Anaerobic metabolism during electrical stimulation of aged rat skeletal muscle.

The ability of human skeletal muscle to generate tension decreases with age.[1-4] Aged human: skeletal muscle can increase in tension-generating capacity following high-intensity resistance training.[5,6] The brief, high-intensity muscle contractions associated with resistance training require rapid resynthesis of adenosine triphosphate (ATP), primarily through use of intramuscular high-energy phosphate stores and anaerobic glycolysis.[7] Little is known, however, about the capacity of aging muscle to anaerobically resynthesize ATP during brief, high-intensity contractions.[8,9]

Studies investigating the effects of age on anaerobic metabolism during brief, high-intensity muscle contractions suggest that in rat muscle there is not an age-related decrease in the muscle's capacity for anaerobic metabolism during brief, high-intensity contractions.[8,9] Campbell and colleagues[8] report similar anaerobic resynthesis of ATP in fast-twitch oxidative glycolytic fibers of gastrocnemius and plantaris muscles and slow-twitch oxidative fibers of soleus muscles of 11- and 25-month-old Fisher 334 rats during I minute of in situ, supramaximal, intermittent, tetanic electrical stimulation. During stimulation, however, less anaerobic resynthesis occurs in fast-twitch glycolytic fibers of the gastrocnemius muscle of 25-month-old rats compared with fast-twitch glycolytic fibers of the gastrocnemius muscle of 11-month-old rats.[8] The maximum tension generated by the gastrocnemius-plantaris-soleus (GPS) muscle group also declines with age.[8] The majority of the tension produced by the GPS muscle group is generated by the fast-twitch glycolytic fibers of the gastrocnemius muscle. The authors conclude that the 25-month fast-twitch glycolytic fibers of the gastrocnemius muscle use less anaerobic substrates during the stimulation bout compared with the 11-month fast-twitch glycolytic fibers of the gastrocnemius muscle due to an age-related decline in the muscle's energy demand.[8]

Anaerobic resynthesis of ATP appears to be greater in aged muscle compared with adult muscle when brief, high-intensity muscle contractions occur over a long time period.[9] Fitts and colleagues[9] found greater decreases in creatine phosphate (CP) and glycogen concentrations in the soleus muscle of 28-month-old Long-evans rats compared with the soleus muscle of 9-month-old Long-evans rats during 30 minutes of in situ, supramaximal, intermittent, tetanic electrical stimulation.

The effect of age on anaerobic metabolism has been studied in muscles that show no age-related differences in the percentage of decline in tension during supramaximal, intermittent stimulation.[8,9] Further investigation is warranted about the effect of age on anaerobic metabolism in muscles where an age-related decline in tension during supramaximal, intermittent electrical stimulation has been observed.

When the flexor digitorum brevis (FDB) muscles of 24- and 36-month-old rats were compared with the FDB muscles of 5-month-old rats, -- an earlier And greater decline in peak tetanic tension during 2 minutes of in vitro, supramaximal, tetanic electrical stimulation was found.[10] The FDB muscle is composed of 90% to 95% fast-twitch oxidative glycolytic fibers and 4% to 5% slow-twitch oxidative fibers, and this muscle has a high capacity for anaerobic metabolism.[11] The purpose of my study was to investigate the utilization of high-energy phosphates and glycogen during repeated short-duration (10 seconds on:20 seconds off) bouts of supramaximal, tetanic electrical stimulation in aging FDB muscle.

