Peroxisome Proliferator-Activated Receptor-α Regulates Fatty Acid Utilization in Primary Human Skeletal Muscle Cells

BSA (essentially fatty acid–free), carnitine, sodium oleate, and oil red O stain were from Sigma (St. Louis, MO). Fetal bovine serum (FBS) and Hank’s balanced salt solution were from Life Technologies (Gaithersburg, MD). Heat-inactivated horse serum was from Hyclone (Logan, UT). Growth media (GM) and differentiation media (DFM) consisted of Dulbecco’s modified Eagles medium (DMEM) from Life Technologies supplemented with human skeletal muscle SingleQuots from BioWhittaker (Walkersville, MD). Biocoat tissue culture plates were from Becton Dickinson (Franklin Lakes, NJ). PCR reagents were from PE Applied Biosystems (Foster City, CA). GW7647 was obtained from Dr. Peter Brown at GlaxoSmithKline.

Protocols were approved by the institutional review board at East Carolina University. Muscle samples weighing 50–200 mg, which were obtained from vastus lateralis by needle biopsy or from rectus abdominis muscle that was excised during surgical procedure, were immediately transferred to ice-cold DMEM and cleaned free of adipose and connective tissues. Satellite cells were isolated by trypsin digestion (13), preplated 1–3 h in 3.0 ml GM on an uncoated T-25 tissue culture flask to remove fibroblasts, and then transferred to a type I collagen–coated T25 flask for attachment. Cells were cultured at 37°C in a humidified atmosphere of 5% CO₂ in GM supplemented with 10% FBS, 0.5 mg/ml BSA, 0.5 mg/ml fetuin, 20 ng/ml human epidermal growth factor, 0.39 μg/ml dexamethasone, and 50 μg/ml gentamicin/amphotericin B. After reaching ∼70% confluence, myoblasts were subcultured onto 6-, 12-, and 24-well type I collagen–coated plates at densities of 100, 50, and 20 × 10³ cells per well, respectively. When cells reached 80–90% confluence, differentiation was induced by changing to low-serum DFM consisting of 2% heat-inactivated horse-serum, 0.5 mg/ml BSA, 0.5 mg/ml fetuin, and 50 μg/ml gentamicin/amphotericin B. Media was changed every 2–3 days, and the PPAR-α–selective compound, GW7647, or DMSO vehicle (0.1% vol/vol) was added to developing myotubes on differentiation days 1–6 or to mature myotubes on days 6–7. Cells were harvested in 1.0 ml Trizol reagent for RNA extraction or 500 μl NP-40 lysis buffer for Western analyses.

Protein (30–50 μg) prepared from total cell lysates was separated by 10% SDS-PAGE, transferred to PVDF membranes (Biorad, Hercules, CA), and then incubated with antibodies diluted in 5% milk in Tris-buffered saline with 0.1% Tween. Proteins were visualized by horseradish peroxidase–conjugated goat anti–rabbit or anti–mouse immunoglobulin G from Santa Cruz Biotechnology (Santa Cruz, CA) using a chemiluminescence Western-blotting detection kit from Pierce (Rockford, IL). Myosin and Myo-D polyclonal antibodies were from Santa Cruz, antibody against medium-chain acyl-CoA dehydrogenase (MCAD) was a gift from Dan Kelly (Washington University, St. Louis, MO), and monoclonal anti–hPPAR-α was synthesized as previously described (11).

