We examined whether skeletal muscle transport rates of long-chain fatty acids (LCFAs) were altered when muscle activity was eliminated (denervation) or increased (chronic stimulation). After 7 days of chronically stimulating the hindlimb muscles of female Sprague-Dawley rats, the LCFA transporter proteins fatty acid translocase (FAT)/CD36 (+43%) and plasma membrane-associated fatty acid-binding protein (FABPpm; +30%) were increased (P < 0.05), which resulted in the increased plasmalemmal content of these proteins (FAT/CD36, +42%; FABPpm +13%, P < 0.05) and a concomitant increase in the LCFA transport rate into giant sarcolemmal vesicles (+44%, P < 0.05). Although the total muscle contents of FAT/CD36 and FABPpm were not altered (P > 0.05) after 7 days of denervation, the LCFA transport rate was markedly decreased (−39%). This was associated with reductions in plasmalemmal FAT/CD36 (−24%) and FABPpm (−28%; P < 0.05). These data suggest that these LCFA transporters were resequestered to their intracellular depot(s) within the muscle. Combining the results from these experiments indicated that changes in rates of LCFA transport were correlated with concomitant changes in plasmalemmal FAT/CD36 and FABPpm, but not necessarily with their total muscle content. Thus chronic alterations in muscle activity can alter the rates of LCFA transport via different mechanisms, either 1) by increasing the total muscle content of FAT/CD36 and FABPpm, resulting in a concomitant increase at the sarcolemma, or 2) by reducing the plasma membrane content of these proteins in the absence of any changes in their total muscle content.
- giant vesicles
- tibialis anterior
- chronic stimulation
skeletal muscles depend on long-chain fatty acids (LCFAs) to maintain ATP production during contractile activity. When muscle activity is increased, progressively more LCFAs are taken up into the muscle (16). It is now recognized that LCFAs cross the sarcolemma via simple diffusion and a protein-mediated system (3, 4, 35, 47). Several proteins have been identified as LCFA transporters. Fatty acid translocase (FAT/CD36) and plasma membrane-associated fatty acid-binding protein (FABPpm) have been shown to facilitate LCFA transport into heart and skeletal muscle (3, 24, 38, 47), whereas fatty acid transport protein 1 (FATP1) appears to be involved in the transport of LCFAs and acylating very long-chain fatty acids (15, 45, 53, 58).
Skeletal muscle metabolism is remarkably capable of adapting to changes in muscle activity pattern (7, 44a). Chronic changes in muscle activity can also alter the expression of several substrate transporters, and hence substrate transport. For example, increasing muscle activity by chronic stimulation enhances glucose and lactate transport and their respective transporters GLUT4 (28, 46, 56) and the monocarboxylate transporter MCT1 (6, 41, 42). Conversely, reductions in muscle activity, induced by denervation, result in decrements in glucose and lactate transport and in their accompanying transporters GLUT4 (14, 23, 43, 54), MCT1, and MCT4 (6, 40, 55). Although it has been shown that increased muscle activity can increase FAT/CD36 (1) and FABPpm (29, 51), as well as LCFA transport (1), the effects of reducing muscle activity on LCFA transport and transporters are not known.
In previous studies, we have shown (3) that LCFA uptake is subject to short-term regulation by a brief period (5–30 min) of muscle contraction, involving the translocation of FAT/CD36 from intracellular stores to the sarcolemma. Subsequently, we demonstrated that insulin is also able to translocate FAT/CD36 in muscle (34) and cardiac myocytes (37) via the phosphatidylinositol 3-kinase-signaling pathway (34, 37). The total muscle content of LCFA transporters has been shown to be a key factor influencing the plasmalemmal content of LCFA transporters, and hence the rate of LCFA transport in skeletal muscle (32, 48). However, we (33a) have also shown, in obese Zucker rats, that rates of LCFA transport into muscle and heart can be increased by a permanent relocation of FAT/CD36 from their intracellular depots to the plasma membrane, without any changes being observed in the total muscle content of this transporter. Thus, just as for glucose transport and GLUT4 (see Ref. 20), it is important to account for changes in LCFA transport by examining both the total muscle content and the plasmalemmal content of FAT/CD36 and FABPpm.
