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Am J Physiol Endocrinol Metab 293: E1256-E1264, 2007. First published August 28, 2007; doi:10.1152/ajpendo.00218.2007
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Selective PPAR{delta} agonist treatment increases skeletal muscle lipid metabolism without altering mitochondrial energy coupling: an in vivo magnetic resonance spectroscopy study

Beat M. Jucker,1 Dewen Yang,5 Warren M. Casey,4 Alan R. Olzinski,1 Carolyn Williams,1 Stephen C. Lenhard,1 Jeffrey J. Legos,3 C. Terrance Hawk,2 Susanta K. Sarkar,5 and Stephen J. Newsholme4

1Cardiovascular and Urogenital Center of Excellence for Drug Discovery, 2Laboratory of Animal Sciences, 3High Throughput Biology, 4Pre-Clinical Development, and 5Technology Development, GlaxoSmithKline, King of Prussia, Pennsylvania

Submitted 7 April 2007 ; accepted in final form 15 August 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Peroxisome proliferator-activated receptor-{delta} (PPAR{delta}) activation results in upregulation of genes associated with skeletal muscle fatty acid oxidation and mitochondrial uncoupling. However, direct, noninvasive assessment of lipid metabolism and mitochondrial energy coupling in skeletal muscle following PPAR{delta} stimulation has not been examined. Therefore, in this study we examined the response of a selective PPAR{delta} agonist (GW610742X at 5 or 100 mg·kg–1·day–1 for 8 days) on skeletal-muscle lipid metabolism and mitochondrial coupling efficiency in rats by using in vivo magnetic resonance spectroscopy (MRS). There was a decrease in the intramyocellular lipid-to-total creatine ratio as assessed by in vivo 1H-MRS in soleus and tibialis anterior muscles by day 7 (reduced by 49 and 46%, respectively; P < 0.01) at the high dose. Following the 1H-MRS experiment (day 8), [1-13C]glucose was administered to conscious rats to assess metabolism in the soleus muscle. The relative fat-vs.-carbohydrate oxidation rate increased in a dose-dependent manner (increased by 52 and 93% in the 5 and 100 mg·kg–1·day–1 groups, respectively; P < 0.05). In separate experiments where mitochondrial coupling was assessed in vivo (day 7), 31P-MRS was used to measure hindlimb ATP synthesis and 13C-MRS was used to measure the hindlimb tricarboxylic acid cycle flux (Vtca). There was no alteration, at either dose, in mitochondrial coupling efficiency measured as the ratio of unidirectional ATP synthesis flux to Vtca. Soleus muscle GLUT4 expression was decreased by twofold, whereas pyruvate dehydrogenase kinase 4, carnitine palmitoyl transferase 1a, and uncoupling protein 2 and 3 expression was increased by two- to threefold at the high dose (P < 0.05). In summary, these are the first noninvasive measurements illustrating a selective PPAR{delta}-mediated decrease in muscle lipid content that was consistent with a shift in metabolic substrate utilization from carbohydrate to lipid. However, the mitochondrial-energy coupling efficiency was not altered in the presence of increased uncoupling protein expression.

peroxisome proliferator-activated receptor-{delta}; intramyocellular lipid; mitochondrial coupling; muscle

THE PEROXISOME PROLIFERATOR-activated receptor (PPAR) subfamilies ({gamma}, {alpha}, and {delta}) are "master" transcriptional regulators for a host of genes that regulate tissue-specific nutrient metabolism and energy homeostasis (2). Whereas PPAR{alpha} and PPAR{gamma} are predominantly expressed in liver and adipose tissue, respectively, PPAR{delta} is ubiquitously expressed (8). Its expression in metabolically active tissues such as skeletal muscle, adipose tissue, liver, intestine, and kidney supports the potential role for PPAR{delta} in regulating metabolic and energy homeostasis.