Myocytes maintained in differentiation medium for 0 days (myoblasts) or 3–9 days (developing myotubes) were incubated at 37°C in sealed 12- or 24-well plates containing 500 or 750 μl serum-free DFM plus 12.5 mmol/l HEPES, 0.2% BSA, 1.0 mmol/l carnitine, 100 μmol/l sodium oleate, 50 μg/ml gentamicin, and 1.0 μCi/ml [¹⁴C]oleate (NEN, Boston, MA). After 3 h, the incubation media was transferred to new dishes and assayed for labeled oxidation products (CO₂ and acid-soluble metabolites [ASMs]) (14). The cells were placed on ice, washed twice with PBS, scraped into a 1.5-ml eppendorf tube in two additions of 0.30 ml 0.05% SDS lysis buffer, and then stored at −80°C. Cell lysates were later assayed for protein, and then total cell lipids were extracted (15). Aliquots of the lipid extracts were spotted on 0.25-mm silica gel G plates from Whatman (Maidstone, Kent, U.K.) and chromatographed with hexane:diethyl ether:acetic acid (80:20:1 vol/vol) in parallel with authentic standards. [¹⁴C]oleate-labeled lipid products were quantified using a BioScan Image 200 System (Bioscan, Washington, DC). All assays were performed in triplicate, and data are presented as the means ± SE of results from 5 to 10 subjects.

Total RNA was prepared using TriZol reagent according the manufacturer’s protocol (Life Technology), treated with DNase I (Ambion, Austin, TX), and quantified using the RiboGreen RNA quantitation kit (Molecular Probes, Eugene, OR). Real-time quantitative PCR (RTQ-PCR) was performed using an ABI PRISM 7700 Sequence Detection System instrument and software (PE Applied Biosystems, Foster City, CA), and primer/probe sets (Table 1) were designed using the manufacturer’s software and the sequences available in GenBank. Expression levels of selected genes were compared between myotubes from the same subject, treated with either vehicle or 1.0 μmol/l GW7647. RNA samples were normalized for comparison by determining 18S rRNA levels by RTQ-PCR. Expression levels were quantified (arbitrary units) by generating a seven-point serial standard curve, as previously described (16). Results are expressed as the fold change, as determined by the ratio of calculated units of RNA in GW7647-treated samples to those in vehicle controls, and are presented as the means ± SE from four subjects per group. For comparison, relative quantitation was also calculated by using the 2^ΔC_T formula, in which ΔC_T equals the difference between C_T (cycle threshold) values for control and treated cells. Results using this formula (not shown) were essentially identical to those generated using the standard curves.

Statistical analyses were performed using JMP Statistical Software (SAS, Cary, NC). Differences between vehicle- and GW7647-treated cells were analyzed by Student’s t test for paired data, and one- or two-way ANOVAs were performed using a standard least squares model to test both the main and interaction effects of GW7647 and etomoxir (where appropriate) on parameters of lipid metabolism.

Preconfluent myoblasts exhibit a mononucleated, spindle-shaped morphology (Fig. 1A). Confluent myoblasts withdraw from the cell cycle, align with one another, and then attach, fuse, and differentiate into multinucleated myotubes (Fig. 1B), a process that is facilitated by switching to a low-serum DFM. Similar to other reports (13), we found that >80% of HSKMCs exhibited elongated, multinucleated myotube morphology within 6–8 days after switching to DFM, indicating low levels of fibroblast contamination in the initial cultures. Although PPAR-α is expressed abundantly in human skeletal muscle (11), it is absent from some stable muscle cell lines (17); thus, we first evaluated PPAR-α protein expression in undifferentiated myoblasts and developing myotubes (Fig. 2A). Expression of PPAR-α protein was low in myoblasts, but it increased markedly by day 3 in DFM and was then maintained throughout myotube development at levels that were similar to those in intact human skeletal muscle (Fig. 2B). Differentiation-dependent expression of PPAR-α coincided with induction of the muscle-specific proteins myosin and Myo-D and the fatty acid oxidative enzyme MCAD. Myocyte differentiation and concomitant induction of PPAR-α were also associated with increased fatty acid oxidative capacity (Fig. 2C).