Because LCFAs are an important substrate for skeletal muscle, and the rate of LCFA transport can regulate the cellular metabolism of this substrate in this tissue (25), we examined the effects of altered muscle activity on LCFA transport and the mechanisms involved. Specifically, we investigated the effects of an increase in muscle activity (7 days of chronic stimulation) and a decrease in muscle activity (7 days of denervation) on 1) the total muscle content of the LCFA transport proteins FAT/CD36 and FABPpm, 2) their plasmalemmal content, and 3) the rates of LCFA transport into giant sarcolemmal vesicles. The contralateral muscles, in each of these treatments, served as controls.
Bovine serum albumin (BSA) (fraction V) and collagenase type VII were purchased from Sigma-Aldrich (St. Louis, MO). Nonfat dry milk and Western blot reagents were from Bio-Rad Laboratories (Hercules, CA), and the enhanced chemiluminescence (ECL) kit was from Amersham Pharmacia Biotech (Buckingham, UK). FAT/CD36 was detected with a monoclonal antibody (MO25) directed against human CD36 (39). A rabbit polyclonal antibody against rat hepatic membrane-associated LCFA-binding protein was used to detect FABPpm (10). A goat polyclonal immuno-A purified GLUT4 antibody and donkey anti-goat horseradish peroxidase (HRP)-conjugated IgG (HRP-IgG) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Goat anti-mouse and donkey anti-rabbit HRP-IgG were obtained from Santa Cruz Biotechnology and Amersham Pharmacia Biotech, respectively.
Female Sprague-Dawley rats weighing 200–250 g were used. Animals were housed in a temperature-controlled room on a reversed 12:12-h light-dark cycle and fed a Purina Chow diet and water ad libitum. Immediately before surgery, rats were anesthetized with halothane, followed by a subcutaneous injection of buprenorphine hydrochloride (0.3 mg/ml; 0.12 μl/kg). After surgery, animals were housed individually (chronic stimulation) or were housed together in a cage (denervation). Ethical approval for all experimental procedures was obtained from the Committee on Animal Care at the University of Guelph.
Chronic Stimulation of Rat Hindlimb Muscles
Rat muscles were prepared and chronically stimulated as we have previously described (1, 6, 28, 41, 42). For these purposes we have routinely stimulated the extensor digitorum longus (EDL) and red and white tibialis anterior (TA) (6, 42). In anesthetized rats, two stainless steel electrodes were sutured to the underlying muscle on either side of the common peroneal nerve. These electrodes were then passed subcutaneously from the thigh, exteriorized at the back of the neck, and subsequently attached to a miniature electronic stimulator. The overlying muscle was sutured and the skin was stapled. Muscles from the contralateral limb were used as a nonstimulated, internal control and were therefore sham operated. Only when animals had regained at least 100% of their preoperative body weight (5 days) and had recovered from surgery for a minimum of 6–7 days was chronic stimulation initiated. The common peroneal nerve, which innervates the EDL and TA muscles, was stimulated at 12 Hz, 24 h/day for 7 days. Thereafter, animals were subdivided into two groups. In the first group (n = 5), total FAT/CD36, FABPpm, and GLUT4 protein levels were measured in chronically stimulated and control EDL and red and white TA muscles. In a second group of rats (n = 15), the EDL and TA muscles were pooled (n = 3 rats/pool) for the purpose of preparing giant sarcolemmal vesicles to measure rates of palmitate uptake. FAT/CD36, FABPpm, and GLUT4 (positive control) were measured in plasma membranes of giant vesicles prepared from these pooled control and chronically stimulated muscles.
