In vitro 31P-NMR spectroscopic studies of rat liver subjected to chronic ethanol administration

151

Biochimica et Biophysica Acta, 1051 (1990) 151-158 Elsevier BBAMCR 12622

In vitro 31p-NMR spectroscopic studies of rat liver subjected to chronic ethanol administration Mingfu Ling and Manfred Brauer Guelph-Waterloo Centrefor Graduate Work in Chemistry, Department of Chemistry and Biochemistry, University of Guelph, Guelph (Canada) (Received 25 September 1989)

Key words: NMR, 31p.; Chronic ethanol; Metabolism; Energy charge; Anoxia

31p_NMR spectroscopy of rat liver perchloric acid extracts was utilized to assess hepatic levels of major phosphorylated metabolites with and without ethanol administration from 0 to 51 days. The results are: (1) 3-phosphoglycerate was largely decreased starting from the first day of ethanol ingestion, (2) in ethanol-treated rats, phosphoethanolamine exhibited a significant decrease relative to the control rats until around day 30, (3) glycerophosphocholine and glycerophosphoethanolamine decreased over the entire period of ethanol ingestion, (4) ATP, AMP, Pi and energy charge showed significant changes due to ethanol effects when a 40-50 s exposure to CO 2 was used prior to freeze-clamping, but ADP did not change, and (5) the other observable metabolites did not show statistically significant change. Some time-dependent changes in metabolite concentration, although not dramatic, were observed from day 1 to day 51 of ethanol administration. Our results indicate that chronic ethanol ingestion results in an inhibition of glycolysis and gluconeogenesis, a decrease in phospholipid breakdown and a lowering of hepatic energy charge.

Introduction Ethanol induces a wide range of metabolic changes in the liver. These changes can eventually become manifest as fatty infiltration, alcoholic hepatitis, fibrosis and cirrhosis [1,2]. One direct result of ethanol metabolism is a redox shift to a higher [ N A D H ] / [ N A D ÷] ratio, which in turn, affects all the major pathways [1]. Other effects, which may or may not be consequences of the redox shift have been reported, including: (i) lowered energy charge and phosphorylation potential [3], (ii) stimulated Na ÷- and K+-dependent ATPase activity [3], (iii) fatty infiltration of the liver as the consequence of inhibited catabolism a n d / o r promoted synthesis of lipids [4], and (iv) inhibition of carbohydrate metabolism, mainly glycolysis and gluconeogenesis [5]. The acute and chronic effects of ethanol

Abbreviations: PCA, perchloric acid; MEOS, microsomal ethanol oxidizing system; FFA, free fatty acid. See Table I for abbreviations of phosphorylated metabolites. Correspondence: M. Brauer, Department of Chemistry and Biochemistry, University of Guelph, Guelph, Ontario, Canada N1G 2Wl.

metabolism on animals are different. For example, ATP levels in the liver were reported to be decreased from chronic effects [6,7], but were lowered [8], did not change [7], or transiently increased [6] after acute administration of ethanol. Adaptive changes in metabolism occur as a function of the length of chronic ethanol exposure. There is an increase in the rate of oxygen consumption [9] and induction of the microsomal ethanol oxidizing system (MEOS) [10] as the liver metabolizes ethanol more rapidly. Furthermore, pathological scores were increased with time of ethanol consumption by the rats [1]. Thus, ethanol has complex effects on the metabolism of the liver, and these effects change as a function of the length of ethanol exposure to the rat. The aim of the present study was to monitor ethanol-induced changes in the levels of the major phosphorus-containing liver metabolites by in vitro 31P-NMR spectroscopy. Rats were pair-fed a nutritionally adequate all-liquid diet containing either ethanol or isocaloric maltose-dextrin [11]. Perchloric acid (PCA) extracts of the rat livers were analyzed by 31p-NMR spectroscopy. 13 phosphorylated metabolites were readily observed. Chronological changes in these metabolites were monitored as a function of the length of control or ethanol feeding.