Until recently, PPAR{delta} was not studied to the same degree as PPAR{alpha} or -{gamma}, in part because selective PPAR{delta} activator availability had been limited. However, recent studies using selective PPAR{delta} agonists have been shown to lower systemic levels of triglycerides (20), increase HDL cholesterol (20, 23), increase tissue-specific lipid oxidation (5, 24), increase insulin sensitivity (17, 26), and protect against diet-induced obesity (26) in a variety of preclinical species. Wang et al. (27) demonstrated that targeted activation of PPAR{delta} in adipose tissue of mice not only induced expression of genes required for fatty acid oxidation but also led to a reduction in adiposity and resistance to weight gain when challenged with a high-fat diet. PPAR{delta} activation leads to an upregulation of energy expenditure by regulating genes involved in fatty acid oxidation (7, 17, 19, 22–24, 26, 27) and mitochondrial uncoupling (7, 19, 22, 24, 26). More recently, it was reported that skeletal muscle-specific PPAR{delta} transgenic mice exhibited an increased number of type 1 muscle fibers and expression of genes implicated in oxidative metabolism (18). Wang et al. (26) further demonstrated that mice overexpressing skeletal muscle-specific PPAR{delta} exhibited increased exercise tolerance, upregulation of mitochondrial gene expression, and decreased propensity for weight gain.

Although these studies do reflect that PPAR{delta}-mediated activation of target gene transcription as well as metabolic regulation occur in skeletal muscle, direct, noninvasive assessment of lipid metabolism in skeletal muscle following selective, pharmacological activation of PPAR{delta} has not been examined. Additionally, whereas thermogenic regulation by PPAR{delta} in skeletal muscle-targeted overexpressing mice (18, 26) or by pharmacological activation of PPAR{delta} in diet-induced obese mice (24, 26) suggests a role for skeletal muscle mitochondrial uncoupling, direct assessment of mitochondrial coupling has not been performed.

In vivo 1H-magnetic resonance spectroscopy (MRS) has been used to noninvasively discriminate intramyocellular lipid (IMCL) from extramyocellular lipid (EMCL) pools in skeletal muscle (21), thereby allowing for temporal assessment of lipid-pool changes. In addition, an MRS assay has been used to assess mitochondrial energy coupling in skeletal muscle, noninvasively, by combining 13C-MRS to measure rates of mitochondrial substrate oxidation and 31P-MRS to assess rates of ATP synthesis in chronic triiodo-L-thyronine-treated rats [a model of increased uncoupling protein (UCP) 3 expression] (14). Therefore, in the present study, we examined the temporal and dose-dependent responses of selective PPAR{delta} agonist treatment on skeletal muscle lipid metabolism and mitochondrial energy coupling in rats by MRS.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Animals. Sprague-Dawley rats (Charles River, Raleigh, NC) were housed in an environmentally controlled room with a 12-h light/dark cycle. Animals were treated with a selective PPAR{delta} agonist, GW610742X (5 or 100 mg/kg once daily for 7 days; n = 24 per group) or vehicle (n = 24) via oral gavage. GW610742X is >200- and >400-fold more selective for the murine PPAR{delta} receptor than for PPAR{alpha} and PPAR{gamma} receptors, respectively (25). Although GW610742X has high protein-binding capacity (>99%) and a resulting low plasma free drug concentration at the doses used in the study, there is still the potential for cellular PPAR{alpha} activation at the high GW610742X dose used. Animals were cared for by following the recommendations contained in the Guide for the Care and Use of Laboratory Animals (National Research Council, 1996). All procedures were approved by the Institutional Animal Care and Use Committee of GlaxoSmithKline and were specifically designed to minimize animal discomfort.