Denervation of Rat Hindlimb Muscles
Muscle activity was eliminated in one hindlimb by denervating the lower leg muscles of that hindlimb. For these purposes we have routinely examined soleus and red and white gastrocnemius in our work (40, 54, 55). Briefly, a small superficial incision was made in the posterior aspect of the hindlimb to expose the sciatic nerve, a 1-cm section of the sciatic nerve was removed, and the incision was closed with sutures and staples. The contralateral leg was sham operated without touching the nerve. Muscles in this limb served as a nondenervated control. After denervation, rats were randomly divided into two groups. In the first group (n = 10), FAT/CD36, FABPpm, and GLUT4 protein levels were measured in denervated and control soleus and red and white gastrocnemius muscles. In a second group of rats (n = 12), the denervated muscles were pooled (n = 2 rats/pool) to prepare giant sarcolemmal vesicles to measure rates of palmitate uptake by giant vesicles. FAT/CD36, FABPpm, and GLUT4 (positive control) were measured in plasma membranes of giant vesicles prepared from control and denervated hindlimb muscles.
Preparation of Giant Sarcolemmal Vesicles
Giant vesicles from control, denervated, and chronically stimulated skeletal muscles were prepared as previously described (2, 3, 30). Briefly, muscles were cut into thin layers (∼1- to 3-mm thick) and incubated for 1 h at 34°C in 140 mM KCl, 10 mM MOPS (pH 7.4), aprotinin (10 mg/ml), and collagenase type VII (150 U/ml) in a shaking water bath. The tissues were then washed with KCl-MOPS and 10 mM EDTA, and the supernatant was collected. Percoll (final concentration 16%) and aprotinin were added to the supernatant. This supernatant was placed at the bottom of a density gradient consisting of a 3-ml middle layer of 4% Nycodenz (wt/vol) and a 1-ml KCl-MOPS upper layer. The samples were centrifuged (GS-15 centrifuge; Beckman, Palo Alto, CA) at 60 g for 45 min at room temperature. After centrifugation, the vesicles were harvested from the interface of the two upper solutions. The vesicles were diluted in KCl-MOPS and recentrifuged (Sorvall MC 12V; DuPont-Mundell Scientific, Guelph, ON, Canada) at 12,000 g for 5 min. Vesicles (∼50 μg) were stored at −80°C until analyzed by Western blotting for FAT/CD36, FABPpm, and GLUT4.
Palmitate Uptake by Giant Vesicles
Palmitate uptake was measured as described previously (3, 4, 30). Briefly, unlabeled and radiolabeled 0.3 μCi [9,10-3H]palmitate and 0.06 μCi [14C]mannitol in a 0.1% BSA KCl-MOPS solution were added to 40 μl of vesicles (∼80 μg protein). The reaction was carried out at room temperature for 15 s. Palmitate uptake was terminated by addition of 1.4 ml of ice-cold KCL-MOPS-2.5 mM HgCl-0.1% BSA. The sample was quickly centrifuged at maximal speed in a microcentrifuge for 1 min. The supernatant was discarded, and radioactivity was measured in the tip of the tube.
Sample Preparation for Western Blotting
Samples were prepared as described in detail elsewhere (3, 4). Briefly, total tissue homogenates were prepared from individual muscles. For these purposes, tissues (∼60 mg) were homogenized in 2 ml of buffer A [in mM: 210 sucrose, 2 EGTA, 40 NaCl, 30 HEPES, 5 EDTA, 2 phenylmethylsulfonyl fluoride (PMSF), pH 7.4] for two interrupted 15-s bursts with a polytron homogenizer (Kinematica, Littau, Switzerland) set at 7. Subsequently, 2 ml of buffer A and 3 ml of buffer B (1.167 M KCl, 58.3 mM tetrasodium pyrophosphate) were added, mixed briefly, and set on ice for 15 min. After centrifugation (XL-90 ultracentrifuge, Beckman) at 50,000 rpm for 75 min at 4°C, the supernatant fluid was discarded, and the pellet was washed thoroughly with 1–2 ml of buffer C (10 mM Tris base, 1 mM EDTA, pH 7.4). The pellet was resuspended in 600 μl of buffer C and homogenized for two interrupted 10-s bursts with a polytron set at 7. Then 200 μl of 16% SDS were added, and samples were removed from ice, vortex mixed, and centrifuged (Sorvall MC 12V, DuPont) at 3,000 rpm for 15 min at room temperature. The supernatant was divided into aliquots, and protein concentration was determined in triplicate by the bicinchoninic acid assay (Sigma, St. Louis, MO) with BSA as standard. Samples were stored at −80°C for immunoblot detection of FAT/CD36, FABPpm, and GLUT4. For detection of proteins by Western blotting, 10 μg of muscle and vesicle protein samples were separated on 10% SDS-polyacrylamide gels (150 V, 1 h). Proteins were then transferred to Immobilon polyvinylidene difluoride membranes (Bio-Rad Laboratories, Hercules, CA) by use of a Trans-Blot SD-Dry Transfer Cell (Bio-Rad Laboratories). Membranes were incubated overnight at 4°C, and immune complexes were detected with ECL (Amersham Pharmacia Biotech). Blots were quantified on a densitometer connected to a computer with appropriate software. Blots were normalized across different gels by using a standard sample and arbitrarily setting control muscle optical density readings to 100 in each gel. These are typical normalization procedures used in our laboratory (3–6, 32–34).
