Different mechanisms can alter fatty acid transport when muscle contractile activity is chronically altered

Debby P. Y. Koonen,1 Carley R. Benton,2 Yoga Arumugam,3 Narendra N. Tandon,4 Jorge Calles-Escandon,5 Jan F. C. Glatz,1 Joost J. F. P. Luiken,1 and Arend Bonen3

1Department of Molecular Genetics, Cardiovascular Research Institute Maastricht (CARIM), Maastricht University, NL-6200 MD Maastricht, The Netherlands; 2Department of Kinesiology, University of Waterloo, Waterloo, Ontario N2L 3G1; 3Department of Human Biology and Nutritional Sciences, University of Guelph, Guelph, Ontario, N1G 2W1, Canada; 4Thrombosis Research LaboratoryOtsuka America Pharmaceutical Inc., Rockville, Maryland 20850; and 5Section of Endocrinology and Metabolism, Wake Forest University School of Medicine & Baptist Medical Center, Winston-Salem, North Carolina 27156

Submitted 20 November 2003 ; accepted in final form 2 February 2004


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
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; gastrocnemius; denervation; 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.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Materials

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.

Animals

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).

Statistical Analyses

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.


    RESULTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
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).



View larger version (16K):
[in this window]
[in a new window]
 
Fig. 1. Long-chain fatty acid (LCFA) transport into giant sarcolemmal vesicles prepared from 7-day chronically stimulated (STIM) and 7-day denervated (DEN) hindlimb muscle. Vesicles were prepared from pooled fast-twitch oxidative-glycolytic (FOG) and fast-twitch glycolytic (FG) chronically stimulated or denervated muscles. For chronic stimulation treatment, tissues from 3 animals were pooled for each independent experiment; for denervation treatment, tissues from 2 animals were pooled for each independent experiment. Values are means ± SE; n = 5 independent experiments for chronically stimulated treatment; n = 6 independent experiments for denervation treatment. *Significantly different from control (P < 0.05).

 
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.



View larger version (29K):
[in this window]
[in a new window]
 
Fig. 2. Muscle homogenate protein expression of fatty acid translocase (FAT)/CD36 and plasma membrane-associated fatty acid-binding protein (FABPpm) in red and white chronically stimulated (left) or denervated (right) muscle (means ± SE). Muscle fiber type composition in rat hindlimb muscles has been reported previously in our studies (5, 42, 43). Red gastrocnemius (RG, denervation) and red tibialis anterior (RTA, chronic stimulation) are primarily composed of FOG fibers. White gastrocnemius (WG, denervation) and white TA (WTA, chronic stimulation) are primarily composed of FG fibers. Extensor digitorum longus (EDL) muscle (chronic stimulation) is composed of similar proportions of FG and FOG fibers. Soleus muscle (denervation) is primarily composed of slow-twitch oxidative (SO) fibers. Values are means ± SE; n = 5–7 muscles/treatment. Control muscles were set to 100, and experimental muscles were expressed relative to this. *Significantly different from control (P < 0.05).

 
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).



View larger version (23K):
[in this window]
[in a new window]
 
Fig. 3. Plasma membrane FAT/CD36 and FABPpm in giant vesicles derived from chronically stimulated and denervated muscle. Vesicles were prepared from pooled FOG and FG chronically stimulated or denervated muscles. For chronic stimulation treatment, tissues from 3 animals were pooled for each independent experiment, and for denervation treatment, tissues from 2 animals were pooled for each independent experiment. Data are expressed relative to control muscles in each experiment (100%). Values are means ± SE; n = 5 independent experiments for chronically stimulated treatment, and n = 6 independent experiments for denervation treatment. *Significantly different from control (P < 0.05).

 
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).



View larger version (15K):
[in this window]
[in a new window]
 
Fig. 4. Relationship between plasma membrane FAT/CD36 (A) and FABPpm (B) and rates of palmitate transport into giant sarcolemmal vesicles (means ± SE). Control data in the 2 experiments did not differ and were therefore pooled. Transport data and plasma membrane data were not always obtained on the same animals; therefore, results from each group of animals were pooled. Thereafter, linear regression analyses were performed by using these mean data. Data are derived from Figs. 1 and 3.

 

    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
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.

Denervation Studies

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 {alpha}-ketoglutarate dehydrogenase complex (17, 50) and catalyzes the following reversible reaction: glutamate + oxaloacetate {leftrightarrow} 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, 2527, 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-{beta}-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.