Method

Animals and Animal Care

Virgin 12- and 32-month-old Fl hybrid Fisher 344 X Norway Brown male rats, obtained through the National Institute on Aging (NIA),(*)[1] were used in this study. These age groups were chosen based on the probability of survival curve for the Fl hybrid male rats.[12] The longevity of ad libitum-fed F1 hybrid male rats is 39 to 40 months.[12] The probability of survival of Fl hybrid male rats is 96% at 12 months of age, and 40% at 32 months of age.[12]

The rats were individually, housed for 2 weeks under pathogen-free conditions in the animal quarters of the University of Illinois at Chicago. The animal room was maintained at a temperature between 21[degrees] and 23[degrees]C, a humidity of 50 [greater than or equal to] 5%, and a light/dark cycle of 12 hours/12 hours. The rats were provided with unrestricted access to feed (18% crude protein, 5% crude fat, and 5% crude fiber) and water. The environmental conditions and diet were similar to those in the NIA colony where the rats were raised.12 Rats that became sick (eg, rapid weight loss following arrival) were excluded from the study. All experiments were begun between 9 and 10 AM to prevent feeding and circadian influences on the intramuscidar substrate concentrations.[13]

Prior to bilateral excision of the FDB muscles from the hind limbs, each rat was anesthetized by an intraperitoneal injection of sodium pentobarbital[dagger] (40 mg per kilogram of body weight). The elapsed time from administration of the anesthetic to the rat to excision of the muscles from the hind limbs was 5 minutes. At the time of muscle excision, the rat was in a deep plane of anesthesia, as indicated by the animal's respiration rate and lack of response to tail pinch.

Experimental Protocol

Twelve FDB muscles from 12-month-old rats and 12 FDB muscles from 32-month-old rats were randomly assigned to one of three experimental groups (8 muscles per experimental group): control (no incubation and no electrical stimulation), incubation (no electrical stimulation), or electrical stimulation (combined with incubation). Bcidy weights of the anesthetized rats were measured prior to the bilateral excisions of the FDB muscles.

Control muscles were frozen within 2 seconds of excision from the rats' hind limbs. Following excision of control muscles, blood was withdrawn from the descending aorta of the live anesthetized rats for measurement of plasma palmitate and glucose concentrations. Plasma palmitate and glucose concentrations in the 12- and 32-month-old rats, from which control muscles had been excised, were measured to determine incubation medium palmitate and glucose contents for the 12- and 32-month-old rats' incubation and electrical stimulation muscles.

Incubation and electrical stimulation muscles were incubated for 15 minutes in Krebs-Henseleit bicarbonate buffer containing glucose,[double dagger] insulin,[sections] palmitate,[parallel] and bovine serum albumin[double dagger] (BSA). During the 15-minute incubation period, the electrical stimulation muscles were stimulated intermittently (10 seconds on:20 seconds off) using direct-current monophasic waveforms at a supramaximal intensity of 80 V, a pulse duration of 200 milliseconds, and a frequency of 5.0 Hz.[14,15] These stimulation settings were chosen because a summation of successive contractions occurs during each 10-second stimulation train when using this setup.[14,15] In addition, during 10 seconds of maximum contractions, the majority of the energy for the muscle contractions comes from anaerobic metabolism.[7] In addition, the rest interval was twice as long as the work interval to ensure sufficient restoration of high-energy phosphates and removal of intramuscular lactate following the 10-second trains and to prevent undue muscle fatigue.[16] Following the 15-minute incubation period, the incubation and electrical stimulation muscles were frozen within 2 seconds of removal from the incubation medium.

Preparation of Incubation Medium

A modification[14,17] of the method of Crass et al[8] was used for preparation of the incubation medium palmitate 3% BSA (98%-99% albumin, essentially fatty acid free) solution. For incubation of the 12- and 32-month-old rats' muscles, the medium palmitate concentrations were 0.50 [greater than or equal to] 0.05 mM and 0.55 [greater than or equal to] 0.05 mM, respectively. Incubation medium- palmitate concentrations were based on the plasma palmitate concentrations in the 12- and 32-month-old rats from which the control muscles were taken (0.51 [greater than or equal to] 0.04 mM and 0. 56 [greater than or equal t] 0.06 mM, respectively). Following preparation, the palmitate 3% BSA solution was dialyzed overnight at 4[degrees]C with Krebs-Henseleit bicarbonate buffer.[14,17] Before use, the solution was warmed to 37[degrees]C and glucose and 100 [mu]U of insulin per milliliter of incubation medium[19] were added. The medium glucose concentration was 5.68 [greater than or equal to] 0.31 mM for incubation of the 12-month-old rats' muscles and 5.95 [greater than or equal to] 0.41 mM for incubation of the 32-month-old rats' muscles. Glucose concentrations were based on the plasma glucose concentrations in the 12-and 32-month-old rats from which the control muscles were taken 5.71[greater than or equal to]O.44 mM and 5.99 [greater than or equal to] 0.36 mM, respectively).