The transport data, as well as the plasma membrane protein content (i.e., separate analyses for FAT/CD36 and FABPpm), were each analyzed using a two-factor (stimulation vs. denervation) repeated-measures (control muscles vs. experimental muscles) analysis of variance (ANOVA). For muscles with the same fiber composition in the two experiments [i.e., muscles rich in either fast-twitch oxidative glycolytic (FOG) or fast-twitch glycolytic (FG) fibers], the homogenate Western blotting data were also analyzed with a two-factor (stimulation vs. denervation) repeated-measures (control muscles vs. experimental muscles) ANOVA. Because some muscles examined do not have the same muscle fiber composition [i.e., chronic stimulation: EDL with mixed FG and FOG; denervation: soleus, primarily slow-twitch oxidative (SO)], we analyzed each of these muscles independently, using a repeated-measures (control muscles vs. experimental muscles) ANOVA. Linear regression analyses were performed to compare plasmalemmal protein content with rates of palmitate transport. All data are presented as means ± SE.
Animal Body Weights
Within the first 3 days after the surgery, the rats lost ∼10% of their body weight. Thereafter, they regained weight, and after 7 days of denervation animals had regained 100% of their presurgical body weight (data not shown). In the chronic stimulation experiments, rats had regained their presurgical body weight after 5 days (data not shown). During the 7 days of chronic stimulation, the rats continued to gain body weight (data not shown), as we have reported previously (6, 40, 41).
Rates of Palmitate Uptake and Muscle and Plasmalemmal Content of FAT/CD36 and FABPpm
Palmitate uptake by giant sarcolemmal vesicles.
Because tissue requirements for preparation of giant vesicles are relatively high, it was necessary to pool the red and white denervated muscles and the red and white chronically stimulated muscles. There were considerable differences in LCFA uptake rates in vesicles from chronically stimulated and denervated hindlimb muscles. After 7 days of chronic stimulation, the uptake of palmitate by giant sarcolemmal vesicles was significantly increased (+44%; P < 0.05; Fig. 1) compared with the contralateral control muscles. In contrast, after 7 days of denervation, there was a marked decrease (−39%) in the uptake rate of palmitate compared with contralateral control muscles (P < 0.05; Fig. 1).
FAT/CD36 and FABPpm protein total muscle content in chronically stimulated and denervated muscles.
Because the muscle fiber composition of the red and white gastrocnemius is similar to that of the red and white TA used in the chronic stimulation studies (5, 42, 43), comparisons between chronically stimulated (red and white TA) and denervated muscles (red and white gastrocnemius) are reported according to their fiber composition, where FOG designates the red gastrocnemius and red TA, and FG designates the white gastrocnemius and white TA. Because we also examined the EDL muscle in the chronic stimulation experiments and soleus muscle in the denervation experiments, we have also reported the data using their fiber composition (EDL: FG/FOG; soleus: SO).
After 7 days of chronic stimulation, the total muscle content of FAT/CD36 was increased +66% in the FG/FOG, +77.5% in the FG and +18% in the FOG muscles (P < 0.05; Fig. 2A). FABPpm total muscle content was also increased in the chronically stimulated muscles (FG/FOG +22.5%; FG +38.8%; FOG +30%; P < 0.05; Fig. 2B). In contrast, 7 days of denervation failed to alter the total muscle content of either FAT/CD36 or FABPpm in any of the muscles examined (P > 0.05; Fig. 2, C and D). As a positive control we also measured the total muscle content of GLUT4. This protein was increased in chronically stimulated muscles and reduced in denervated muscles (data not shown), as we (28, 43, 46, 54) have previously reported in these two models.