Summary

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.


    GRANTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
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).


    ACKNOWLEDGMENTS
 
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.


    FOOTNOTES
 

Address for reprint requests and other correspondence: A. Bonen, Dept. of Human Biology and Nutritional Sciences, Univ. of Guelph, Guelph, Ontario, N1G 2W1, Canada (E-mail: abonen{at}uoguelph.ca).

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.


    REFERENCES
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Bonen A, Dyck DJ, Ibrahimi A, and Abumrad NA. Muscle contractile activity increases fatty acid metabolism and transport and FAT/CD36. Am J Physiol Endocrinol Metab 276: E642–E649, 1999.[Abstract/Free Full Text]
  2. Bonen A, Dyck DJ, and Luiken JJ. Skeletal muscle fatty acid transport and transporters. Adv Exp Med Biol 441: 193–205, 1998.[ISI][Medline]
  3. Bonen A, Luiken JJ, Arumugam Y, Glatz JF, and Tandon NN. Acute regulation of fatty acid uptake involves the cellular redistribution of fatty acid translocase. J Biol Chem 275: 14501–14508, 2000.[Abstract/Free Full Text]
  4. Bonen A, Luiken JJ, Liu S, Dyck DJ, Kiens B, Kristiansen S, Turcotte LP, Van Der Vusse GJ, and Glatz JF. Palmitate transport and fatty acid transporters in red and white muscles. Am J Physiol Endocrinol Metab 275: E471–E478, 1998.[Abstract/Free Full Text]
  5. Bonen A, Miskovic D, Tonouchi M, Lemieux K, Wilson MC, Marette A, and Halestrap AP. Abundance and subcellular distribution of MCT1 and MCT4 in heart and fast-twitch skeletal muscles. Am J Physiol Endocrinol Metab 278: E1067–E1077, 2000.[Abstract/Free Full Text]
  6. Bonen A, Tonouchi M, Miskovic D, Heddle C, Heikkila JJ, and Halestrap AP. Isoform-specific regulation of the lactate transporters MCT1 and MCT4 by contractile activity. Am J Physiol Endocrinol Metab 279: E1131–E1138, 2000.[Abstract/Free Full Text]
  7. Booth FW and Thomason DB. Molecular and cellular adaptation of muscle in response to exercise: perspectives of various models. Physiol Rev 71: 541–585, 1991.[Free Full Text]
  8. Bradbury MW and Berk PD. Mitochondrial aspartate aminotransferase: direction of a single protein with two distinct functions to two subcellular sites does not require alternative splicing of the mRNA. Biochem J 345: 423–427, 2000.[CrossRef][ISI][Medline]
  9. Brozinick JT Jr, Etgen GJ Jr, Yaspelkis BB III, Kang HY, and Ivy JL. Effects of exercise training on muscle GLUT-4 protein content and translocation in obese Zucker rats. Am J Physiol Endocrinol Metab 265: E419–E427, 1993.[Abstract/Free Full Text]
  10. Calles-Escandon J, Sweet L, Ljungqvist O, and Hirshman MF. The membrane-associated 40 KD fatty acid binding protein (Berk's protein), a putative fatty acid transporter is present in human skeletal muscle. Life Sci 58: 19–28, 1996.[CrossRef][ISI][Medline]
  11. Cechetto JD, Sadacharan SK, Berk PD, and Gupta RS. Immunogold localization of mitochondrial aspartate aminotransferase in mitochondria and on the cell surface in normal rat tissues. Histol Histopathol 17: 353–364, 2002.[ISI][Medline]
  12. Chabowski A and Bonen A. Regulation of fatty acid transport and transporters by insulin and AICAR (Abstract). Can J Appl Physiol 28: S32, 2003.
  13. Clarke DC, Miskovic D, Han X-X, Calles-Escandon J, Glatz JFC, Luiken JJFP, Heikkila JJ, and Bonen A. Overexpression of membrane associated fatty acid binding protein (FABPpm) in vivo increases fatty acid sarcolemmal transport and metabolism. 10.1152/physiolgenomics.00190.2003. In press.