In Vitro FDB Muscle Preparation

The FDB muscles in the incubation and electrical stimulation experimental groups were placed in 21[degrees]C Krebs-Henseleit bicarbonate buffer (pH 7.4) bubbled with 95% oxygen/5% carbon dioxide within 2 seconds of the muscles' removal from the hind limbs.[14,15,17] While in the 21[degrees]C Krebs-Henseleit bicarbonate buffer, adherent adipose tissue was removed from the muscles. The muscles were then positioned, with their long axes vertical, on muscle holders between two platinum sheet electrodes (3x18 mm).[14,15,17] The proximal tendon of the muscle was clamped to the bottom of the muscle holder while a suture was tied to the three distal tendons.[14,15,17] The muscle holder was then transferred to an incubation chamber in an isolated organ tissue bath maintained at 37[degrees]C.[14,15,17] An average of 300[greater than or equal to] 30 seconds elapsed between placement of the muscle in the 21[degrees]C Krebs-Henseleit bicarbonate buffer and placement of the muscle holder, containing the muscle, in the incubation chamber. Each incubation chamber contained 25 mL of incubation medium, which was continuously gassed with a gentle stream of 95% oxygen/5% carbon dioxide (2 ml/min).[14,15,17]

When the muscle holder was positioned in the incubation chamber, the suture attached to the three distal tendons was tied directly to a force-displacement transducer.[14,15,17] The output of the transducer was connected to a low-level direct-current preamplifier of a chart recorder.[14,15,17] The output of the amplifier of the chart recorder was connected to a recorder. Lead wires attached to the platinum sheet electrodes were then connected to an electrical stimulator.[14,15,17] The length-tension relationship for each muscle was determined using a supramaximal, monophasic waveform of 80 V intensity and 200 milliseconds' pulse duration.[14,15,17] The muscle was positioned at the length that produced the maximum force displacement.

Muscle Contractile Measurements

In the incubation muscles, peak isometric twitch tension ([P.sub.1]) was determined at 0 and 15 minutes of incubation using a single supramaximal impulse of 80 V intensity and 200 milliseconds' pulse duration. In the electrical stimulation muscles, peak train tension ([P.sub.tr), or the peak tension during each 10-second stimulation train, was measured.

Biochemicol Procedures

Muscle ATP and CP contents were measured by the procedure of Lamprecht et al.[20] Glycogen content was determined by the method of Lo et al.[21] Lactate concentrations in muscle extracts and incubation medium were determined by the method of Gutmann and Wahlefeld.[22] Plasma and incubation medium glucose contents were determined by the method of Bergmeyer et al.23 Plasma and incubation medium palmitate were extracted according to the procedure of Dole.[24] The concentrations of plasma and incubation medium palmitate were determined by the method of Duncombe.[25]

Expression of Results and Statistical Anolysis

Contractile and biochemical data are expressed per gram wet weight (gww) of muscle. Muscle wet weights were determined prior to biochemical analyses. Mean values ([greater than or equal to] SD) were computed. When 12- and 32-month-old rats' body weights (12 rats per age group) or FDB muscle weights (%4 muscles per age group) were compared, the Student t test was used.[26] Comparisons of 12-and 32-month-old rats' muscle tensions and substrate and metabolite values (8 muscles per experimental group) were made using a two-way analysis of variarice with time as the repeated measure.[26] When significance was found at P<.05, means were compared using Tukey's post hoc test.[26]