152 Materials and Methods

Ethanol administration and liver sample pretreatment Wistar male rats weighing 200-250 g were grouped into pairs consisting of one ethanol-treated rat and one control rat of similar weight, and fed a nutritionally adequate liquid diet modified from Ref. 11. The diet for ethanol-treated rats contained 36% (in calories) ethanol, while the one for control rats had isocaloric dextrinmaltose substitution for ethanol. The rats were pair-fed to ensure that the control and ethanol-treated rats received the same number of calories. All ingredients in the liquid diets were purchased from Bioserve, New Jersey, except that carrageenan (type I), corn oil and olive oil were from Sigma. Growth curves for rats transferred from lab chow directly to 36% ethanol (and their pair-fed controls) were compared with curves for rats introduced gradually to 36% ethanol (9% ethanol for 3 days, then 18% ethanol for 3 days, then 27% ethanol for 3 days). The body weights of the two groups were within 10% throughout the 51 days time-course experiment. After a certain period of feeding, the rats were killed in a CO 2 atmosphere for 40-50 s followed by cervical dislocation. Euthanasia methods using ether or pentobarbital anaesthesia or simply rapid cervical dislocation alone were also used. A liver sample was excised in less than 15 s after the rat was unconscious and frozen immediately with W o l l e n b e r g e r - t y p e aluminum tongs precooled in liquid nitrogen. Experiments showed that the time needed to dissect a portion of the liver (less than 15 s) did not cause a measurable breakdown of labile metabolites such as ATP. Extraction of hepatic metabolites A perchloric acid (PCA) extraction procedure modified from Ref. 12 was used whereby the PCA concentration was 6%. Prior to lyophilization, the neutralized extracts were eluted through a Chelex-100 exchange resin to remove divalent metal ions. Paramagnetic metal ions broaden N M R resonances and worsen spectral resolution [13,14]. The lyophilizate was then dissolved in the solution (buffer A) of 20 m M Hepes buffer, 20 m M E D T A and 0.5 M KC1 in 30% D 2 0 at p H 7.5. The purpose of adding 0.5 M KC1 was to precipitate excess perchlorate ions to ensure that the ion concentration product of KC104 was low enough to prevent potential precipitation in the N M R tube. After being adjusted to p H 7.5, the resulting 2 ml solution of liver PCA extract was clarified with a bench-top centrifuge. A p H of 7.5 was selected to carry on all measurements on the extracts because at this p H spectral resolution was optimal and variation of chemical shifts with minor changes in p H was minimal. NMR spectroscopy and resonance assignment A Bruker W H 400 N M R spectrometer was used for

acquisition of 3~P-NMR spectra at 160.2 MHz in thc Fourier-transform mode. Free induction decays were obtained with an excitation pulse of 45 ° and a delay between acquisitions of 3 s, to minimize T~ effects on signal intensity. Residual T1 effects were corrected by comparing signal intensities to fully relaxed spectra (recycle time of 50 s). Proton scalar coupling interactions were removed using two-level broad-band proton decoupling. All measurements were carried out at room temperature. To assign resonances, known compounds were prepared in buffer A and N M R spectra were acquired using the same set of spectral parameters as for PCA extracts. A coincident chemical shift between a test compound and the known compound, combined with knowledge of physiological levels of the known metabolite and the published chemical shift value, were used to identify a peak in the N M R spectrum. If necessary, a known compound was added to the liver extract to see whether the resonance in question would have an increased intensity to make certain of the assignment.

Quantitive analysis of NMR data 31p-NMR spectra from 19 rat liver PCA extracts were acquired with a 2 M methylenediphosphonic acid capillary concentrically mounted on a vortex plug in a 10 m m N M R tube. Spectral peaks were integrated using the integration program in the W H 400 spectrometer. The areas were then converted into concentrations by comparison to the areas of standard solutions of ATP, AMP, and Pi. Against this external concentration reference, the total concentration of all 31p-NMR observable metabolites was almost constant at 20.5 + 1.3 (S.E.M., n = 19) /*mol/g of wet liver for rats fed with control or ethanol formula liquid diets or a lab chow diet regardless of the length of feeding. Because the insertion of the capillary decreased the spectral resolution, total 31p resonance intensity was used as the working concentration reference for all subsequent experiments. Two-way analysis of variance (ANOVA) was used to determine whether ethanol treatment had a statistically significant effect and whether the metabolite levels showed a significant time dependence. Enzymatic analysis of the PCA extracts were done to assess the redox ratios of lactate to pyruvate [15]. The ratios of lactate to pyruvate were 26.6 + 2.8 and 9.9 + 1.7 (mean + S.E., n = 5) # m o l / g of liver for control and ethanol-treated rats killed using the CO 2 method. Using the pentobarbital method, the lactate to pyruvate ratios were 5 . 8 + 0 . 8 and 3 . 6 + 0 . 3 ( m e a n + S . E . , n = 3 ) # m o l / g of liver for control and ethanol-treated rats. This confirms the anoxia associated with the CO2 method. The effects of chronic ethanol relative to control redox ratios are in agreement with other results [16,17]. Our pentobarbital method was not anoxic, since

153 our lactate/pyruvate ratios agree with those of Faupel et al., extrapolated to zero anoxia [18].

