BIOCHEMICAL MEDICINE 28, 210-223 (1982)

Depressed Oxidative Metabolism in Skeletal Muscle after Chronic Alcohol Consumption

SPENCER SHAW, MARIA ANNA LEO,’ ELIZABETH JAYATILLEKE, AND

CHARLES S. LIEBER~

Alcohol Research and Treatment Center, VA Medical Center, Bronx, New York 10468,

and the Mount Sinai School of Medicine (CUNY), New York, New York 10029

Received February 4. 1982

The effects of chronic alcohol consumption on hepatic mitochondria are well defined (l-3) but the effects on skeletal muscle mitochondria are relatively unknown despite their potentially large contribution to overall intermediate metabolism because the muscle mass constitutes a large percentage of body weight. Mitochondrial inclusions as well as a decrease in the volume fraction of mitochondria have been noted in muscle biopsies obtained from alcoholics under uncontrolled dietary con- ditions (4, 5). Acutely, acetaldehyde depresses respiration in skeletal muscle mitochondria (6). In studies in which alcohol was given chron- ically along with an adequate diet to humans, an increase in size and distortion of mitochondrial shape was observed (7, 8). Chronic ethanol abuse in man results in decreased triosphophate dehydrogenase and lactic dehydrogenase in skeletal muscle (5) and chronic alcohol feeding in rats results in depressed oxidation of ketones (9). The effects of chronic alcohol consumption on more general oxidative metabolism is unknown.

The syndrome of alcoholic myopathy has been well defined (10) but is relatively uncommon. The metabolic alterations in muscle due to al- cohol that precede myopathy are unknown. In order to investigate such changes we studied the metabolic effects of alcohol on skeletal muscle, especially with respect to oxidative metabolism. We studied the mor- phological and biochemical effects of chronic alcohol consumption on skeletal muscle mitochondria in rats and baboons fed alcohol along with adequate diets.

’ Recipient of a Clinical Investigator Award (AM-00698). * To whom correspondence should be addressed at the Alcohol Research and Treatment

Center, VA Medical Center, 130 West Kingsbridge Road, Bronx, NY 10468.

210

0006-2944/82/050210-14$02.00/O Copyright (B 1982 by Academic Press. Inc. All rights of reproduction in any form reserved.

MUSCLE RESPIRATION AND ETHANOL 211

METHODS AND MATERIALS

Animals

Male Sprague-Dawley rats were obtained from the Charles River Breeding Laboratories (Wilmington, Mass.). Littermates were pair fed a nutritionally adequate liquid diet containing either ethanol as 36% of total calories or with ethanol substituted isocalorically by carbohydrate as previously described (11) for a period of 8 to 10 weeks.

Baboons were housed at the Laboratory for Experimental Medicine and Surgery in Primates (LEMSIP), Tuxedo, New York. Animals were pair fed nutritionally adequate liquid diets containing either alcohol at 50% of total calories or with carbohydrate substituted isocalorically for ethanol as previously described (11) for a period of 1 to 4 years.

Sampling

Rats were sacrificed by decapitation following an overnight fast. Ba- boons were subjected to a muscle biopsy under light ketamine anesthesia following an overnight fast. At the time of sampling both rats and baboons had no measurable ethanol or acetaldehyde in the blood using a gas-liquid chromatography method (sensitivity:ethanol 0.5 mM, acetaldehyde lug) (12).

Morphological Studies

Muscle samples obtained from rat hindquarter muscles and gastroc- nemius muscles in the baboons were fixed in 10% buffered Formalin and stained with hematoxylin and eosin for light-microscopic study. For electron-microscopic studies the muscle fibers were clamped to maintain the isometric conditions and fixed overnight in ice-cold 2.5% glutaral- deyde in 0.1 M cacodylate buffer. Subsequently the tissues were trimmed into small cubes and fixed for 3 more hr in fresh glutaraldehyde (pH 7.4) and postfixed in 1% osmium tetroxide (13). After dehydration and infil- tration the tissues were embedded in Epon 812 (14). Following uranyl acetate and lead citrate staining (15, 16), ultrathin longitudinally cut sections were examined under a Hitachi HS-8 electron microscope.