Results

Body and Muscle Weights

The body weight of 32-month-old rats (539.88[greater than or equal to]5.84 g) was 18% greater (P<.05) than the body weight of 12-month-old rats (457.22[greater than or equal to] 4.33 g). In contrast, the wet weight of the FDB muscles from the 32-month-old rats 40.20[greater than or equal to]O.55 mg) was 12% less (P<.05) than that of the 12-month-old rats (45.55[greater than or equal to]O.44 mg). In both age groups, there were no differences in muscle wet weights immediately following excision or following 15 minutes of incubation or stimulation.

Muscle Performance

At the beginning of stimulation, the 12- and 32-mondiold rats' muscle [P.sub.t] and [P.sub.tr] values were not different (Tab. 1). During small, there was an earlier (5 minutes versus 8 minutes) and larger (46% versus 40%) decline in the 32-month-old rats' muscle Pt, compared with the 12-month-old rats' muscle [P.sub.tr] (Tab. 1, Figure).

[TABULAR DATA 1 OMITTED]

Intramuscular Phosphote Concentrations

Adenosine triphosphate concentrations were similar in the 12- and 32-month-old rats' muscles in the two control preparations (Tab. 2). In addition, ATP concentrations in the 12- and 32-month-old rats' stimulated muscles were less than the control Values but not different from each other (Tab. 2).

[TABULAR DATA 2 OMITTED]

Creatine phosphate concentrations were also similar in the 12- and 32-month-old rats from which the control muscles were taken (Tab. 2). In addition, CP concentrations in the 12- and 32-month-old rats' stimulated muscles were less than the control values but not different from each other (Tab. 2). During recovery (2 minutes after the end of 15 minutes of stimulation), CP concentrations in the stimulated muscles of five 12-month-old rats and five 32-month-old rats were only 17% 11.86 [greater than or equal to] 0.47 bkmol/gww) and 19% (10.98 [greater than or equal to] 0.45 [mu]mol/ gww) less than the control values.

Ancierobic Glycolysis

Glycogen concentrations were similar in the 12- and 32-month-old rats from which the control muscles were taken (Tab. 2). Glycogen concentrations in the 12- and 32-month-old rats' stimulated muscles were less than the control values and different from each other (Tab. 2). The decrease following stimulation in glylogen content in the 32-month-old rats' muscles was greater than the decrease in the 12-month-old rats' mus.cles (Tab. 2). Glycogen use during stimulation of [mu]bies in the 12-month-old rats (0.97 mg/gww) and the 32-month-old rats (1.65 mg/gxvw) can also be expressed as mic-pomoles per gram wet weight. Using a formula weight for glucose of 180.2 g, 5.39 btmol/gww and 9.17 blmol/gww were used by the 12- and 32-month-old rats' muscles during stimulation.

Lactate concentrations were similar in the 12- and 32-month-old rats' control muscles (Tab. 2). No measurable amounts of lactate could be obtained in the incubation medium from the 12- and 32-month-old rats' incubation muscles (Tab. 2). Following 15 minutes of stimulation, lactate production in the 12- and 32-monthold rats' muscles were greater than in the control muscles and different from each other (Tab. 2). The production of lactate in the 32-month-old rats' stimulated muscles was greater than in 12-month-old rats' stimulated muscles (Tab. 2). Total lactate (muscle 4nd incubation medium) in the muscle of the 12-month-old rats (4.10 [mu]mol/gww) and the 32-month-old rats (9.28 [mu]mol/gww) followed a pattern similar to that of glycogen in the muscles of the 12-month-old rats (5.39 [mu]mol/gww) and the 32-month-old rats (9.17 [mu]mol/ gww)