CO2 prior to cervical dislocation and rapid freeze-clamping, dramatic and reproducible spectral changes could be observed between ethanol-treated and control rats, especially for the resonances of ATP, AMP and Pi (Fig. 2A and B). A fixed period of anoxia associated with the CO 2 exposure allowed the effects of ethanol on bioenergetic metabolites to become more apparent. Hence, CO 2 killing was used for all subsequent experiments. The assignments of the easily observable a l p r e s o n a n c e s in Fig. 2 to the phosphorus-containing metabolites are shown in Table I. Most lines in the spectra were identified on a one peak-one assignment basis. However, anomeric isomerism results in two resonances for glucose 6-phosphate (G6P) [14], as confirmed by coincident enhancement of two peaks at 4.41 and 4.37 ppm after adding G6P to the extract. The PCA extracts were adjusted to p H 7.5 for 31p-NMR analysis, because the major 31p resonances were optimally resolved at this pH. The possibility that a single resonance may corre-

Results

Initial 31P-NMR spectra of rat liver extracts were obtained after killing using ether, direct rapid cervical dislocation or pentobarbital anaesthesia (Fig. 1). The spectra of control rats (Fig. 1A, C and E) showed a ratio of ATP to ADP (doublet at - 5 . 7 ppm compared to doublet at - 6 . 0 ppm) of 3 - 4 to 1, consistent with normal literature values [7,8,12]. This indicates that our freeze-clamping and PCA extraction methods did not lead to appreciable ATP hydrolysis. Spectral changes for the bioenergetic metabolites observed for ethanoltreated rats (Fig. 1B, D and F) compared to their pair-fed controls (Fig. 1A, C, and E) were relatively minor. Each killing method has its own inherent effects on other metabolite levels [18]. When animals were killed via 40-50 s exposure to

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Fig. 1. 160.2 MHz 31p-NMR spectra of PCA extracts of the livers of rats fed control (A, C, E) or ethanol-containing (B, D, F) liquid diets. For spectra A and B, 2 min ether anaesthesia preceeded freeze-clamping and PCA extraction. For spectra C and D, rapid cervical dislocation preceeded freeze-clamping and PCA extraction. For spectra E and F, pentobarbital anaesthesia (56 m g / k g body weight, i.p.) was initiated 20 min before freeze-clamping and PCA extraction. The primary chemical shift reference was 85% orthophosphoric acid at 0 ppm. The typical number of scans was 2000. Spectra were taken at 25°C. See Table I and Fig. 2 for assignments.

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Fig. 2. 160.2 MHz 31P-NMR spectra of perchloric acid extracts of the livers of rats fed with liquid diets for 8 days in (A) control formula, (C) ethanol formula. B and D are expanded views of A and C between 4.60 and 2.00 ppm. The primary chemical shift reference was 85% orthophosphoric acid at 0 ppm. Typical number of scans was around 2000. The spectra were acquired at 25°C. See Table I for assignments.

spond to two or more compounds with very close chemical shifts and comparable intensities is slight, because pH titration did not reveal any new previously overlapping resonances (Fig. 3). Peaks from - 1 0 . 9 0 to

- 11.16 ppm were contributions from all forms of nicotinamide adenine dinucleotides (NAD, NADH, NADP and N A D P H ) represented as diphosphodiester 1 (DPDE1). A family of diphosphodiesters (DPDE2) including primarily UDP-glucose presented a cluster of peaks from - 1 1 . 3 6 to - 1 2 . 7 3 ppm. The twelve most abundant hepatic metabolites were determined quantitatively by the N M R techniques described in Materials and Methods. Contents of hepatic metabolites per unit wet weight (1 g) of liver were plotted against the length of liquid diet feeding to rats, for both ethanol-treated and isocaloric control animals (Figs. 4-6). Fig. 4 describes those phosphorylated metabolites involved mainly in carbohydrate metabolism. Fig. 5 includes intermediary metabolites in lipid metabolism. Fig. 6 depicts the levels of phosphorylated substrates involved in bioenergetics as well as the values of energy charge, total adenylate pool and phosphorylation potential over the full treatment period in both groups. In the first day of switching diet from the lab chow to liquid diets, rats in both the ethanol-treated and control groups had a prompt shift in most metabolites, suggesting that a dietary adaptation was occurring in this time period. However, judging from the levelling of the metabolite contents in the control animals after the first day (Figs. 4-6), the acute effects of the ethanoland control liquid diet were over quickly and chronic effects could be studied from day 1 to day 51.