GW7647 is a novel PPAR-α ligand with an EC₅₀ value of 6 nmol/l, as determined in PPARx/gal4 chimeric transfection assays (12). GW7647 exhibits 1,000-fold selectivity for PPAR-α over PPAR-γ and -δ and is therefore more potent and selective than many commonly used PPAR-α–targeted compounds. To evaluate the role of PPAR-α in regulating muscle lipid metabolism, we treated day 6 myotubes for 48 h with 0–1,000 nmol/l GW7647, and then day 8 myocytes were incubated for 3 h with 100 μmol/l [¹⁴C]oleate. Fatty acid oxidation, measured as the sum of [¹⁴C]oleate oxidized to CO₂ (complete oxidation) plus ASM (incomplete oxidation), was 1.52 ± 0.14 nmol · mg⁻¹ · h⁻¹ in vehicle-treated myotubes. GW7647 increased myotube oleate oxidation to CO₂ and ASMs in a dose-dependent manner (P < 0.001) (Figs. 3A and B). Myotubes responded maximally at 1.0 μmol/l GW7647 by increasing oleate oxidation 2.7-fold to 4.01 ± 0.32 nmol · mg⁻¹ · h⁻¹. This response was similar in cells from different muscle sources (vastus lateralis versus rectus abdominus); therefore, results were pooled together.

In PPAR-α–null mice, pharmacological inhibition of fatty acid oxidation results in marked accumulation of triacylglycerol (TAG) in liver and heart (18). In contrast, when wild-type mice are treated with the same inhibitor, they exhibit only minor increases in tissue TAG (18). These findings suggest not only that PPAR-α is required for adaptive adjustments in TAG homeostasis, but also that TAG regulation occurs via mechanisms that are independent of lipid catabolism. To determine whether PPAR-α might play a similar role in human muscle, we evaluated the effects of GW7647 on [¹⁴C]oleate esterification into glycerolipids. Basal rates of oleate incorporation into TAG and phospholipids were 12.95 ± 1.26 and 8.25 ± 0.76 nmol · mg⁻¹ · h⁻¹, respectively, and only small amounts (<10% of the total) were incorporated into diacylglycerol and other minor lipid species (not shown). Opposite to its effect on fatty acid oxidation, GW7647 decreased oleate esterification into TAG, up to 45% (P < 0.001), without affecting labeling of phospholipids (Figs. 3C and D).

To quantify the partitioning of fatty acid between opposing metabolic pathways, we divided the rate (nmol · mg⁻¹ · h⁻¹) of oleate esterified into TAG by the rate of oleate oxidized, thereby providing an index of fatty acid utilization. Because GW7647 regulated oxidation and esterification in opposite directions, the partitioning index decreased 75%, from a TAG-to-oxidation ratio of 8:1 to 2:1 (Fig. 3E). The impact of this marked adjustment in fatty acid partitioning on myotube lipid homeostasis was then examined histologically, by oil red O staining of neutral lipids (Fig. 4). In vehicle-treated myocytes exposed to an overnight fatty acid load, the intensity of the stain increased as a function of increasing oleate concentration (Fig. 4A). In comparison, 48-h pretreatment with GW7647 attenuated fatty acid–induced accumulation of neutral lipids. In Fig. 4B, higher magnification of stained cells confirmed that neutral lipids had accumulated in mature myotubes and indicated that PPAR-α activation decreased the number cells exhibiting large lipid droplets.

Our observation that GW7647 specifically reduced TAG, but not phospholipid, suggested that this was not a mass action effect caused by changes in the myocellular [¹⁴C]oleate pool. To determine whether the TAG-depleting effects of GW7647 depended on reciprocal changes in oleate oxidation, we repeated previous experiments but performed the 3-h [¹⁴C]oleate incubation in either the presence or absence of etomoxir, a potent inhibitor of fatty acid oxidation. In the absence of etomoxir, 1.0 μmol/l GW37647 increased oleate oxidation approximately threefold, recapitulating our earlier result. As expected, etomoxir inhibited oleate oxidation to negligible levels in both groups (Fig. 5A). During short-term exposures (3 h), blocking oxidation with etomoxir did not result in a compensatory increase in [¹⁴C]oleate esterification (Fig. 5B). Moreover, [¹⁴C]oleate incorporation into myocyte TAG was lower in GW7647-treated cells, regardless of whether etomoxir was present. These data are consistent with lipid studies in PPAR-α–null mice (18) and indicate that the TAG-depleting effects of PPAR-α activation do not require opposing changes in fatty acid oxidation.