Intramyocellular lipid measurement. Rats (n = 8 per group) were anaesthetized with 2.0% isoflurane (Abbott Laboratories, Chicago, IL) during the nuclear magnetic resonance (NMR) experiments. Magnetic resonance imaging (MRI) and 1H-MRS were performed on days 0 and 7. All in vivo 1H-MRS/MRI experiments were performed on a Bruker Biospec 4.7T system (horizontal, 40-cm-diameter bore magnet; Bruker Medical, Billerica, MA). A 1H radio-frequency (RF) surface coil (42 mm diameter) tuned to 200.21 MHz was used. The rat was positioned prone to the RF coil/platform assembly (horizontal in plane), and the right hindlimb was placed so that the lower hindlimb was placed in the magnet isocenter. Coronal and axial gradient echo [repetition time (TR)/echo time (TE) = 120/9 ms] pilot images were acquired before performing spatially localized IMCL and EMCL measurements. Proper placement of a spectroscopic region of interest (ROI) in the soleus and tibialis anterior muscles for each imaging session was achieved by using the following anatomical landmarks: 15 mm distal to the head of the tibia in the coronal slice and ~1–3 mm from the tibia in the axial slice for the soleus or tibialis anterior voxel placement. The muscle-fiber orientation along the magnetic field lends itself to optimal discrimination of IMCL (1.3 ppm) from EMCL (1.5 ppm) NMR peaks. The fiber orientation along the magnetic field is critical in obtaining spectral resolution between the IMCL and EMCL NMR peaks (3). 1H point-resolved spatially localized spectroscopy [PRESS; TR/TE = 1,000/20 ms, no. of scans (NS) = 512, sweep width (SW) = 5,000 Hz] with chemical shift selective (CHESS) water suppression was performed over a 3 x 3 x 3-mm voxel in the soleus and tibialis anterior. For relative IMCL, EMCL, and total creatine (tCr) peak-area analysis, 1H-magnetic resonance spectra were processed by using a Gaussian filter, and baseline flattening before automated peak deconvolution was performed by a simplex routine (Nuts NMR processing software; Acorn NMR, Fremont, CA).

Mitochondrial energy-coupling measurement. In a separate group of animals (n = 8 per group), the efficiency of mitochondrial energy coupling was assessed. Because the tricarboxylic acid (TCA)-cycle activity is coupled to O2 consumption via a stoichiometric relationship, the flux through the TCA cycle may be used as an index of substrate oxidation at steady state. Therefore, the ratio of the measured unidirectional ATP-synthesis flux to the TCA cycle flux (Vtca) may be used as a qualitative index of the degree of coupling between mitochondrial substrate oxidation and ATP synthesis (14). Because the extent of basal mitochondrial uncoupling present as a result of combined proton transport and leaks across the inner mitochondrial membrane is unknown, this ratio was normalized to the vehicle-dose group. As a positive control to illustrate that increased mitochondrial uncoupling could be measured by using the MRS assay, triiodo-L-thyronine and 2,4-dinitrophenol were previously shown to increase uncoupling by increasing TCA cycle flux in the absence of changes in unidirectional ATP synthesis (14).