TABLE I Resonance assignment of a 3tp - N M R spectrum of the PCA extract of rat liver from a day 8 control rat Peak number a

Chemical shift (ppm)

Assignment

Abbreviation

1 2b 3b 4 b 5b 6 7 b 8b 9 b 10 b 11 b 12 b 13 b 14 b 15 16 17 b

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glucose 6-phosphate sn-glycerol 3-phosphate 3-phosphoglycerate phosphoethanolamine adenosine monophosphate fructose 6-phosphate P3, 2,3-bisphosphoglycerate phosphocholine P2, 2,3-bisphosphoglycerate inorganic phosphate glycerophosphoethanolamine glycerophosphocholine "/-P of ATP (doublet) /3-P of ADP (doublet) a-P of A D P (doublet) a-P of ATP (doublet) diphosphodiesters mainly N A D ( H ) and N A D P ( H ) diphosphodiesters mainly UDP-glucose etc. B-P of ATP (apparent triplet)

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155 which was very close to the reported value of 0.28/zmol per gram wet liver in Faupel et al. [18]. The glycolytic pathway in which 3PGA is generated involves a NAD ÷dependent oxidation of glyceraldehyde 3-phosphate to 1,3-diphosphoglycerate, which undergoes substrate level phosphorylation to produce ATP and 3PGA. Ethanol metabolism inhibits the oxidation of glyceraldehyde 3phosphate by consuming free NAD ÷ in its own metabolism. Because of their low concentrations relative to the most abundant phosphorylated metabolites, glucose 6phosphate (G6P) and fructose 6-phosphate (F6P) were difficult to analyze quantitatively. However, it is apparent from comparison of peak intensities by looking at the spectra that as a result of ethanol metabolism, G6P was lowered and F6P raised (Fig.2). A crossover plot containing G6P, F6P and 3PGA was presented by Ontko et al. [19], in which F6P and G6P levels in alcohol-treated rat livers had both increased with respect to control levels and 3PGA was decreased. The

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Two-way analysis of variance (ANOVA) was used to determine whether ethanol treatment had a statistically significant effect on the various phosphorus-containing metabolites and whether the metabolite levels changed significantly with time (Table II). Mean metabolite concentrations for ethanol and control rats were obtained by averaging values from day 1 to day 51, but statistical significance was evaluated from ANOVA rather than from a comparison of means. In Table II, we can see that with chronic ethanol administration: (i) 3-phosphoglycerate (3PGA) and phosphoethanolamine (PE) decreased to 29 and 70% of control values, respectively (ii) glycerophosphocholine (GPC) and glycerophosphoethanolamine (GPE) decreased significantly, (iii) AMP and Pi levels increased significantly and (iv) ATP, energy charge, phosphorylation potential and total adenylate pool level decreased significantly. Changes of most of these metabolites were not time-dependent ( P < 0.05), but ATP and energy charge did show a statistically significant decrease with time of ethanol feeding.

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157 TABLE II Effects of chronic ethanol ingestion on the levels of phosphorylated metabolites Level a ( # m o l / g of wet liver) Control

Ethanol

3PGA G3P DPDE1 DPDE2 Phosphocholine PE GPC GPE ATP(y-P) ADP(fl-P) AMP Pi energy charge b total adenylate pool phosphorylation potential c

0.24 + 0.02 0.77 + 0.05 3.09+0.25 0.95 + 0.07 1.06-t-0.07 0.56+0.05 1.16 + 0.09 0.85 +0.09 1.20+0.07 1.08+0.06 0.64+0.04 3.79+0.23 0.57+0.03 2.92 + 0.14 0.28+0.03

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n.s., not significant.

from 3PGA (see Fig. 7). 3PGA is converted by the combined action of a phosphatase, a dehydrogenase and transaminase to serine [22]. Serine biosynthesis has been shown to be decreased by ethanol metabolism [23]. Serine is then converted to ethanolamine and phosphorylated to PE, which is used in the biosynthesis of the major phospholipid phosphatidylethanolamine [24]. PE can also be produced from the breakdown of phosphatidylethanolamine by phospholipase C to a minor degree [25]. The breakdown of phosphatidylethanolamine via GPE can be another source of ethanolamine (see Fig. 7). Our results show an ethanol-induced decrease in GPE. Thus the decrease in hepatic PE concentration may be coupled to a decrease in 3PGA a n d / o r GPE. While chronic ethanol administration resulted in lower hepatic levels of PE, phosphocholine, the precursor for another major phospholipid, phosphatidylcholine, did not change. No change in phosphocholine was seen because choline is required from the diet rather than synthesized endogenously. (PhosphatidylPhosphatidylserine (membrane)

