Limb muscles from rats flown in space and after hindlimb unloading (HU) show an increased fatigability, and spaceflight has been shown to result in a reduced ability to oxidize fatty acids. The purpose of this investigation was to determine the effects of HU on the substrate content in fast- and slow-twitch fibers and to assess the substrate utilization patterns in single slow type I fibers isolated from control and HU animals. A second objective was to assess whether HU altered the ability of the heart or limb muscle to oxidize pyruvate or palmitate. After 2 wk of HU, single fibers were isolated from the freeze-dried soleus and gastrocnemius muscles. HU increased the glycogen content in all fiber types, and it increased lactate, ATP, and phosphocreatine in the slow type I fiber. After HU, the type I fiber substrate profile was shifted toward that observed in fast fibers. For example, fiber glycogen increased from 179 ± 16 to 285 ± 25 mmol/kg dry wt, which approached the 308 ± 23 mmol/kg dry wt content observed in the post-HU type IIa fiber. With contractile activity, the type I fiber from the HU animal showed a greater utilization of glycogen and accumulation of lactate compared with the control type I fiber. HU had no effect on the ability of crude homogenate or mitochondria fractions from the soleus or gastrocnemius to oxidize pyruvate or palmitate. The increased fatigability after HU may have resulted from an elevated glycolysis producing an increased cell lactate and a decreased pH.
- tissue oxidation
- adenosine 5′-triphosphate
a primary objective of the US and international space programs is to undertake a human explorative mission to Mars by the year 2014. For this to become a reality, the deleterious effects of microgravity on human biology must be solved. It is clear from the Skylab and more recently the Russian space station Mir and US Spacelab missions that the loss of bone and the loss of muscle mass in the microgravity environment pose two of the most serious problems inherent in prolonged space travel. The microgravity-induced muscle atrophy is associated with a reduced force and power and with an increased fatigability (3, 5, 7). In prolonged space missions, these cellular alterations could limit the crew's ability to work in space and/or on the surface of Mars and to rapidly egress in an emergency on return to earth.
When the rat soleus muscle was stimulated in situ after microgravity or after the spaceflight-model hindlimb unloading (HU), an increased rate and extent of fatigue were observed (3, 20). Additionally, HU has been shown to reduce the in vivo exercise run time to exhaustion and to increase the rate of glycogen depletion (26, 31). The observed change in substrate utilization during submaximal exercise was not caused by a change in muscle blood flow (20). HU did induce a decline in maximal cardiac output (31), and HU animals failed to fully redistribute blood from visceral tissues to the working skeletal muscles (19). Thus muscle blood flow during maximal exercise would likely be reduced compared with control animals.
After a 9-day exposure to microgravity, Baldwin et al. (2) observed a 37% decline in the ability of both the high- and low-oxidative regions of the vastus muscle to oxidize long-chain fatty acids. In contrast, they found no change in this muscle's ability to oxidize pyruvate or in key marker enzymes of the tricarboxylic acid cycle, the malate shuttle, or β-oxidative pathway. Collectively, the observations from microgravity and HU experiments suggest that working skeletal muscles have an increased dependence on carbohydrates and a reduced ability to oxidize fats after unloading. Both HU and spaceflight have been shown to increase the percentage of fast-twitch fibers in the slow rat soleus muscle from ∼15 to ∼30% (4, 18). Consequently, the increased carbohydrate utilization during exercise may simply result from an increased population of fast fibers that are known to have a higher glycolytic capacity. Alternatively, unloading may induce an increased utilization of muscle glycogen in the slow type I fiber.
The purpose of this investigation was to determine 1) the extent to which 2 wk of HU alters the substrate profile of individual slow- and fast-twitch fibers in rat hindlimb muscles; 2) whether the slow type I fiber utilizes more glycogen and produces higher lactate during 10 min of in situ contractile activity; and3) the effect of HU on the ability of the soleus and gastrocnemius to oxidize fats and carbohydrates under optimal substrate conditions.
MATERIALS AND METHODS
Animal care and suspension procedure.
Male Sprague-Dawley rats (∼275 g) were obtained from Sasco (Madison, WI) and randomly assigned to either the HU or cage-control group. The hindlimb unloading was carried out exactly as described previously (10). The hindlimbs of the HU animals were elevated for 14 days with use of a harness attached to the proximal two-thirds of the tail. The height of suspension was adjusted to prevent the hindlimbs from contacting supportive surfaces. The forelimbs maintained contact with a grid floor, which allowed the animals to move about to obtain food and water. The HU animals were fed Purina rat chow and water ad libitum, whereas the control rats were pair fed to maintain weights similar to those of the HU group. Both groups were housed at 23°C with a 12:12-h light-dark cycle.