  14. Coburn CT, Knapp FF Jr, Febbraio M, Beets AL, Silverstein RL, and Abumrad NA. Defective uptake and utilization of long chain fatty acids in muscle and adipose tissues of CD36 knockout mice. J Biol Chem 275: 32523–32529, 2000.[Abstract/Free Full Text]
  15. Coderre L, Monfar MM, Chen KS, Heydrick SJ, Kurowski TG, Ruderman NB, and Pilch PF. Alteration in the expression of GLUT-1 and GLUT-4 protein and messenger RNA levels in denervated rat muscles. Endocrinology 131: 1821–1825, 1992.[Abstract]
  16. Coe NR, Smith AJ, Frohnert BI, Watkins PA, and Bernlohr DA. The fatty acid transport protein (FATP1) is a very long chain acyl-CoA synthetase. J Biol Chem 274: 36300–36304, 1999.[Abstract/Free Full Text]
  17. Dyck DJ and Bonen A. Muscle contraction increases palmitate esterification and oxidation and triacylglycerol oxidation. Am J Physiol Endocrinol Metab 275: E888–E896, 1998.[Abstract/Free Full Text]
  18. Fahien LA, Davi JW, and Laboy J. Interactions between pyruvate carboxylase and other mitochondrial enzymes. J Biol Chem 268: 17935–17942, 1993.[Abstract/Free Full Text]
  19. Febbraio M, Abumrad NA, Hajjar DP, Sharma K, Cheng W, Pearce SF, and Silverstein RL. A null mutation in murine CD36 reveals an important role in fatty acid and lipoprotein metabolism. J Biol Chem 274: 19055–19062, 1999.[Abstract/Free Full Text]
  20. Glatz JF and van der Vusse GJ. Cellular fatty acid-binding proteins: their function and physiological significance. Prog Lipid Res 35: 243–282, 1996.[CrossRef][ISI][Medline]
  21. Goodyear LJ and Kahn BB. Exercise, glucose transport, and insulin sensitivity. Ann Rev Med 49: 235–261, 1998.[CrossRef][ISI][Medline]
  22. Hamilton JA. Transport of fatty acids across membranes by the diffusion mechanism. Prostaglandins Leukot Essent Fatty Acids 60: 291–297, 1999.[CrossRef][ISI][Medline]
  23. Hamilton JA, Guo W, and Kamp F. Mechanism of cellular uptake of long-chain fatty acids: Do we need cellular proteins? Mol Cell Biochem 239: 17–23, 2002.[CrossRef][ISI][Medline]
  24. Henriksen EJ, Rodnick KJ, Mondon CE, James DE, and Holloszy JO. Effect of denervation or unweighting on GLUT-4 protein in rat soleus muscle. J Appl Physiol 70: 2322–2327, 1991.[Abstract/Free Full Text]
  25. Hui TY and Bernlohr DA. Fatty acid transporters in animal cells. Front Biosci 2: 221–231, 1997.
  26. Ibrahimi A, Bonen A, Blinn WD, Hajri T, Li X, Zhong K, Cameron R, and Abumrad NA. Muscle-specific overexpression of FAT/CD36 enhances fatty acid oxidation by contracting muscle, reduces plasma triglycerides and fatty acids, and increases plasma glucose and insulin. J Biol Chem 274: 26761–26766, 1999.[Abstract/Free Full Text]
  27. Ibrahimi A, Sfeir Z, Magharaie H, Amri EZ, Grimaldi P, and Abumrad NA. Expression of the CD36 homolog (FAT) in fibroblast cells: effects on fatty acid transport. Proc Natl Acad Sci USA 93: 2646–2651, 1996.[Abstract/Free Full Text]
  28. Isola LM, Zhou SL, Kiang CL, Stump DD, Bradbury MW, and Berk PD. 3T3 fibroblasts transfected with a cDNA for mitochondrial aspartate aminotransferase express plasma membrane fatty acid-binding protein and saturable fatty acid uptake. Proc Natl Acad Sci USA 92: 9866–9870, 1995.[Abstract]
  29. Johannsson E, McCullagh KJ, Han XX, Fernando PK, Jensen J, Dahl HA, and Bonen A. Effect of overexpressing GLUT-1 and GLUT-4 on insulin- and contraction-stimulated glucose transport in muscle. Am J Physiol Endocrinol Metab 271: E547–E555, 1996.[Abstract/Free Full Text]