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Fig. 7. M a j o r hepatic pathways i n v o l ~ n g PE and GPE.

ethanolamine and phosphatidylcholine themselves were not detected in our present study because they are not extracted by the PCA). From Table II, the level of several metabolites involved in bioenergetics changed significantly with ethanol ingestion: (i) ATP decreased, (ii) AMP increased, (iii) Pi increased and (iv) ADP did not change. From the concentrations of cellular Pi, AMP, ADP, ATP, values of energy charge ([ATP] + [ A D P ] / 2 ) / ([AMP] + [ADP] + [ATP]) and phosphorylation potential [ATP]/[ADP][Pi] also had decreases of very obvious statistical significance. The total adenylate pool showed a slight increase. Most of these changes are consistent with results from other researchers. Helzberg et al. [7], Bernstein et al. [3], Miyamoto and French [26] and G o r d o n [16] have all reported decreased ATP levels, increased levels of Pi and variable results for AMP and ADP levels after chronic ethanol ingestion. These studies all showed a decrease in energy charge a n d / o r phosphorylation potential as a result of chronic ethanol treatment. The magnitude of the decrease varied from study to study due to differences in diet, ethanol concentration and degree of hypoxia associated with killing of the animal and freezing of the tissue. Faupel et al. [18] have shown that the liver bioenergetic metabolites are extremely sensitive to the length of hypoxia. Our results show that ethanol induced only minor changes in the bioenergetic metabolite levels under conditions which minimized liver hypoxia (Fig. 1). Ethanol-induced bioenergetic changes were dramatic only in conjunction with some hypoxia. Thus, at least part of ethanol's hepatotoxic action must involve its sensitization of the liver to hypoxic stress.

158 The mechanism for this sensitization to hypoxia is likely to have at least two stages. From our study, phosphoenergetic metabolites were significantly altered after just one day of ethanol treatment. Ethanol, being a pharmacologically active xenobiotic, must be preferentially metabolized and detoxified by the liver. This metabolism catalyzed by alcohol dehydrogenase and acetaldehyde dehydrogenase generates N A D H which is reoxidized via oxidative phosphorylation, consuming oxygen. The immediate increase in oxygen consumption by the liver after ethanol administration is well documented [1,2,6]. Uncoupling of oxidative phosphorylation and increased consumption of ATP via a 'hypermetabolic state' [1,3] have been proposed as methods whereby excessively high levels of ATP are prevented. An increased rate of 02 consumption due to metabolism of ethanol would make the ethanol-treated liver more susceptible to hypoxic stress. Our study also showed a statistically significant time-dependent change in bioenergetic status over the course of the 51-day study. This may involve the induction of the MEOS system, which requires oxygen directly to metabolize ethanol. French has observed a maximal oxygen consumption by the rat after 18 to 21 days of chronic ethanol treatment [6]. MEOS system induction leads to faster metabolism of ethanol, generating higher levels of acetaldehyde which has toxic effects on mitochondrial function [1,2,19]. Finally, fatty infiltration of the liver occurs over a period of few weeks [11]. The accumulation of hepatic triacylglycerols, phospholipids and protein has been studied by a variety of classical [2,19,28] and contemporary methods, including magnetic resonance imaging [29]. This gradual build-up may constitute a physical barrier to blood flow, again exacerbating hypoxic tissue damage. Acute ethanol effects happen immediately after ethanol reaches animal tissues and organs. For example, Cook et al. found that in only 15 rain after intraperitoneal injection of ethanol, N A D H levels in the liver increased [28], while the rise in blood ethanol levels needed at least 5 min. Our preliminary results also showed that a single intraperitoneal dose of ethanol lowered ATP levels and greatly suppressed the level of 3-phosphoglycerate in 10 min (results not shown). The characteristic of quick response of the metabolites to the presence of ethanol compared to relatively long-term ethanol-induced morphological hepatic changes indicates that a shift in [NADH]/[NAD+], together with other metabolic alterations induced by the redox potential shift are the driving forces of later mitochondrial dysfunction [26] and other biochemical and morphological changes leading to irreversible liver damage.