All in vivo NMR experiments were performed on a Bruker Biospec 4.7T system. Both 13C observe/1H decouple and 31P observe/1H decouple MRS were performed by using concentric surface RF coils [the outer 1H RF coil (42 mm) tuned to 200.21 MHz and the inner dual-frequency 13C or 31P RF coil (25 mm) tuned to 50.34 or 81.05 MHz, respectively]. Rats were anaesthetized with 2% isoflurane, and the hindlimb was positioned over the 13C/31P RF coil (horizontal in plane) and placed in the magnet isocenter. Because of the limited 13C/31P sensitivity in the hindlimb experiments, it was necessary to measure the bulk signal from the larger tissue beds of mixed fiber type, including the gastrocnemius and biceps femoris. The ATP-synthesis flux was assessed by using a saturation-transfer experiment as previously described (15). To measure the kinetics of Pi -> ATP, a continuous-wave selective saturation of the {gamma}-ATP resonance was used (TR = 4.3 s, NS = 208, SW = 5,000 Hz). The unidirectional ATP-synthesis flux was calculated as unidirectional rate constant x Pi concentration. The Pi concentration was extrapolated from the baseline NMR spectrum (comparing peak integrals from Pi and {gamma}-ATP) and ATP concentration. Following the 31P measurement, 13C-MRS was used to measure the Vtca by examining labeled glutamate turnover (Fig. 1) as follows: [2-13C]acetate (sodium salt, 99% enriched; Cambridge Isotope Laboratories, Cambridge, MA) was administered to study the glutamate-labeling kinetics for Vtca measurements (15). The [2-13C]acetate infusion consisted of an intravenous bolus (100 mg/kg body wt) for 1 min followed by a 138 µmol·kg–1·min–1 continuous infusion for 120 min (sufficient time required to achieve isotopic steady state). The 13C observe/1H decouple experiment was performed as previously described (15). Briefly, a 15-min lipid-suppressed baseline spectrum was followed by subsequent 15-min acquisitions throughout the duration of the experiment (TR = 0.5 s, NS = 1,455, SW = 10,000 Hz). At the end of the in vivo MRS experiment, animals were anesthetized with Nembutal (50 mg/kg; Abbot Laboratories, Chicago, IL). Superficial skin was rapidly removed from the left hindquarter, followed by in situ freeze clamping of the gastrocnemius and biceps femoris muscles. Rats were euthanized with a lethal dose of Nembutal.


Figure 1
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  		Fig. 1. Intramuscular label flow schematic resulting from [2-13C]acetate precursor administration. Labeling of [4-13C]glutamate/glutamine (Glu/Gln) occurs as a result of label entering tricarboxylic acid (TCA) cycle via condensation of [2-13C]acetyl-CoA (AcCoA) with oxaloacetate (Oaa). With multiple turns of TCA cycle, label will appear at C2, C3, and C4 isotopomer positions of Glu and Gln and C2 and C3 isotopomer positions of Oaa. Labeled intermediates and associated fluxes were used to develop isotopic steady-state mathematical model for calculating TCA cycle flux (Vtca) (15). Cit, citrate; Mal, malate; Pyr, pyruvate; Asp, aspartate.

 
The absolute Vtca was calculated by using metabolic steady-state equations that were derived for isotopic mass flow into the TCA cycle as previously described (15).

Relative fat vs. carbohydrate oxidation measurement in muscle. On day 8 following the 1H-MRS experiment and IMCL measurement, overnight-fasted rats were subjected to a glucose-somatostatin infusion via jugular catheter. [1-13C]glucose (8 mg·kg–1·min–1, 99% enriched; Cambridge Isotope Laboratories), set to match the hepatic glucose-production rate, was administered together with somatostatin (0.15 µg/min) to maintain euinsulinemia for 120 min in the awake rat. This time period was sufficient for glycolytic and TCA cycle intermediates to achieve steady-state enrichments. At the end of the experiment, rats were anaesthetized with Nembutal (50 mg/kg). Superficial skin was rapidly removed from the left hindlimb, followed by harvesting of the soleus muscle for biochemical and gene-expression analysis. Proton observe-carbon edited 1H-MRS measurement of metabolite 13C enrichment in tissue extracts was performed on a 9.4T Bruker system (16). Relative carbohydrate and fat oxidation in terms of relative substrate contribution to acetyl-CoA oxidation was assessed from the metabolite-pool enrichments (28, 29).

Analytical procedures. Plasma clinical chemistry endpoints, including glucose, triglyceride, nonesterified free fatty acids (NEFA), LDL, HDL and total cholesterol concentrations, were determined enzymatically by using an Olympus AU600 analyzer (Olympus America, Mellville, NY).

The skeletal muscle Pi concentration was extrapolated from the baseline 31P-NMR spectrum (comparing peak integrals from Pi and {gamma}-ATP and measured ATP concentration; ATP bioluminescence assay kit FL-AA from Sigma, modified for tissue analysis). Skeletal muscle glutamate and lactate concentrations were measured in a similar manner as previously described (13).