Respirometer Studies

Oxygen respiration was measured with muscle slices from rat hind- quarter muscles. Hindquarter muscles were dissected free of connective tissue and sliced in a Stadie-Riggs microtome in iced saline into pieces of approximately 0.5-mm thickness. Approximately 100-200 mg of tissue was placed in a 25-ml respirometer flask containing 3 ml of phosphate buffer (17). Additional sets of flasks contained added glucose (final con- centration 100 mg/dl) or leucine (60 +T). Incubations were carried out in a Gilson G-30 respirometer at 37°C and oxygen consumption was mea-

212 SHAW ET AL

sured as outlined in the Gilson Respirometer Manual (18) at S-min in- tervals over a period of 45 min and the mean rate was determined. Flasks with boiled muscle tissue and with no added tissue were used as controls.

Mitochondrial Studies

Preparation. Mitochondria from rats and baboons were prepared ac- cording to the method of Sirotzky (19). Hindquarter muscle in the rat and gastrocnemius muscle in the baboon were freed of connective tissue and finely minced in ice-cold KC1 (0.15 M). The mince was then incubated for 15 min in a modified Chapel&Perry medium which included 2 mg bacterial proteinase per gram muscle (Nagarse, Teikoku Chemical In- dustry, Osaka, Japan) (19). The mince was then homogenized with a Potter-Elvehjem homogenizer and filtered through cheese cloth.

Mitochondria were prepared by centrifugation of the filtrate and re- suspension in a modified Chapell-Perry medium to which 1 mg/ml of bovine albumin was added according to the method of Sirotzky et al. (19).

Oxygen consumption. Oxygen consumption was measured polaro- graphically at 27°C using a Clark-type oxygen electrode. The reaction mixture (3 ml) contained mannitol(0.25 M), KC1 (10 mM), K2HP04-Na2HP04 (5 mM), pH (7.4), Tris-HCl (10 mM) (pH 7.4), and EDTA (2 mM). Ap- proximately 3 mg of mitochondrial protein was added to each reaction mixture. Substrates included glutamate (10 mM), glycerol-l-phosphate (10 mM), and malonate (5 mM) + alphaketoglutarate (5 mM).

State 3 respiration (according to the nomenclature of Chance and Williams) (20) was measured by the addition of GDP to the reaction mixture in a final concentration of 150 PM. A second addition of ADP was made after the return of the reaction mixture to State 4 respiration and the respiratory control ratios and ADP:O ratios were calculated according to the method of Estabrook (21).

Recoveries. Mitochondrial recovery was estimated by the measure- ment of glutamate dehydrogenase as previously described (22). Protein concentrations of mitochondria were determined by the method of Lowry et al. (23).

Statistics. All data were expressed as mean t standard error of the mean. Significance of differences was assessed by the paired Student t test.

RESULTS

Morphology

By light microscopy, no abnormalities were found in the muscles of ethanol-fed rats, ethanol-fed baboons, and their respective pair-fed con- trols. By electron microscopy, however, compared to the control animals

MUSCLE RESPIRATION AND ETHANOL 213

which were normal (Fig. l), all alcohol-fed rats showed irregular and swollen mitochondria (Fig. 2) with rarefaction of the matrix, alterations of the cristae, and occasional myelin figures. Focal sarcomere abnor- malities were also present such as fragmentation of the Z bands and disorganization of the myofilaments (Fig. 3). Compared to the control baboons (Fig. 4), in the ethanol-treated animals severe mitochondrial lesions were present, ranging from simple enlargement (Fig. 5) to an “empty” appearance of the matrix, with alterations of membranes and cristae (Fig. 6). Edema and numerous fat droplets were also noted in the intermyofibrillar space. In both species, muscle specimens of alcohol- fed animals were in various stages of contraction, possibly due to a slight ethanol withdrawal state.