  30. Kiens B, Kristiansen S, Jensen P, Richter EA, and Turcotte LP. Membrane associated fatty acid binding protein (FABPpm) in human skeletal muscle is increased by endurance training. Biochem Biophys Res Commun 231: 463–465, 1997.[CrossRef][ISI][Medline]
  31. Koonen DP, Coumans WA, Arumugam Y, Bonen A, Glatz JF, and Luiken JJ. Giant membrane vesicles as a model to study cellular substrate uptake dissected from metabolism. Mol Cell Biochem 239: 121–130, 2002.[CrossRef][ISI][Medline]
  32. Lehninger AL, Nelson DL, and Cox MM. Principles of Biochemistry. New York: Worth, 1993.
  33. Luiken JJ, Arumugam Y, Bell RC, Calles-Escandon J, Tandon NN, Glatz JF, and Bonen A. Changes in fatty acid transport and transporters are related to the severity of insulin deficiency. Am J Physiol Endocrinol Metab 283: E612–E621, 2002.[Abstract/Free Full Text]
  34. Luiken JJ, Arumugam Y, Dyck DJ, Bell RC, Pelsers MM, Turcotte LP, Tandon NN, Glatz JF, and Bonen A. Increased rates of fatty acid uptake and plasmalemmal fatty acid transporters in obese Zucker rats. J Biol Chem 276: 40567–40573, 2001.[Abstract/Free Full Text]
  35. Luiken JJFP, Coort SML, Willems J, Coumans WA, Bonen A, van der Vusse GJ, and Glatz JFC. Contraction-induced fatty acid translocase/CD36 translocation in rat cardiac myocytes is mediated through AMP-activated protein kinase signaling. Diabetes 52: 1627–1634, 2003.[Abstract/Free Full Text]
  36. Luiken JJ, Dyck DJ, Han XX, Tandon NN, Arumugam Y, Glatz JF, and Bonen A. Insulin induces the translocation of the fatty acid transporter FAT/CD36 to the plasma membrane. Am J Physiol Endocrinol Metab 282: E491–E495, 2002.[Abstract/Free Full Text]
  37. Luiken JJ, Glatz JF, and Bonen A. Fatty acid transport proteins facilitate fatty acid uptake in skeletal muscle. Can J Appl Physiol 25: 333–352, 2000.[ISI][Medline]
  38. Luiken JJ, Koonen DP, Coumans WA, Pelsers MM, Binas B, Bonen A, and Glatz JF. Long-chain fatty acid uptake by skeletal muscle is impaired in homozygous, but not heterozygous, heart-type-FABP null mice. Lipids 38: 491–496, 2003.[ISI][Medline]
  39. Luiken JJ, Koonen DP, Willems J, Zorzano A, Becker C, Fischer Y, Tandon NN, Van Der Vusse GJ, Bonen A, and Glatz JF. Insulin stimulates long-chain fatty acid utilization by rat cardiac myocytes through cellular redistribution of FAT/CD36. Diabetes 51: 3113–3119, 2002.[Abstract/Free Full Text]
  40. Luiken JJ, Turcotte LP, and Bonen A. Protein-mediated palmitate uptake and expression of fatty acid transport proteins in heart giant vesicles. J Lipid Res 40: 1007–1016, 1999.[Abstract/Free Full Text]
  41. Matsuno K, Diaz-Ricart M, Montgomery RR, Aster RH, Jamieson GA, and Tandon NN. Inhibition of platelet adhesion to collagen by monoclonal anti-CD36 antibodies. Br J Haematol 92: 960–967, 1996.[CrossRef][ISI][Medline]
  42. McCullagh KJ and Bonen A. Reduced lactate transport in denervated rat skeletal muscle. Am J Physiol Regul Integr Comp Physiol 268: R884–R888, 1995.[Abstract/Free Full Text]
  43. McCullagh KJ, Juel C, O'Brien M, and Bonen A. Chronic muscle stimulation increases lactate transport in rat skeletal muscle. Mol Cell Biochem 156: 51–57, 1996.[ISI][Medline]
  44. McCullagh KJ, Poole RC, Halestrap AP, Tipton KF, O'Brien M, and Bonen A. Chronic electrical stimulation increases MCT1 and lactate uptake in red and white skeletal muscle. Am J Physiol Endocrinol Metab 273: E239–E246, 1997.[Abstract/Free Full Text]
  45. Megeney LA, Neufer PD, Dohm GL, Tan MH, Blewett CA, Elder GC, and Bonen A. Effects of muscle activity and fiber composition on glucose transport and GLUT-4. Am J Physiol Endocrinol Metab 264: E583–E593, 1993.[Abstract/Free Full Text]