Changes in plasmalemmal FAT/CD36 and FABPpm in chronically stimulated and denervated muscles.
The sarcolemmal content of FAT/CD36 and FABPpm was measured on plasma membranes of giant vesicles prepared from chronically stimulated and denervated pooled rat hindlimb muscles, as well as from pooled contralateral control muscles. After 7 days of chronic stimulation, both plasmalemmal FAT/CD36 (+42%) and plasmalemmal FABPpm (+13%) were significantly increased (P < 0.05; Fig. 3). In contrast, in denervated muscles, plasmalemmal FAT/CD36 (−24%) and FABPpm (−28%) were reduced (P < 0.05; Fig. 3).
Comparison of LCFA transport and plasma membrane FAT/CD36 and FABPpm.
Since the plasmalemmal content of the LCFA transporters is expected to be related to rates of LCFA transport, we compared the plasmalemmal FAT/CD36 and FABPpm with rates of palmitate uptake into giant vesicles prepared from chronically stimulated and denervated hindlimb muscles. Because the LCFA transport rates in the control vesicles did not differ in the chronically stimulated and denervation groups, the control data were pooled. There was a highly linear relationship between plasma membrane FAT/CD36 and rates of palmitate transport into giant sarcolemmal vesicles (Fig. 4A) and between plasma membrane FABPpm and rates of palmitate transport into giant sarcolemmal vesicles (Fig. 4B).
By examining chronically stimulated muscles and denervated muscles, we have shown 1) that muscle activity patterns can regulate LCFA transport into skeletal muscle of healthy animals, because LCFA transport is downregulated when muscle activity is inhibited and upregulated when muscle activity is increased. In addition, our studies show 2) that protein-mediated LCFA transport can be regulated by several means, either by increasing total muscle LCFA transport protein content, thereby increasing plasmalemmal LCFA transporters, or by reducing plasmalemmal LCFA transporters while not altering their total muscle content.
Muscle activity is completely abolished with the denervation protocol used in the present investigation. In previous studies, denervation has led to a reduced insulin-stimulated glucose transport due to the repression of GLUT4 (14, 23, 43, 54) and reductions in lactate transport as a result of the repression of MCT1 and MCT4 (6, 40, 55). In marked contrast, we observed that chronic muscle inactivity does not repress the LCFA transporters FAT/CD36 and FABPpm despite the fact that GLUT4 protein was reduced in these muscles (data not shown). Thus denervation does not regulate LCFA transporters in the same manner as has been observed for glucose and MCTs. However, despite the unaltered LCFA transporters in denervated muscle, LCFA transport was markedly reduced. This was associated with concomitant reductions in plasmalemmal FAT/CD36 and FABPpm.
Considerable care needs to be taken when alterations in LCFA transport are associated with changes in total muscle FABPpm content. The reason is that FABPpm is also known as mitochondrial aspartate aminotransferase (mAspAT) (8, 11, 49). mAspAT is present on the inner mitochondrial membrane, where this protein binds to the α-ketoglutarate dehydrogenase complex (17, 50) and catalyzes the following reversible reaction: glutamate + oxaloacetate ↔ aspartate + 2-oxoglutarate (31). However, giant sarcolemmal vesicles do not contain mitochondria; therefore, the reductions in plasmalemmal FABPpm in denervated muscle are not confounded by mitochondrial FABPpm/mAspAT.