Experimental design and muscle preparation.
In this investigation three experiments were conducted. Inexperiment 1, three control and three HU animals were studied to establish the resting substrate profile for the slow type I, fast type IIa, and fast type IIx/IIb fiber types. In the second experiment, the soleus muscles from control and HU animals were setup in situ and electrically stimulated for 0, 2, 5, or 10 min as described below. Three control and three experimental animals were assigned to each stimulation duration. In the third experiment, 16 control and 16, 14-day HU animals were utilized to determine the effect of HU on substrate oxidation. For all three experiments, the control and 14-day HU animals were anesthetized with pentobarbital sodium (50 mg/kg body wt ip). In experiment 1, the soleus and gastrocnemius muscles were excised from the left hindlimb and placed on ice (4°C). The soleus (source of type I fibers) was divided into four equal-sized bundles, which in turn were aligned longitudinally on small (2.5 × 0.5-cm) index cards and quick frozen in liquid nitrogen. Bundles were similarly prepared from the deep red (source of type IIa fibers) and superficial white (source of type IIx/IIb fibers) regions of the lateral and medial heads of the gastrocnemius, respectively. The frozen samples were then freeze-dried under vacuum at −35°C and stored under vacuum at −80°C. Individual fiber segments (∼2 mm long) were dissected free at room temperature and stored separately under vacuum at −80°C.
In experiment 2, the in situ soleus muscle preparation was prepared exactly as described previously (20). The soleus muscle was exposed and dissected free of surrounding tissue, with its blood and nerve supply intact. The isolation of the soleus involved dividing the gastrocnemius into its medial and lateral heads and carefully pulling back the two heads of the gastrocnemius and the plantaris. These muscles were denervated such that they did not contribute to the force developed during activation of the soleus. A silk thread (4-0) was secured to the distal end of the soleus tendon under the cut ends of the gastrocnemius tendon such that the latter acted as stop to prevent slippage of the tie. A small loop was tied in the thread and used for attachment of the preparation to a model 305 Cambridge Technology force transducer.
The soleus muscle was stimulated indirectly through the distal portion of the cut sciatic nerve exactly as described previously (20). Fatigue was produced by applying 100-Hz, 100-ms supermaximal trains at a train rate of 120/min. This protocol has previously been shown to induce significant fatigue in both groups with a greater decline in train force in the HU muscles (20). After 0, 2, 5, or 10 min of stimulation, the muscles were quick frozen with small tongs cooled to liquid nitrogen temperatures. The frozen samples were then divided longitudinally into three or four segments, freeze-dried, and single fiber segments isolated and stored as described above.
The individual fibers isolated from experiment 1 animals were assayed for glycogen, lactate, ATP, and phosphocreatine (PCr), whereas, for experiment 2, the fibers were assayed for glycogen and lactate. The individual fibers were warmed to room temperature, and divided into three pieces. The individual pieces were weighed on a quartz-fiber balance and used to assay for fiber high-energy phosphates (ATP and PCr), glycogen, or lactate (29). One piece was always run on 5 and 12% SDS-PAGE gels for fiber-type determination. The gel procedures were exactly as described by McDonald et al. (18). Because of difficulty in consistently separating the type IIx and type IIb myosin heavy chain, these fibers were considered as a single group. The assays were conducted as described by Thompson and Fitts (29) by using modifications of the assays published by Passonneau and Lowry (22). The procedures employ enzyme methods based on the fluorometric determination of pyridine nucleotides. Sensitivity was increased by enzymatic cycling (22), and concentrations were expressed as millimoles per kilogram dry weight.
Determination of pyruvate and palmitate oxidation rates.
In these experiments, substrate oxidation rate was determined by using crude homogenate and isolated mitochondria. The soleus, red (RG) and white (WG) regions of the gastrocnemius, and heart muscles were removed from the three experimental animals and were placed on ice. The deep region of the lateral head and superficial region of the medial head of the gastrocnemius were used for preparation of the RG and WG sample (1). The tissues were weighed, finely minced, and homogenized in buffer (1:10; wt/vol) containing 175 mM KCl and 2 mM EDTA (pH 7.2). The homogenate was either used for tissue respiration studies or mitochondrial isolation. The mitochondrial were isolated exactly as described by Terjung et al. (27).