Discussion

During a series of repeated, short-duration, high-intensity contractions of the FDB muscle, the age-related difference in the decline of muscle tension is not accompanied by an age-related decrease in intracellular anaerobic metabolism. The FDB muscle's capacity for intramuscular utilization of high-energy phosphates is maintained with age. There is also an age-related increase in intramuscular glycogen depletion and in the accumulation of intramuscular and incubation medium lactate. Intramuscular glycogen depletion has been associated with the inability of Sprague-Dayley rat soleus muscle to maintain tension during a series of repeated contractions.[27] This association, however, occurs only during prolonged muscle activity of 2 hours or more.[27] To explain that greater decrease in tension during stimulation in aged rat FDB muscle compared with adult rat FDB muscle, other intramuscular factors, such as age-related changes in muscle mass, connective tissue concentration, excitation/contraction coupling, or sarcoplasmic reticulum [Ca.sup.+2] content and release, need to be investigated.

Alterations in aged human skeletal muscle metabolism are thought to be, in part, due to decreases in activity levels.28 The activity levels of the adult and aged rats were not measured in my study. To provide insight into whether activity levels might affect anaerobic metabolism in aging rat skeletal muscle, the findings of my study and those of a study by Fitts and colleagues[9] can be compared with findings of a study of a rat disuse muscle model.[29] In the study of the disuse model, high-energy phosphate utilization and lactate production increased during 30 minutes of stimulation of rat hind-limb muscles previously immobilized for 6 weeks.[29] Thus, the increase in anaerobic glycolysis in stimulated aged FDB muscle in my study and the increase in anaerobic glycolyses and the decrease in CP content in stimulated aged soleus muscle in Fitts and colleagues' study[9] appear to be similar to the anaerobic metabolism findings of the study of the rat disuse muscle model.[29] The increase in anaerobic metabolism with age during the stimulation bout may be due not only to age, but also to a decrease in the activity levels of the aged rats. Further investigation of the how an aging organism's activity level affects skeletal muscle anaerobic metabolism is warranted.

Investigations into the effect of age on metabolism and tension generation during supramaximal, intermittent stimulation have used in situ preparations.[8,9] When using an isolated in vitro muscle preparation, such as the FDB muscle preparation, inadequate oxygen supply and diffusion of substrates can affect metabolism and tension generation independent of other experimental variables such as age.[30] In my study, lactate concentrations following stimulation were higher in the incubation medium than in the muscles in both age groups. In addition, restitution of CP in the 12- and 32-month-old rats' stimulated FDB muscles during recovery was similar to restitution of CP in in situ exercised muscles during recovery.[31] These findings support previous findings[10,11,14,15,17] that the isolated in vitro FDB muscle preparation is viable for the study of metabolism and tension generation during a bout of electrical stimulation.

The results of my study and those of previous studies[8,9] of rats suggest that aged muscle has the capacity to rapidly, anaerobically resynthesize ATP during high-intensity resistance training. The question arises as to whether the metabolic findings in aged rat muscles can be applied to aged human muscle undergoing supramaximal neuromuscular electrical stimulation or performing bouts of high-intensity exercise during resistance training. Few studies have attempted to investigate the effect of age on human muscle anaerobic metabolism.[32-34] Larsson and Karlsson[32] found an age-related increase in human quadriceps femoris muscle endurance during 50 maximal dynamic contractions. Taylor et al,[33] using 1311P nuclear magnetic resonance to measure high-energy phosphate contents in the elbow flexors of individuals 70 to 80 years of age and young adults, found no change in resting concentrations of ATP and CP. The findings of Larsson and Karlsson[32] and Taylor et al[33] suggest that there is no age-related decline in the capacity of human muscle to produce ATP anaerobically during high-intensity resistance exercise. In vastus lateralis muscle biopsy samples from asymptomatic men and women, 52 to 79 years of age, intracellular concentrations of ATP and CP decreased 5%, however, compared with contents in muscle biopsy samples from men and women 40 years younger.[34] Moller et al[34] suggest that there may be an age-related decline in the ability of muscle to produce ATP anaerobically. Based on these limited findings,[32-34] further investigation of human muscle endurance and anaerobic metabolism during high-intensity, short-duration activity is necessary before definitive conclusions can be made.