  46. Paulsen SR, Rubink DS, and Winder WW. AMP-activated protein kinase activation prevents denervation-induced decline in gastrocnemius GLUT-4. J Appl Physiol 91: 2102–2108, 2001.[Abstract/Free Full Text]
  47. Pette D. J. B. Wolffe Memorial Lecture. Activity-induced fast to slow transitions in mammalian muscle. Med Sci Sports Exerc 16: 517–528, 1984.[ISI][Medline]
  48. Richards MR, Listenberger LL, Kelly AA, Lewis SE, Ory D, and Schaffer JE. Oligomerization of the murine fatty acid transport protein 1. J Biol Chem 278: 10477–10483, 2003.[Abstract/Free Full Text]
  49. Roy D, Johannsson E, Bonen A, and Marette A. Electrical stimulation induces fiber type-specific translocation of GLUT-4 to T tubules in skeletal muscle. Am J Physiol Endocrinol Metab 273: E688–E694, 1997.[Abstract/Free Full Text]
  50. Schaffer JE. Fatty acid transport: the roads taken. Am J Physiol Endocrinol Metab 282: E239–E246, 2002.[Abstract/Free Full Text]
  51. Steinberg GR, Dyck DJ, Calles-Escandon J, Tandon NN, Luiken JJ, Glatz JF, and Bonen A. Chronic leptin administration decreases fatty acid uptake and fatty acid transporters in rat skeletal muscle. J Biol Chem 277: 8854–8860, 2002.[Abstract/Free Full Text]
  52. Stump DD, Zhou SL, and Berk PD. Comparison of plasma membrane FABP and mitochondrial isoform of aspartate aminotransferase from rat liver. Am J Physiol Gastrointest Liver Physiol 265: G894–G902, 1993.[Abstract/Free Full Text]
  53. Teller JK, Fahien LA, and Valdivia E. Interactions among mitochondrial aspartate aminotransferase, malate dehydrogenase, and the inner mitochondrial membrane from the heart, hepatoma, and liver. J Biol Chem 265: 19486–19494, 1990.[Abstract/Free Full Text]
  54. Turcotte LP, Swenberger JR, Tucker MZ, and Yee AJ. Training-induced elevation in FABP(PM) is associated with increased palmitate use in contracting muscle. J Appl Physiol 87: 285–293, 1999.[Abstract/Free Full Text]
  55. Uphues I, Chern Y, and Eckel J. Insulin-dependent translocation of the small GTP-binding protein rab3C in cardiac muscle: studies on insulin-resistant Zucker rats. FEBS Lett 377: 109–112, 1995.[CrossRef][ISI][Medline]
  56. Watkins PA, Lu JF, Steinberg SJ, Gould SJ, Smith KD, and Braiterman LT. Disruption of the Saccharomyces cerevisiae FAT1 gene decreases very long-chain fatty acyl-CoA synthetase activity and elevates intracellular very long-chain fatty acid concentrations. J Biol Chem 273: 18210–18219, 1998.[Abstract/Free Full Text]
  57. Wilkes JJ and Bonen A. Reduced insulin-stimulated glucose transport in denervated muscle is associated with impaired Akt-{alpha} activation. Am J Physiol Endocrinol Metab 279: E912–E919, 2000.[Abstract/Free Full Text]
  58. Wilson MC, Jackson VN, Heddle C, Price NT, Pilegaard H, Juel C, Bonen A, Montgomery I, Hutter OF, and Halestrap AP. Lactic acid efflux from white skeletal muscle is catalyzed by the monocarboxylate transporter isoform MCT3. J Biol Chem 273: 15920–15926, 1998.[Abstract/Free Full Text]
  59. Yaspelkis BB III, Castle AL, Farrar RP, and Ivy JL. Contraction-induced intracellular signals and their relationship to muscle GLUT-4 concentration. Am J Physiol Endocrinol Metab 272: E118–E125, 1997.[Abstract/Free Full Text]
  60. Zakim D. Fatty acids enter cells by simple diffusion. Proc Soc Exp Biol Med 212: 5–14, 1996.[Medline]
  61. Zou Z, DiRusso CC, Ctrnacta V, and Black PN. Fatty acid transport in Saccharomyces cerevisiae. Directed mutagenesis of FAT1 distinguishes the biochemical activities associated with Fatp1. J Biol Chem 277: 31062–31071, 2002.[Abstract/Free Full Text]