3. LIEBER, C. S., AND DECARLI, L. M. (1972) J. Pharmacol. Exp. Ther. 181, 279-287. 4. THIEDEN, H. I. D. (1971) Acta Chem. Sand. 25, 3421-3427. 5. ROGNSTAD, R. (1974) Arch. B&hem. Biophys. 163, 544-551. 6. PAPENBERG, J., VON WARTBURG, J. P., AND AEBI, H. (1970) Enzymol. Biol. Clin. 11, 237250. 7. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S. (1974) Biochem. Biophys. Res. Commun. 60, 851-857. 8. HILDEBRANDT, A. G., SPECK, M., AND ROOTS, I. (1974) Biochem. Biophys. Res. Commun. 60, 851-857. 9. TESCHKE, R., HASUMURA, Y., JOLY, J.-G., ISHII, H., AND LIEBER, C. S. (1972) Biochem. Biophys. Res. Commun. 49, 1187-1193. 10. MEZEY, E., POTTER, J. J., AND REED, W. D. (1973) J. Biol. Chem. 248, 1183-1187. 11. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S. (1974) Arch. Biochem. Biophys. 163, 404-415. 12. TESCHKE, R., HASUMURA, Y., AND LIEBER, C. S (1975) J. Biol. Chem. 250, 7397-7404. 13. SCHOLZ, R., HANSEN, W., AND THURMAN, R. G. (1971) in Metabolic Changes Induced by Alcohol (Martini, G. A., and Bode, Ch., eds.), pp. 101-107, Springer-Verlag, Berlin. 14. THURMAN, R. G., AND MCKENNA, W. (1974) Hoppe-Seyler’s Z. Physiol. Chem. 355, 336340. 15. DE LUCA, H. F., AND COHEN, P. P. (1964) in Manometric Techniques (Umbreit, W. W., Burris, R. H., and Stauffer, J. F., eds.), pp. 131-133, Burgess, Minneapolis. 16. KORSTEN, M. A., MATSUZAKI, S., FEINMAN, L., AND LIEBER, C. S. (1975) N. Engl. J. Med. 292, 386-389. 17. BONNICHSEN, R. K., AND BRINK, N. G. (1955) in Methods in Enzymology (Colowick, S. P., and Kaplan, N. O., eds.), Vol. 1, pp. 495-500, Academic Press, New York. 18. LOCK, H. (1963) in Methods of Enzymatic Analysis (Bergmeyer, H. U., ed.). pp. 885-894, Academic Press, New York.
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19. LOWRY, 0. H., ROSEBROUGH, N. J., FARR, A., AND RANDALL, R. J. (1951) J. Biol. Chem. 193, 265-275. 20. ARSLANIAN, M. J., PASCOE, E., AND REINHOLD, J. G. (1971) &o&em. J. 125, 1039-1047. 21. JOLY, J.-G., FEINMAN, L., ISHII, H., AND LIEBER, C. S. (1973) J. L@pid Res. 14, 337-343. 22. ROGNSTAD, R., AND CLARK, D. G. (1974) Eur. J. Biochem. 42, 51-60. 23. GRUNNET, N., AND THIEDEN, H. I. D. (1972) Life sci. 11, 983-993. 24. MATSUZAKI, S., AND LIEBER, C. S. (1975) Gustroenterology 69, 845. 25. MAKAR, A. B., AND MANNERING, G. J. (1970) Biochem. Pharmucol. 19, 2017-2022. 26. REYXIER, M. (1969)Actu Chem. Stand. 23,11191129.
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