Microarray gene-expression analysis. Frozen soleus muscle samples were lysed in TRIzol reagent (Invitrogen, Carlsbad, CA), RNA was precipitated with isopropanol, and the pellet was washed with 80% ethanol and resuspended in diethyl pyrocarbonate-H2O. RNA quality was checked by using an Agilent bioanalyzer (Agilent Technologies, Palo Alto, CA). RNA was purified through a Qiagen column (Qiagen, Valencia, CA) and was quantified by using OD260 absorbance. RNA samples were analyzed by using Affymetrix GeneChip RAE230A (Affymetrix, Santa Clara, CA) with standard protocols. Affymetrix GCOS 1.1 with MAS5 were used to extract data from the GeneChip image files. Data were loaded into Rosetta Resolver (Rosetta Biosoftware, Seattle, WA) for selection of differentially expressed probe sets, which were identified by using the normalization and statistical package included in Rosetta Resolver v. 5.1, build 5.1.0.0.163. Disregulation of selected Affymetrix probe sets was confirmed by using RT-PCR.

Statistical analysis. All data are reported as means ± SE. An ANOVA was performed to determine significance at a minimum P < 0.05 threshold. A multiple-comparison Newman-Keuls post hoc test was used when necessary to determine significance at different time points.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Physiological parameters. The body weights remained constant throughout the short-duration dosing regimen in the vehicle and low-dose (5 mg·kg–1·day–1 GW610742X) groups (Table 1). However, there was a temporal decrease in body weight observed in the high-dose (100 mg·kg–1·day–1 GW610742X) group, resulting in an 11% loss in body weight by day 7 (Table 1). Although there was a trend toward decreased NEFA in both GW610742X groups, there were no significant differences in glucose and lipid parameters following 7-day treatment in either GW610742X group (Table 1).


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  		Table 1. Summary of body weight and plasma lipids

 
IMCL measurement. Coronal and axial slice images of the rat hindlimb are shown in Fig. 2A. Scout images such as those shown in Fig. 2A were used to position the spectroscopic ROI (3 x 3 x 3 mm) in the soleus and tibialis anterior muscles. Typical 1H spectra obtained from the soleus muscle are shown in Fig. 2, B and C. The IMCL, EMCL, and tCr MRS peaks appear at 1.3, 1.5, and 3.1 ppm, respectively. An example of superimposed baseline and 7-day vehicle-treatment spectra obtained from the soleus muscle is shown in Fig. 2B. There were no differences in IMCL/tCr ratio in the soleus throughout the 7-day treatment period in the vehicle group (Fig. 2D). An example of superimposed baseline and 7-day high-dose group spectra obtained from the soleus is shown in Fig. 2C. Although there was a slight, nonsignificant decrease in the soleus IMCL/tCr ratio at day 7 in the low-dose group, there was a significant decrease in the IMCL/tCr ratio in the soleus muscle by day 7 ({downarrow}49%, P < 0.001, Fig. 2D) in the high-dose group. Additionally, there also was a slight, nonsignificant decrease in the tibialis anterior IMCL/tCr ratio at day 7 in the low-dose group, whereas there was a significant decrease in the IMCL/tCr ratio in the tibialis anterior muscle by day 7 (46%; P < 0.01; Fig. 2D) in the high-dose group.