In these experiments, vehicle or GW7647 was administered to developing myotubes (days 1–6) or to mature myotubes for 48 h (days 6–8). Using RTQ-PCR, we evaluated changes in steady-state mRNA expression of several candidate target genes that are known or predicted to regulate muscle energy utilization. Pyruvate dehydrogenase kinase (PDHK), which phosphorylates and inactivates the pyruvate dehydrogenase (PDH) complex, is thought to facilitate preferential oxidation of lipid over glucose substrates. Remarkably, GW7647 increased expression of PDHK4 12- and 45-fold in cells that were treated for 48 h and 6 days, respectively (Table 2). GW7647 also induced highly consistent but less robust increases (1.7- to 2.4-fold) in mRNA expression of malonyl-CoA decarboxylase (MCD) and the muscle isoform of carnitine palmityltransferase 1 (CPT1), enzymes that promote uptake of fatty acid into the mitochondria for β-oxidation. Surprisingly, mRNA abundance of PPAR coactivator 1 was decreased 15% (P < 0.05) in mature myotubes that were treated with GW7647 for 48 h; however, this effect was reversed when GW7647 was added to developing myotubes for 6 days. Notably, we found that mRNA expression of the peroxisomal marker enzyme acyl-CoA oxidase (ACO) was unaffected by treatment with GW7647. PPAR-α mRNA abundance was similar in vehicle and GW7647-treated cells; thus, hPPAR-α does not appear to regulate its own expression.

Data from our metabolic studies suggested that GW7647 might repress genes that promote myotube TAG biosynthesis. To address this result, we examined expression of genes involved in TAG synthesis, including sterol regulatory element binding protein 1 (SREBP1), mitochondrial glycerol-3-phosphate acyltransferase (GPAT), and diacylglycerol acyltransferase (DGAT). Contrary to our expectations, mRNA expression of GPAT was similar between control and GW7647-treated cells, and DGAT mRNA abundance was low or undetectable (C_T values >35) in HSKMCs from six of eight subjects, regardless of the treatment (Table 2). SREBP1 actually showed a tendency to increase in response to GW7647, but this effect was statistically significant (P < 0.05) only when results from both the 48-h and 6-day treatments were pooled. Collectively, these data suggest the TAG-depleting effects of GW7647 were not caused by direct repression of glycerolipid biosynthetic enzymes.

In this study, we used a primary HSKMC system to investigate the role of hPPAR-α in regulating muscle fuel metabolism. In contrast to previous animal studies that examined the effects of PPAR-α activators in vivo, the use of primary HSKMCs allowed us to discern direct, muscle-specific effects of hPPAR-α activation. To our knowledge, this is the first report that characterizes PPAR-α expression and its metabolic target genes in developing human myotubes. Our results indicate that hPPAR-α is robustly induced upon myocyte differentiation and plays an important role in regulating lipid utilization in human muscle.

The therapeutic utility of PPAR-α activators in treating dyslipidemia has been well established, but their mechanisms of action, particularly in humans, are still unclear. We found that treating HSKMCs with the highly selective PPAR-α agonist GW7647 increased fatty acid oxidation by approximately threefold, suggesting that the hypolipidemic actions of PPAR-α activators are at least partly mediated by increased muscle clearance and utilization of circulating lipids. In addition to stimulating β-oxidation, GW7647 also decreased fatty acid esterification into myocyte TAG. Similarly, animal studies have shown that PPAR-α agonists ameliorate diet-induced increases in muscle TAG content (5,19), but it was unclear whether this effect was due to primary targeting of muscle gene regulation per se or was secondary to the systemic lipid-lowering actions of the drug. Our results are the first to show that PPAR-α−selective compounds regulate TAG content in human myocytes directly, a finding that has considerable clinical relevance (discussed below).