Summary

From this study, it can be concluded that aged rat skeletal muscle has the capacity to resynthesize the ATP during brief, high-intensity resistance exercise. The results also suggest that inadequate ATP resynthesis through anaerobic metabolism does not appear to contribute to the decline in tension during supramaximal, intermittent electrical stimulation of aged rat skeletal muscle.

Acknowledgments

I thank Christine K Fleming and Sharon T McIlvain for their help in the biochemical analyses of muscle samples.

References

[1] Burke WE, Tuttle WW, Thompson CW, et al. The relation of grip strength and grip strength endurance to age. J Appl Physiol. 1954;5: 628-630. [2] Davies CTM, White MJ. Contractile properties of elderly human triceps surae. Gerontology. 1983;29:19-25. [3] Larsson L, Grimby G, Karlsson JO. Muscle strength and speed of movement in relation to age and muscle morphology. J Appl Physiol. 1979;46:451-456. [4] Vandervoort AA, McComas AJ. Contractile changes in opposing muscles of the human ankle joint with aging. J Appl Physiol. 1986;61: 361-367. [5] Frontera WR, Meredith CN, O'Reilly KP, et al. Strength conditioning in older men: skeletal muscle hypertrophy and improved function. J Appl Physiol. 1988;64:1038-1044, [6] Fiatarone MA, Marks EC, Ryan ND, et al. Strength conditioning in older men: skeletal muscle. JAMA. 1990;263:3029-3034. [7] Edwards RHT. Physiological analysis of skeletal muscle weakness and fatigue. Clin Sci Mol Med. 1978;54:463-470. [8] Campbell CB, Marsh DR, Spriet LL. Anaerobic energy provision in aged skeletal muscle during tetanic stimulation. J Appl Physiol. 1991; 70:1787-1795. [9] Fitts RH, Troup JP, Witzmann FA, et al. The effect of ageing and exercise on skeletal muscle function. Mech Ageing Dev. 1984;27:161-172. [10] Carlsen RC, Walsh DA. Decrease in force potentiation and appearance of alpha adrenergic mediated contracture in aging rat skeletal muscle. Pflugers Arch. 1987;408:224-230. [11] Carlsen RC, Larson DB, Walsh DA. A fast-twitch oxidative glycolytic muscle with a robust inward calcium current. Can J Physiol Pharmacol. 1985;63:958-965. [12] Information Notice to Recipients of NIA Aged Mice & Rats. Kingston, NY: Charles River Laboratories; August 1993. [13] Oscai LB, Palmer WK. Cellular control of triacylglycerol metabolism. In: Terjung RL, ed. Exercise and Spott Science Review. Philadelphia, Pa: Franklin Institute Press; 1983;11:1-23. [14] Hopp JF, Palmer WK. Electrical stimulation alters fatty acid metabolism in isolated skeletal muscle. J Appl Physiol. 1990;68:2473-2481. [15] Hopp JF, Palmer WK Effect of electrical stimulation on triacylglycerol in isolated skeletal muscle. J Appl Physiol. 1990;68:348-354. [16] McArdle WD, Katch Fl, Katch VL. Exercise Physiology: Energy, Nutrition, and Human Performance. Philadelphia, Pa: Lea & Febiger; 1991: 443-444. [17] Hopp JF, Palmer WK. Preliminary report: effect of glucose and insulin on triacylglycerol metabolism in isolated normal and diabetic skeletal muscle. Metabolism. 1991;40:223-225. [18] Crass MF, McCaskill ES, Granata AL. Effect of pressure development on glucose and palmitate metabolism in perfused heart. Am J Physiol. 1969;216:1569-1576. [19] Ivy JL, Young JC, McLane JA, et al. Exercise training and glucose uptake by skeletal muscle in rats. J Appl Physiol. 