The tissue (crude homogenate or isolated mitochondria) oxygen uptake with pyruvate or palmitate as substrate was determined at 30°C by using a Clark oxygen electrode and Yellow Springs Instruments model 5301 temperature bath. For pyruvate oxidation, 2.8 ml of the reaction cocktail were added to the reaction vessel and equilibrated at 30°C. The reaction cocktail contained (in mM) 5 MgCl2, 100 KCl, 50 potassium phosphate buffer, 6.9 EDTA, and 0.0075 cytochromec. For the soleus, RG and heart 20 mg of tissue, and for the WG 40 mg of tissue, were added to the reaction and equilibrated for 3 min. The reaction was initiated by the addition of 200 μl of 100 mM pyruvate and 200 μl of 20 mM ADP, and the oxygen uptake was recorded by using a MacLab computer.
The reaction medium for the determination of palmitate oxidation was exactly as described By Molé et al. (21) except the oxygen consumption was measured by using a Clark electrode. The tissue amount added was the same as for the pyruvate assay, and the concentration of palmitic acid, CoA, and l-carnitine was 0.75 mM, 25 μM, and 0.5 mM, respectively. The reaction was initiated by the addition of substrate, CoA, and ADP to the media (30°C and pH 7.2), and oxygen uptake recorded by using a MacLab computer.
Data are presented as means ± SE. Treatment effects for each muscle were analyzed with a one-way analysis of variance. When significantF ratios were obtained, a post hoc Student's unpaired two-tailed t-test was employed using the 0.05 level of confidence.
Substrate profile of individual fast- and slow-twitch fibers. The mean glycogen, lactate, ATP, and PCr concentration of individual type I, IIa, and IIb fibers are shown in Table1. The primary difference between fiber types was the significantly higher high-energy phosphate content (ATP and PCr) in the fast- compared with the slow-twitch fiber types. Although all four substrates were higher in the fast type IIb compared with the fast type IIa fibers, and the fast fibers contained higher glycogen than did the slow type I fibers, these differences were not significant at the <0.05 level.
The major effect of the 2-wk HU was a significant increase in all four substrates in the slow type I fiber and an increase in glycogen in the fast fiber types. In fast fibers, the other substrates increased, but, except for PCr in the type IIa fiber, the increases were not significant (Table 1). The HU-induced increase in the ATP and PCr content of the slow type I fiber caused this population of fibers to resemble fast type II fibers from the perspective of their high-energy phosphate profile. This fact can be clearly seen in Fig.1 where PCr is plotted against ATP content for individual fibers isolated from the soleus of control and HU animals. For comparison, fibers representing the range of ATP and PCr values for individual fast fibers are shown.
Glycogen and lactate changes with muscle fatigue.
By using the exact stimulation protocol used here, we have previously demonstrated that 2 wk of HU significantly increased both the rate and extent of soleus muscle fatigue (20). Figure2 confirms the result obtained inexperiment 1 (Table 1) that HU significantly increased soleus type I fiber glycogen (178 ± 10 vs. 213 ± 18 mmol/kg dry wt for an observation number of 65 and 41 fibers, respectively). Importantly, Fig. 2 also shows that the soleus type I fibers from the HU animals metabolized glycogen at a faster rate and reached a significantly lower glycogen content after 10 min of electrical stimulation (71 ± 5 vs. 36 ± 5 mmol/kg dry wt). The effect of HU on the initial rate of glycogen utilization is best illustrated by comparing the difference between the 0- (prestimulation) and 2-min stimulation time points (Fig. 2). The mean glycogen used during this period was 54 and 125 mmol/kg dry wt for the control and HU groups, respectively.
The type I fibers from the HU animals also contained significantly higher resting lactate levels (time 0 data, Fig. 3). Consistent with an increased rate of glycogen utilization, the HU group showed an increased lactate production with stimulation such that after 10 min of contractile activity lactate levels were 54 ± 7 and 90 ± 9 mmol/kg dry wt for the control and HU group, respectively (Fig.3).
Whole muscle pyruvate and palmitate oxidation.