In the present study, we also examined several candidate target genes that are involved in regulating muscle fatty acid oxidation. Most remarkable was our finding that treating developing myotubes with GW7647 induced a 45-fold increase in PDHK4 mRNA expression. This result is consistent with previous studies showing that PDHK4 mRNA levels respond more robustly to changes in muscle fatty acid flux than other PPAR-α target genes (9,16,20,21). Together with these reports, our results indicate that the PDHK4 gene is highly sensitive to both endogenous and pharmacological PPAR-α ligands. Whether this regulatory property of the gene might be related to distinguishing features of the PPAR binding element within the PDHK4 promoter remains to be determined. PDHK4 functions by phosphorylating and inactivating PDH, which is a multienzyme complex that catalyzes the oxidation of pyruvate to acetyl-CoA (22). Because this is an irreversible reaction that prevents conversion of acetyl-CoA back to glucose, inactivation of the PDH complex facilitates glucose sparing during states of energy depletion. There are at least four PDHK isoenzymes that exhibit distinct tissue distributions and kinetic properties (23). Skeletal muscle expresses PDHK2, a lower specific activity enzyme that is ubiquitously expressed in the fed state, and PDHK4, a higher specific activity enzyme that is less sensitive to inhibition by pyruvate and is robustly induced by increased fatty acid flux (21,24). Our finding that GW7647 selectively upregulated PDHK4 without affecting expression of PDHK2 is consistent with the physiological regulation of these genes in vivo. Increased expression and activity of PDHK4, which occurs in response to exercise (20), starvation (21), and a low-carbohydrate diet (25), is thought to promote fatty acid oxidation and spare pyruvate for nonoxidative, anaplerotic entry into the tricarboxylic acid (TCA) cycle. Conversely, refeeding (21) and insulin treatments (9) decrease PDHK4 expression, which favors preferential oxidation of carbohydrates (26). These studies indicate that the PDH reaction is positioned at a key metabolic branch point. Furthermore, our results, together with these earlier studies, support the hypothesis that PDHK isoform switching imparts a mechanism by which the muscle can rapidly adjust the source of substrate that supplies acetyl-CoA to the TCA cycle (21).

RTQ-PCR analyses also showed that GW7647 increased expression of muscle CPT1 and MCD approximately twofold. The magnitude of these responses is highly consistent with the in vivo effects of pharmacological (10) and physiological (20) activation of PPAR-α. CPT1 catalyzes the initial and rate-limiting step in the transport of fatty acid into mitochondria, whereas MCD disposes of the potent CPT1 inhibitor malonyl-CoA. Thus, activation of these genes increases fatty acid catabolism by promoting their entry into the mitochondria. In contrast to the effects of PPAR-α activation in mouse liver (3) and heart (18), but consistent with reports in humans (6), treating HSKMCs with GW7647 did not induce mRNA expression of the peroxisomal marker enzyme ACO. Although this result suggests that GW7647 might preferentially upregulate mitochondrial over peroxisomal β-oxidation, it does not rule out the possibility that other extramitochondrial pathways might participate in PPAR-α–mediated regulation of cellular lipid balance.

Our finding that GW7647 reduced HSKMC esterification into TAG, even when fatty acid oxidation was inhibited by etomoxir, suggests that the TAG-depleting effects of the compound do not depend on reciprocal increases in lipid catabolism. However, we were unable to explain this observation at the molecular level. GPAT and DGAT catalyze the first and final committed steps, respectively, in the pathway of TAG biosynthesis, and SREBP1 is a transcription factor that coordinately stimulates expression of several lipogenic enzymes (27). Expression levels of these candidate genes were unchanged by the PPAR-α agonist. Thus, we found no evidence that PPAR-α represses expression of TAG biosynthetic enzymes. Alternatively, PPAR-α activation might decrease TAG content by reducing the supply of lipogenic precursors, stimulating hydrolysis, or perhaps by stimulating lipid export via lipoprotein particles, a pathway known to operate primarily in liver but that has also been described in heart (28). These possibilities warrant further investigation.