Acknowledgements Financial support from the Research Board, University of Guelph and from the Natural Sciences and

Engineering Research Council of Canada are gratefully acknowledged. The authors would like to thank technical support from Rita Barbieri and Bill Klimstra.

References 1 Lieber, C.S. (1985) Acta Med. Scand. (suppl.) 703, 11-55. 2 Weiner, F.R., Czaja, M.J. and Zern, A. (1988) in The Liver: Biology and Pathobiology, Second Edn. (Arios, I.M., Jakoby, W.B., Popper, H., Schachter, D. and Shafriz, D.A., eds.), pp. 1169-1193, Raven Press, New York. 3 Bernstein, J., Videla, L. and Israel, Y. (1973) Biochem. J. 134, 515-521. 4 Nikkla, E.A. and Ojala, K. (1963) Proc. Soc. Exp. Biol. Med. 113, 814-817. 5 Krebs, H.A. Freedland, R.A., Hems, R. and Stubbs, M. (1969) Biochem. J. 112, 117-124. 6 French, S.W. (1966) Proc. Soc. Exp. Biol. Med. 121,681-685. 7 Helzberg, J.M., Brown, M.S., Smith, D.J., Gore, J.C. and Gordon, E.R. (1987) Hepatology 7, 83-88. 8 Desmoulin, F., Canioni, P., Crotte, C., G6rolami A. and Cozzone, P.J. (1987) Hepatology 7, 315-323. 9 Yuki, T. and Thurman, R.G. (1980) Biochem. J. 186, 119-126. 10 Alderman, J., Takaki, T. and Lieber, C.S. (1987) J. Biol. Chem. 262, 7497-7503. 11 Lieber, C.S. and DeCarli, L.M. (1970) J. Clin. Nutr. 23, 474-478. 12 Barany, M. and Glonek, T. (1982) Methods Enzymol. 85, 624-676. 13 Glonek, T. and Marotta, S.F. (1987) Horm. Metab. Res. 10, 420-424. 14 Navon, G. Ogawa, S., Shulman, R.G. and Yamane, T. (1977) Proc. Natl. Acad. Sci. USA 74, 87-91. 15 Lamprecht, W. and Heinz, F. (1984) In Methods of Enzymatic Analysis (3rd edn.) Vol. VI (Bergmeyer, H.U., Bergmeyer, J. and Grassl, M., eds.), Verlag-Chemie, Weiheim. 16 Gordon, E.R. (1973) J. Biol. Chem. 248, 8271-8280. 17 Ammon, H.P.T. and Estler, C.J. (1967) Nature 14, 158-159. 18 Faupel, R.P., Seeitz, H.J. and Tarnowski, W. (1977) Arch. Biochem. Biophys. 148, 509-522. 19 Ontko, J.A., Cook, G.A. and Sullivan, A.C. (1973) in Alcohol and Aldehyde Metabolizing Systems, VIII (Thurman, R.G., Williamson, J.R., Fron, H. and Chance, B., eds.) pp. 137-146, Academic Press, New York. 20 Dawson, R.M.C. (1955) Biochem. J. 59, 5-8. 21 Spanneer, S. and Ansell, G.B. (1982) Biochem. J. 208, 845-850. 22 Devlin, M. (1986) Textbook of Biochemistry, p. 459, John Wiley & Sons, New York. 23 LaBaume, L.B., Merrill, D.K and Guynn, R.W. (1987) Arch. Biochem. Biophys. 250, 569-77. 24 Devlin, T.M. (1986) Textbook of Biochemistry, p. 398, John Wiley & Sons, New York. 25 Evanochko, W.T., Sakai, T.T., Ng, T.C., Krishna, N.R., Hyun, D.K., Zeidler, R.B., Ghanta, V.K., Brockman, R.W., Schiller. L.W., Braunschweiger, P.G. and Glickson, J.D. (1984) Biochim. Biophys. Acta 805, 104-116. 26 Miyamoto, K. and French, S.W. (1988) Hepatology 8, 53-60. 27 Cohen, S.M. (1983) J. Biol. Chem. 258, 14294-14308. 28 Cook, G.A., King, M.T. and Veech, R.L. (1980) in Alcohol and Aldehyde Metabolizing Systems IV. (Thurman, R.G., ed.) pp. 433-440, Plenum Press, New York. 29 M. Brauer and M. Ling (1989) Can. Fed. Biol. Soc. 32, Abstr. No. 268.