1983;55:1393-1396. [20] Lamprecht W, Stein P, Heinz F, et al. Creatine phosphate: determination with creatine phosphate, hexokinase, and glucose-6-phosphate dehydrogenase. In: Bergmeyer HU, ed. Methods of Enzymatic Analysis. New York, NY: Academic Press Inc; 1974:1777-1781. [21] Lo S, Russell JC, Taylor AW. Determination of glycogen in small tissue samples. J Appl Physiol. 1970;28:234-236. [22] Gutmann I, Wahlefeld AW. L- (+) lactate determination with lactate dehydrogenase and NAD. In: Bergmeyer HU, ed. Methods of Enzymatic Analysis. New York, NY: Academic Press Inc; 1974:1464-1468. [23] Bergmeyer HU, Brent E, Schmidt F, et al. D-glucose: determination with hexokinase and glucose-6-phosphate dehydrogenase. In: Bergmeyer HU, ed. Methods of Enzymatic Analysis. New York, NY: Academic Press Inc; 1974:1196-1201. [24] Dole VP. A relation between non-esterfied fatty acids in plasma and the metabolism of glucose. J Clin Invest. 1956;35:150-154. [25] Duncombe WG. The colormetric micro-determination of longchain fatty acids. Biochem J. 1963;88:7-10. [26] Steel RGD, Torrie JH. Principles and Procedures of Statistics. New York, NY: McGraw-Hill Inc; 1960:109-110, 308-310. [27] Fitts RH, Booth FW, Winder WW, et al. Skeletal muscle respiratory capacity, endurance, and glycogen utilization. Am J Physiol. 1975;38: 273-291. [28] Grimby G, Banneskiold-Samsoe B, Hvid K, et al. Morphology and enzymatic capacity in arm and leg muscles in 78- to 81-year-old men and women. Acta Physiol Scand. 1982;115:125-134. [29] Witzmann FA, Kim DH, Fitts RH. Effect of hindlimb immobilization on the fatigability of skeletal muscle. J Appl Physiol 1983;54:1242-1248. [30] Hill AV. Trials and Trials of Physiology. Baltimore. Md: Williams Wilkins; 1965:208-241. [31] Harris RC, Edwards RHT, Hultman E, et al. The time course of phosphotylcreatine resynthesis during recovery of the quadriceps muscle in man. Pflugers Arch. 1976;367:137-142. [32] Larsson L, Karlsson JO. Histochemical and biochemical changes in human skeletal muscle with age in sedentary males, age 22-65 years. Acta Physiol Scand. 1978;104:129-136. [33] Taylor DJ, Crowe M, Bore PJ, et al. Examination of the energetics of ageing skeletal muscle using nuclear magnetic resonance. Gerontology. 1984;30:2-7. [34] Moller P, Bergstrom J, Furst P, et al. Effect of aging on energy-rich phosphagens in human skeletal muscles. Clin Sci. 1980;58:553-555.

(*) Charles River Laboratories, Kingston, NY 12401. [dagger] Abbott Laboratories, Pharmaceutical Products Division, North Chicago, IL 60064. [double dagger] Sigma Chemical, PO Box 14508, St Louis, MO 63178. [sections] Eli Lilly and Company, Lilly Corporate Center, Indianapolis, IN 46285. [parallel] Aldrich Chemical, 1001 W Saint Paul Ave, Milwaukee, WI 53233.

JF Hopp, PhD, PT, is Associate Professor and Director of Physical Therapy, Carroll College, 100 N East Ave, Waukesha, WI 53186-9988 (USA) ([e-mail]). She was Assistant Professor, Department of Physical Therapy and the Section of Geriatric Medicine, Department of Medicine, University of Illinois at Chicago, Chicago, IL 60612, at the time this study was conducted.

This study was approved by the Animal Care Committee at the University of Illinois at Chicago.

This research was supported by National Science Foundation Grant No. 9009371.

This article was submitted June 1, 1994 and was accepted November 15, 1995.