Recently, there has been heightened interest in the lipid oversupply hypothesis that links increased muscle lipid content with the development of insulin resistance (29). Several rodent studies have shown that insulin sensitivity indexes correlate inversely with changes in muscle lipid content (5,30). Similarly, in humans, [¹H-¹³C] nuclear magnetic resonance studies have demonstrated that intramyocellular TAG content correlates inversely with insulin resistance, and that multiple regression analyses select muscle TAG as the strongest predictor of insulin resistance, independent of BMI, adiposity, and waist-to-hip ratio (31,32). These data suggest that muscle lipid dysregulation, which is marked by increased muscle TAG content, is causally related to insulin resistance. The underlying factors contributing to muscle lipid accumulation are still obscure but may be related to reduced oxidative capacity (rev. in 33). We and others have reported that fatty acid oxidation rates (14) and fatty acid oxidative enzyme activities (14,33) are up to 50% lower in muscle from obese compared with lean subjects, and that markers of fatty acid oxidative capacity, including CPT1, correlate inversely with insulin resistance (34). These reports imply that diminished lipid oxidation precedes muscle lipid accumulation and insulin resistance; thus, pharmacological interventions designed to enhance muscle lipid oxidation might facilitate weight loss, lower muscle lipid content, and promote insulin sensitivity. Although this prediction contradicts the classic model by Randle et al. (35), which hypothesizes that increased fatty acid oxidation contributes to insulin resistance through product inhibition of hexokinase, recent studies have challenged this convention by showing that glucose-6-phosphate does not accumulate during increased fatty acid substrate utilization (29). Moreover, in rodent models of obesity and insulin resistance, administration of PPAR-α agonists has been shown to increase whole-body lipid catabolism (8,36) while improving glucose tolerance (5,19,36). Clinical trials in humans have shown either improvement (37,38) or no change (39) in insulin sensitivity indexes. Clearly, future studies using the HSKMC model to evaluate the effects of PPAR-α activation on myocyte glucose homeostasis should provide additional insight into the therapeutic potential of PPAR-α activators in treating patients with type 2 diabetes.

In summary, PPAR-α is a nuclear hormone receptor that is expressed abundantly in human skeletal muscle, but until now, its function in this tissue was essentially unknown. In the present study, we demonstrated that PPAR-α was induced during myocyte differentiation and that its activation stimulated lipid oxidation and decreased TAG accumulation in primary human myotubes. Consistent with these biochemical observations, we identified three hPPAR-α target genes, CPT1, MCD, and PDHK4, each of which is involved in promoting muscle lipid catabolism. These results not only suggest a mechanism for the hypolipidemic effects of PPAR-α–selective drugs in humans but also implicate hPPAR-α as a transcription factor that mediates fatty acid–induced upregulation of these genes in vivo. Additionally, we found that hPDHK4 mRNA expression was more responsive to PPAR-α activation than other known target genes, suggesting a critical role for this gene in regulating muscle’s adaptive response to changes in fuel availability.

This work was supported by Grants NIDDK 46121-06 (to G.L.D.), NIH F32DK 10017-01 (to D.M.), and HL57354 (to W.E.K.) from the National Institutes of Health and a grant from the North Carolina Institute of Nutrition (to D.M. and J.H.).

We thank Dr. Dan Kelly for the MCAD antibody, Jason Berggren for assistance with myocyte isolations, and Dr. Rosalind Coleman for allowing us to perform the lipid analyses in her laboratory.