For both the crude homogenate and isolated mitochondrial preparations, oxidation rate was highest in the heart and lowest for the WG (Table2). HU had no significant effect on the ability of the heart, soleus, or WG to oxidize pyruvate or palmitate, and this was true for both the crude homogenate and isolated mitochondrial preparations (Table 2). In the case of palmitate oxidation by the crude homogenate, there was a small but nonsignificant decline after HU in all muscles; the decrease in the RG was significant at the P < 0.1 level. Mitochondria isolated from the RG showed a significantly higher pyruvate oxidation but no significant change in palmitate oxidation after HU (Table 2).
Resting substrate profile.
The mean soleus type I fiber glycogen content observed here was in good agreement with previously published values obtained from whole muscle analyses (15). Additionally, the type I fiber glycogen appears to be similar across species lines. For example, when slow soleus type I fibers were assayed under identical conditions the mean fiber glycogen content was 179, 148, and 217 mmol/kg dry wt for rats, monkeys, and humans, respectively (11, 12). For fast fibers isolated from the gastrocnemius, the glycogen levels averaged 218, 239, and 227 mmol/kg dry wt for these same three species (11, 12). These comparisons demonstrate that, when fiber glycogen is assayed under identical conditions and without carbohydrate loading or food restriction, species differences are minimal and the fast fibers have somewhat higher glycogen contents (24).
In agreement with the results of Hintz et al. (16), the rat fast fibers contained higher ATP and PCr content than did the slow type I fiber type (Table 1). In contrast, larger species such as nonhuman primates and humans demonstrate no significant difference in high-energy phosphate content between fiber types (12, 17). When rats are compared with the larger monkeys or humans, it is apparent that the fiber ATP content is greater in the rat. For example, we found the rat IIa and IIb fiber ATP to be 37 and 40 mmol/kg dry wt, while human and monkey fast fibers averaged 20 and 22 mmol/kg dry wt, respectively.
Effect of HU on fiber substrate profile.
It is apparent from the results in Table 1 and Fig. 1 that HU-induced a substrate pattern in the slow type I fiber that more resembles that observed in fast fibers. This substrate change was not a result of fiber type shifting because all of the slow fibers in Table 1 contained only the slow myosin isozyme. The increased slow type I fiber glycogen is consistent with the results of Henriksen et al. (13) and Henriksen and Tischler (15) that demonstrated an increased soleus muscle glycogen after HU. Our results demonstrate that the increase in fiber glycogen with HU occurs in fast- as well as slow-twitch fibers. Henriksen and Tischler have shown that the HU-induced increase in muscle glycogen occurs rapidly and reaches a maximum by 24 h. The increased muscle glycogen occurred despite a decline in the rate of glucose uptake (14). It has been suggested that the HU-induced increase in muscle glycogen can be attributed to the reduced contractile activity during HU, which inhibits glycogen phosphorylase and glycolysis (15). Additionally, the increased substrate content of the type I fiber after HU could in part reflect selective atrophy of the contractile proteins, such that, the concentration of other cell components increased (9, 28).
Muscle fatigue and substrate changes with electrical stimulation.
We have previously shown that HU increases the fatigability of the soleus (20), and Caiozzo et al. (3) demonstrated a similar effect after spaceflight. Although different stimulation protocols were used in these studies, the extent of fatigue was similar. The control groups showed 66 and 64% and the experimental groups 48 and 36% of the initial force for the HU and the flight studies, respectively (3, 20). An important question is what contributes to the accelerated fatigue after spaceflight or HU. Our hypothesis is that both spaceflight and HU cause an accelerated glycolysis with the onset of contractile activity. Although the soleus muscle blood flow achieved during the electrical stimulation was not altered by HU (20), the possibility exists that the initial rate of rise in blood flow was slowed after HU. If this occurred, one would expect to observe a greater rate of decline in PCr and rise in Pi. The latter along with an elevated cell glycogen would activate phosphorylase and increase the glycolytic rate. The elevated lactate production observed in this study, and an increased phosphofructokinase activity after HU supports this hypothesis (9). An increased Pi would contribute to fatigue by directly inhibiting the transition of the cross bridge from the low to the high-force state (8). It will be important for future studies to determine whether HU accelerates the rise in muscle Piduring the onset of contractile activity. In addition to Pi, a decreased cell pH is also known to directly inhibit cross-bridge force (8). Although pH was not measured in this study, a high inverse correlation is known to exist between muscle lactate and intracellular pH (25). Thus the 90 mmol/kg dry wt lactate in the fatigued type I fibers of the HU animals was likely associated with a lower pH than the control group.