Chronic Stimulation Studies
The increase in the total content of FAT/CD36 in chronically stimulated muscle was observed previously in our laboratory (1). However, the increase both in the total content of FABPpm and at sarcolemma in chronically active muscle has not been reported previously. An increase in the total content of FABPpm has been observed after a period of exercise training (29, 51). In previous studies, the total muscle content of other transport proteins, such as MCT1 (6, 41, 42, 55) and the glucose transporter GLUT4 (28, 46, 56), were also increased by chronic low-frequency stimulation. We also observed an increase in GLUT4 in the present studies (data not shown). However, such an upregulation of transport proteins is not a generalized response in this chronic contraction model, because the MCT4 transporter content is not increased (6). The increase in LCFA transporter content in chronically stimulated muscle would seem to account for its increased presence in the plasma membrane, since the relative increases in the plasma membrane transporter FAT/CD36 and the total pool of this transporter were of the same order of magnitude. Nevertheless, the relative increase in plasmalemmal FABPpm was smaller than the relative increase in the total FABPpm content in muscle homogenates. This suggests that there was a substantially larger increase in mitochondrial FABPpm/mAspAT than in plasmalemmal FABPpm.
Correlation Between Muscle Activity and Plasmalemmal LCFA Transporters
A strong correlation was found between the rates of palmitate uptake and the plasma membrane content of FAT/CD36 and FABPpm in giant vesicles prepared from chronically stimulated and denervated hindlimb muscles. Clearly, chronically altered muscle activity regulates the plasmalemmal content of LCFA transporters. Given that both FAT/CD36 and FABPpm were altered in concert in both denervated and chronically stimulated muscles, this might indicate that FAT/CD36 and FABPpm cooperate in a joint fashion to translocate LCFA across the sarcolemma. Although there is some preliminary evidence for this suggestion (38), other data indicate that LCFA transport can be altered when only plasmalemmal FAT/CD36 is increased (33a) or when only FABPpm is increased (12b). These studies (33a and 12b), however, do not rule out that it is still necessary to have both FAT/CD36 and FABPpm present at the sarcolemma to facilitate the uptake of LCFAs. It remains to be established whether FAT/CD36 and FABPpm are indeed two protein components of a single LCFA transport system.
Intracellular Cycling of LCFA Transporters
Despite the belief that LCFAs enter the cell via diffusion (21, 22, 57), there is now considerable molecular information demonstrating that LCFA transport is regulated, in part, by LCFA transport proteins (3, 13, 18, 25–27, 34, 37, 48). Our present studies show clearly that changes in LCFA transport are directly associated with concomitant changes in plasmalemmal FAT/CD36. This supports previous studies from our laboratory, in which we showed that plasmalemmal FAT/CD36 transporters are altered in direct relation to the changes in their total muscle content (36, 48). However, we have also observed previously that LCFA transport into muscle is increased when only plasmalemmal FAT/CD36 is increased, whereas no changes occurred in the total content of this protein in muscle (33a). This can occur because FAT/CD36 can cycle between an intracellular depot and the plasma membrane in skeletal muscle and in the heart (3, 34, 37). In the present study, the plasmalemmal FAT/CD36 was reduced with denervation despite the fact that FAT/CD36 content in muscle was not altered. These observations suggest that FAT/CD36 was resequestered to its intracellular depot(s).
In the face of the unaltered total FABPpm content in the denervated muscle, the reduction in plasmalemmal FABPpm in denervated muscle was unexpected. It had been thought that this protein was present only at the plasma membrane, aside from being identical to mAspAT and being present in mitochondria (8, 27, 49). The present studies, however, indicated that FABPpm might also be present in an intracellular depot, because the plasmalemmal FABPpm was altered, whereas no change in total muscle FABPpm content was observed. We have now been able to confirm that there is an intracellular FABPpm depot in both muscle and heart (12). Thus it would seem that, in denervated muscle, the reduced LCFA uptake was also associated with a resequestering of FABPpm to its intracellular depot.
This strategy of not altering LCFA transport protein content while still altering the plasmalemmal protein content [denervated muscles in the present studies and in obese Zucker rats (33a)] has also been observed with GLUT4. In obesity, total muscle GLUT4 content is not altered, but insulin-induced GLUT4 translocation is impaired (9, 20, 52), and thus plasmalemmal GLUT4 is reduced. Clearly, the cycling of transport proteins between the muscle's surface and intracellular depots is an important mechanism regulating substrate transport rates.