Respirometer studies. Oxygen consumption in muscle slices from rat hindquarter muscles was depressed in alcohol-fed animals compared to controls both in the absence and presence of added substrate: no added substrate 455 + 35 vs 595 2 40 ~1 O2 g-l hr-1 n = 6, P < 0.001; glucose (100 mg/dl) 412 + 41 vs 531 +- 51, n = 7, P < 0.01; leucine (60 PM) 471 -C 59 vs 561 t 72, n = 7, P < 0.02. The percent depression was 24.1 + 2.2, 23.7 + 7.3, and 18.1 + 5.7%, respectively.

Mitochondrial Studies

Rats. Mitochondrial recovery estimated by the measurement of glu- tamate dehydrogenase was not significantly different in alcohol-fed an- imals (29.5 +- 3.4%) and controls (37.5 + 4.2%).

Oxygen consumption was depressed approximately 40% in mitochon- dria obtained from skeletal muscle from alcohol-fed animals compared to controls. Differences were similar in State 3 with various substrates including glutamate, glycerol-l-phosphate, and malonate with alpha-ke- toglutarate (Table 1). Similar trends were noted in State 4 respiration but differences were only significant with glutamate as substrate (Table 1). Respiratory control ratios and ADP:O ratios were not significantly different in alcohol-fed animals compared to controls (Table 1).

Baboons. Mitochondrial recovery estimated by the measurement of glutamate dehydrogenase was not significantly different between alcohol- fed animais (16.7 ? 2.3%) and controls (10.0 + 1.9%).

Oxygen consumption was depressed approximately 50% in mitochon- dria of skeletal muscle from alcohol-fed animals compared to controls. Differences were similar in States 3 and 4 with the two substrates tested: glutamate and glycerol-l-phosphate (Table 2). The respiratory control ratios and ADP:O ratios were not significantly different in alcohol-fed animals compared to controls (Table 2).

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MUSCLE RESPIRATION AND ETHANOL 215

216 SHAW ET AL.

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218 SHAW ET AL.

MUSCLE RESPIRATION AND ETHANOL 219

220 SHAW ET AL.

TABLE I EFFECT OF CHRONIC ALCOHOL CONSUMPTION ON RESPIRATION OF SKELETAL MUSCLE

MITOCHONDRIA IN THE RAT

Oxygen uptake (micro atoms 0, hr- I

mg- 1 mit. prot.) Respiratory

Substrate State 3

Glutamate (10 mhi) Ethanol

fed 9.2 r 0.97 (n = 6)

Control 14.2 r 1.97 (n = 6)

P co.01 Glycerol-l-

phosphate (10 mM) Ethanol

fed 2.44 ix 0.31 (n = 6)

Control 3.29 k 0.35 (n = 6)

P co.02 Malonate +

alpha-keto glutarate (5 rnM)

Ethanol fed 2.55 + 0.13 (n = 4)

Controls 3.95 k 0.40 (II = 4)

P co.02 I_~

State 4 control ratio -~

1.25 f 0.15 7.47 k 0.65

1.55 2 0.31 7.02 2 0.94

co.02 NS

1.49 2 0.20

1.61 2 0.30

NS

1.66 r 0.08

2.07 +- 0.22

NS

0.94 2 0.15

1.46 ? 0.23

NS

2.93 t 0.43

2.93 2 0.39

NS -

DISCUSSION

The results of this study demonstrate that chronic alcohol consumption of relatively brief duration is associated with striking ultrastructural changes in skeletal muscle, especially in the mitochondria. These mor- phological alterations were accompanied by depressions in oxidative metabolism in skeletal muscle slices as well as isolated mitochondria. The changes were observed following an overnight fast when alcohol and acetaldehyde were no longer measurable in the blood.