During maximal exercise, the increased fatigability after spaceflight and HU would likely be exacerbated because the maximal cardiac output is known to be depressed by both conditions (5, 31). Furthermore, after HU the redistribution of blood flow from the internal organs to the working skeletal muscles is blunted (19). The reduced cardiac output and incomplete redistribution of blood flow would combine to lower skeletal muscle blood flow during maximal exercise. This would lead to an increased rate of glycolysis and of lactate, H+, and Pi production.
Substrate oxidation and muscle fatigue.
The increased fatigue after HU and spaceflight was not likely caused by a depressed oxidative enzyme capacity. We have previously demonstrated that HU in rats and spaceflight in humans and monkeys has no effect on marker enzymes of either the Krebs cycle or β-oxidative pathways (Ref. 9; unpublished observations). Additionally, we found no significant change in the capacity of crude homogenates of the soleus or gastrocnemius muscles to oxidize palmitate or pyruvate. In contrast, after a 9-day spaceflight, Baldwin et al. (2) observed a 37% decline in the capacity of limb skeletal muscle to oxidize long-chain fatty acids. Similar to HU, they observed no effect of spaceflight on the ability of rat skeletal muscle to oxidize pyruvate. These results suggests that spaceflight inhibited the ability of limb skeletal muscle to oxidize fatty acids. Because the inhibition was observed under optimal substrate conditions, the result suggests that some enzyme(s) involved in fatty acid oxidation was depressed. The reduced fatty acid oxidation might result from a depression in carnitine palmitoyltransferase (CPT I), a rate-limiting enzyme for fatty acid oxidation (6).
The differences between our result and those of Baldwin et al. (2) might be explained by a fundamental difference between HU and spaceflight or alternatively by the different methodologies employed. Baldwin et al. measured the capacity of the whole homogenate to oxidize [U-14C]palmitate, and thus the oxidation of this substrate could be evaluated independently of any endogenous substrate. In our preparation, endogenous carbohydrate may have contributed to the total substrate oxidized, thus masking any reduction in palmitate oxidation in the post-HU muscles. To test this possibility, we assayed the capacity of isolated mitochondria to oxidize pyruvate and palmitate. In agreement with the crude homogenate data, HU had no effect on the maximal respiratory rate with either substrate. The mitochondrial yields were low (∼5% of the total pool), and thus the possibility exists that the mitochondria assayed were not representative of the entire cell population. As described above, the increased utilization of muscle glycogen and lactate production after HU may have resulted from substrate level (high Pi and glycogen) activation of glycolysis. The failure to observe any significant decline in palmitate oxidation suggests that the enzymes required for optimal fat oxidation were not altered. Thus if HU does inhibit fatty acid oxidation, the mechanism likely involves substrate- level control rather than direct effects on the enzyme content of the muscle. Winder and colleagues (30) and Rasmussen and Winder (23) have suggested that fatty acid oxidation increases with the onset of contractile activity due to a decline in malonyl-CoA, a known inhibitor of CPT I. Thus one possibility is that HU increased glycolysis and the production of acetyl-CoA. Because acetyl-CoA acts as a substrate for malonyl-CoA, this could reduce the exercise-induced decline in malonyl-CoA, maintaining the inhibition of CPT I and fatty acid oxidation.
The primary findings of this investigation were that after HU the slow type I fiber of the soleus showed a substrate profile shifted toward that of a fast fiber and that when stimulated it utilized more glycogen and accumulated higher levels of lactate than did type I fibers from control animals. We hypothesize that the increased fatigability after HU was at least in part due to an increased glycolysis producing an increased Pi, lactate, and free H+. The apparent increased dependence on glycogen metabolism during contractile activity could not be attributed to a reduced ability to oxidize carbohydrates or fatty acids, and thus it likely resulted from substrate level activation of glycolysis and inhibition of fatty acid oxidation.
This study was supported by National Aeronautics and Space Administration Grant NAGW-4376 (R. H. Fitts). K. R. Kidd was supported by a Howard Hughes Medical Institute undergraduate research program.
Address for reprint requests and other correspondence: R. H. Fitts, Marquette University, Dept. of Biology, Wehr Life Sciences Bldg., P.O. Box 1881, Milwaukee, WI 53201-1881.
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