Possible Muscle Activity-Related Mechanisms That Regulate LCFA Transporter Content and Subcellular Distribution
The mechanism for inducing the changes in LCFA transporters in the present study is speculative. With chronic stimulation, the increased metabolic rate is well known to act as a very strong stimulus to rapidly upregulate the muscle content of many proteins (44b). Presumably, this involves activation of the AMP kinase (AMPK)-signaling pathway (44a), although this remains to be determined for specific proteins, including LCFA transporters. Because of the increased metabolic rate in chronically stimulated muscles, more substrate for ATP production is also required; hence, increasing the plasmalemmal content of LCFA transporters can effectively deliver more LCFAs into the muscle, where they can be oxidized. This process of providing more LCFA transport proteins at the sarcolemma may also be due to AMPK activation in chronically stimulated skeletal muscle, because we have recently shown (33b) that FAT/CD36 translocation is induced by AMPK activation in cardiac myocytes.
In contrast to chronic stimulation, the muscles' energetic demands are lowered with denervation. This may involve the downregulation of AMPK, since activating AMPK in denervated muscle with AICAR (5-aminoimidazole-4-carboxamide-1-β-d-ribofuranoside) prevented the decline in GLUT4 in denervated gastrocnemius muscles, although not in denervated soleus muscles (44a). In our studies, the total LCFA transporter content in denervated muscle was not reduced. This may suggest that the reduction in AMPK activation in denervated muscle (44a) is not sufficiently large to affect the LCFA transport protein content. Or, alternatively, AMPK is not involved in regulating LCFA transporter content in denervated muscles.
On the other hand, LCFA provision into the denervated muscles was reduced because there were fewer plasmalemmal LCFA transporters. This failure to maintain plasmalemmal LCFA transporters, as distinct from maintaining the total pool of LCFA transporters, could well be due to the lowered AMPK activity in denervated muscle, since AMPK is known to induce the translocation of LCFA transporters (33b). Thus the reduced AMPK activation in denervated muscle (44a) may be seen as reduction in the stimulus that induces the translocation of LCFA transporters to the plasma membrane. We have previously reported (3) that the translocation of FAT/CD36 to the plasma membrane is regulated by the intensity of muscle contraction (30 min), and therefore, presumably by the extent of AMPK activation. Clearly, an examination of the involvement of AMPK signaling with respect to LCFA transporter content and subcellular localization in skeletal muscle is warranted.
We have shown that alterations in LCFA uptake in chronically stimulated and denervated hindlimb muscles occur via different mechanisms. The present data reveal that, whereas the rates of LCFA transport are associated with concomitant changes in plasmalemmal FAT/CD36 and FABPpm, the mechanisms involved in altering these transport proteins at the sarcolemma are fundamentally different in denervated and chronically stimulated muscles. The increase in LCFA uptake induced by chronic stimulation is attributable to an increased content of FAT/CD36 and FABPpm, which results in an increased sarcolemmal content of these LCFA transporters. In contrast, the reduction in LCFA transport in denervated hindlimb muscles cannot be explained by a repression of the LCFA transporters. Instead, a diminished abundance of the LCFA transporters at the plasma membrane suggests that they were resequestered to their intracellular depots. The present studies indicate that LCFA transport in muscle can be regulated by altering 1) the total muscle content of LCFA transporters and 2) the distribution of these proteins between the plasma membrane and their intracellular depots.
These studies were funded by the Van Walree Fund granted by the Royal Netherlands Academy of Arts and Sciences and by grants from the Netherlands Organization for Scientific Research (ZonMW: 903-39-194), the Canadian Institutes of Health Research, the Natural Sciences and Engineering Research Council of Canada, and the Canada Research Chair program.
J. J. F. P. Luiken is the recipient of a VIDI-Innovation Research Grant from the Netherlands Organization for Scientific Research (NWO-ZonMw Grant 016.036.305).
We acknowledge M. M. A. L. Pelsers for valuable technical assistance.
J. F. C. Glatz is Netherlands Heart Foundation Professor of Cardiac Metabolism.
A. Bonen is Canada Research Chair in Metabolism and Health.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
- Copyright © 2004 by American Physiological Society