ADP:O ratio -

2.76 f 0.11

2.69 f 0.18

NS

1.6 k 0.14

1.56 2 0.20

NS

3.25 2 0.17

3.20 ” 0.25

NS

The changes we observed occurred when alcohol was given along with an adequate diet, Since dietary deficiencies may alter mitochondrial func-

MUSCLE RESPIRATION AND ETHANOL 221

TABLE 2 EFFECT OF CHRONIC ALCOHOL CONSUMPTION ON SKELETAL MUXLE MIT~CHONDRIA

IN THE BABOONS -___

Oxygen uptake (micro atoms Oz hr- I

mg- 1 mit. prot.) Respiratory control ADP:O

Substrate State 3 State 4 ratio ratio

Glutamate (10 mM) Ethanol fed 2.84 2 0.65 0.55 -c 0.13 5.15 f 0.56 2.46 2 0.25

(n = 4) Controls 5.83 -c 0.94 1.10 k 0.11 4.77 2 0.42 2.88 t 0.36

(n = 4) P co.01 co.02 NS NS

Glycerol- l-phosphate (10 mm) Ethanol fed 0.72 “- 0.14 0.45 2 0.13 1.53 + 0.18 1.79 !z 0.30

(n = 4) Controls 1.59 + 0.44 1.0 k 0.30 1.40 i 0.10 1.53 2 0.24

(n = 4) P co.05 NS NS NS

a Animals were fed ethanol as 50% of total calories for 1 to 4 years.

tions (24-26), the changes we observed may be potentiated in alcoholics by concomitant dietary deficiencies. Furthermore, in our studies animals were withdrawn from alcohol to rule out acute affects of alcohol. Because acetaldehyde, a product of alcohol metabolism, has been shown to de- press respiration in skeletal muscle mitochondria (6), the changes we observed may be increased during acute ethanol consumption. The mor- phological changes reported here were similar to those observed in al- coholic volunteers given alcohol along with adequate diets (7,8). In these studies the morphological abnormalities persisted for 6 weeks beyond cessation of drinking. Thus, the animal changes we observed can be considered as a model of corresponding human abnormalities.

The changes in mitochondrial respiration we observed in muscle dif- fered from those reported in hepatic mitochondria. For instance, with glutamate or alpha-ketoghrtarate as substrates, we did not observe the decrease in the respiratory control ratios or ADP:O ratios found with hepatic mitochondria (2). Thus, in contrast to liver organelles, muscle mitochondria may be damaged at a site other than coupling site 1 of the respiratory chain. The changes we observed in rats after 8 to 12 weeks were similar in magnitude to those observed in baboons given a greater amount of alcohol (as percentage of calories) for 1 to 4 years.

The significance of the observed morphological abnormalities and de- pressed oxidative metabolism remains to be evaluated. Such changes

222 SHAW ET AL

might explain, at least in part, the subclinical myopathy seen in alcholics which includes muscle cramps or weakness (27). Similarly, the relatively large proportion of body mass that is comprised of muscle (40%) and the increasing awareness of the importance of this organ in overall fuel metabolism suggest that the changes we observed may have important implications for overall intermediate metabolism.

In conclusion, chronic alcohol administration along with an adequate diet results in mitochondrial abnormalities and depression of respiration in skeletal muscle. These changes are observed in the absence of alcohol or its metabolites and occur after as short a period of time as 8 weeks of drinking.

SUMMARY

Morphological and biochemical studies were performed on skeletal muscle obtained from rats and baboons fed alcohol chronically along with an adequate diet. Muscle mitochondria from both species demon- strated marked alterations: enlargement and distortion of shape, loss of cristae, and vacuolization of the matrix. Following an overnight fast, when alcohol was no longer measurable in the blood, respiration in muscle slices obtained from the rats was depressed 25-40% (P < 0.01-0.05) both in the presence and absence of added substrate. Isolated mitochondria from rats and baboons demonstrated depressions in oxygen consumption ranging from 45 to 60%. These changes were present with various substrates: glutamate (10 mM), glycerol-l-phosphate (10 mM), and malonate with alpha-ketoglutarate (5 mM + 5mM). Changes were similar in States 3 and 4 of respiration. ADP:O ratios and respiratory control ratios were not significantly different in alcohol-fed and control animals. Thus, chronic alcohol consumption results in morphological changes in muscle mitochondria associated with a marked depression in oxidative metabolism.

ACKNOWLEDGMENTS We gratefully acknowledge the expert assistance which the LEMSIP staff provided for

the baboon studies and Steven A. Mortillo for the technical help with the electron microscopy. The work in this study was supported by the Veterans Administration and USPHS Grants AA-03508 and AA-00698.

REFERENCES 1. Cederbaum, A. I., Lieber, C. S., Beattie, D. S., and Rubin, E., J. Eiol, Chem. 250,

5122 (1975). 2. Cederbaum, A. I., Lieber, C. S., and Rubin, E., Arch. Biochem. Biophys. 165, 560

(1974). 3. Kiessling, K. H., and Tobe, U., Cell Res. 33, 350, (1964). 4. Fisher, E. R., Puntereris, A. J., Jung, Y.. Corredor, D. G.. and Danowski, T. S.,

Amer. J. Med. Sci. 261. 85 (1971).

MUSCLE RESPIRATION AND ETHANOL 223

5. Kiessling, K. H., Pilstrom, I., Byhmd, A. C., Piehl, K., and Saltin, B. Stand. J. Clin. Lab. Invest. 35, 601 (1975).

6. Cederbaum, A. I., and Rubin, E., Biochem. Pharmacol. 26, 1349 (1977). 7. Rubin, E., Katz, A. M., Lieber, C. S., Stein, E. P., and Puszkin, S., Amer. J. Pathol.

83, 499 (1976). 8. Song, S. K., and Rubin, E., Science 175, 327 (1972). 9. Lefevre, A., Adler, H., and Lieber, C. S., J. Clin. Sci. 49, 1775 (1970).

10. Perkoff, G. T., Hardy, P., and Velez-Garcia, E., N. Engl. J. Med. 274, 1277 (1966). 11. Lieber, C. S., and DeCarli, L. M.,Fed. Proc. 35, 1232, 1976. 12. Korsten, M. A., Matsuzaki, S., Feinman, L.. and Lieber, C. S. N. Eng. J. Med. 292,

386 (1975). 13. Sabatini, D. D., Bensch, K., and Barrnett, R. J., J. Cell Biol. 17, 19 (1963). 14. Luft, J. H., J. Biophys. Biochem. Cytol. 9, 409 (1961). 15. Reynolds, E. S., J. Cell Biol. 17, 208 (1963). 16. Stempak, J. G., and Ward, R. T., J. Cell Biol. 22, 697 (1964). 17. Adibi, S. A., Krzysik, A., Morse, E. L.. Amin, P. F., and Allen, E. R., J. Lab. C/in.

Med. 83, 548 (1974). 18. Gilson Differential Respirometer, Directions for Operations, Gilson Medical Electron-

ics, Middleton, Wisconsin, Feb. 1974. 19. Sirotzky, S., DeFavelukes, S., Schwartz de Tarlovsky, M., and Stoppani, A. 0. M.,

Acta Physiol. Lat. Amer. 21, 30 (1971). 20. Chance, B., and Williams, G. R., Adv. Enzymol. Rel. Sub. Biochem. 17, 65 (1956).

21. Estabrook, R. V., in “Methods of Enzymology,” (R. W. Estabrook and M. E. Pullman, Eds.), Vol. 10, p. 41. Academic Press, New York, 1967.

22. Hasumura, Y., Teschke, R., and Lieber, C. S., J. Biol. Chem. 251, 4908 (1976). 23. Lowry, 0. H., Rosebrough, N. J., Fat-r, A. L., and Randall, R. J., J. Biol. Chem.

193, 265 (1951). 24. Burch, H. B., Hunter, F. E., Combs, A. M., and Schutz, B. A., J. Biol. Chem. 235,

1540 (1960). 25. Gold, A. J., and Costello, L. C., J. Nutr. 108, 208 (1975). 26. Houtsmuller, U. M. T., VanDerBeek, A.. and Zaalberg, J., Lipids 4, 571 (1969). 27. Perkoff, G. T., Dioso, M. M., Bleisch, V., and Klinkerfuss, G., Ann. Intern. &fed.

67, 481 (1967).