Figure 2
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  		Fig. 2. Volume-selective 1H-magnetic resonance spectroscopy (MRS) in soleus and tibialis anterior muscle beds. Coronal and axial gradient echo scout images were acquired to achieve proper placement of spectroscopic region of interest (ROI) in soleus or tibialis anterior muscle bed (A). 1H point-resolved spatially localized spectroscopy MRS with chemical shift selective water suppression was performed over a 3 x 3 x 3 mm ROI in muscle. Total creatine (tCr), extramyocellular (EMCL), and intramyocellular (IMCL) peaks appear at 3.1, 1.5, and 1.3 ppm, respectively. Representative spectra from soleus illustrating baseline and 7-day vehicle (B) and high-GW610742X-dose (C) treatment spectra superimposed are shown. IMCL/tCr ratio in vehicle, low-, and high-GW610742X-dose groups at baseline (closed bars) and day 7 (open bars) is shown for both soleus and tibialis anterior muscles (D). Data are presented as means ± SE. *P < 0.01 vs. baseline; **P < 0.001 vs. baseline.

 
Relative fat vs. carbohydrate oxidation. One day after the IMCL MRS experiment, [1-13C]glucose was administered to conscious rats to assess the relative fat vs. carbohydrate oxidation in the soleus muscle. The relative fat-oxidation rate, represented as the percentage of the acetyl-CoA units generated by fatty acids and ketone bodies, increased in a dose-dependent manner (by 52% and 93% in the 5 and 100 mg·kg–1·day–1 groups, respectively; P < 0.05; Fig. 3). Conversely, the relative carbohydrate oxidation, represented as the percentage of acetyl-CoA units generated by aerobic glycolysis, decreased in both the low- and high-dose groups (by 20 and 36%, respectively; P < 0.05).


Figure 3
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  		Fig. 3. Relative carbohydrate vs. fat oxidation in soleus muscle performed on day 8. Relative carbohydrate and fat oxidation, presented as relative contribution of each of these substrate sources to acetyl-CoA oxidation, was assessed in vehicle (open bars), low- (closed bars), and high- (gray bars) GW610742X-dose groups. This measurement was achieved by assessing steady-state [3-13C]lactate, [3-13C]alanine, and [4-13C]glutamate enrichments in tissue extracts following an in vivo administration of [1-13C]glucose as described in MATERIALS AND METHODS. Data are presented as means ± SE. *P < 0.05 vs. vehicle for relative carbohydrate and fat oxidation.

 
Mitochondrial energy coupling. A 15-min acquired 13C-MRS spectrum of the rat hindlimb taken between 105 and 120 min may be seen in Fig. 4A. The 13C-label turnover of [4-13C]glutamate followed by slower turnover of [2-13C]glutamate may be seen in Fig. 4A, inset. The 4-13C- and 2-13C-labeled glutamate peaks appeared at 34.4 and 55.5 ppm, respectively, in the first spectrum and increased until steady-state signal intensities were achieved (~60–90 min). Labeling of [4-13C]glutamate was the result of [2-13C]acetyl-CoA condensing with oxaloacetate to produce [4-13C]citrate, which in turn labeled {alpha}-ketoglutarate and glutamate at the C4 position (Fig. 1). The [2-13C]glutamate peak appears as a result of additional turns of the TCA cycle, which allows for scrambling the C4 label of glutamate to 2-13C- and 3-13C-labeled glutamate (27.9 ppm). The [3-13C]glutamate and [2-13C]acetate (24.2 ppm) peaks were not observable, because they reside in the frequency bandwidth that was partially suppressed by the aliphatic lipid-suppression pulse sequence.


Figure 4
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  		Fig. 4. 13C-MRS glutamate-turnover study. A 15-min acquired spectrum from rat hindlimb at 105–120 min is shown (A). [2-13C]glutamate peak appears at 55.5 ppm on shoulder of creatine/phosphocreatine peak (54.4 ppm), and [4-13C]glutamate peak obscured by a coresonating aliphatic lipid peak appears at 34.4 ppm. Turnover of [2-13C]- and [4-13C]glutamate peaks in hindlimb muscles of a vehicle-treated rat is illustrated in baseline-subtracted spectra shown in inset. [2-13C]acetate label incorporates rapidly into [4-13C]glutamate and more slowly into [2-13C]glutamate (second turn of TCA cycle). Calculated TCA cycle flux (Vtca) in vehicle, low-, and high-GW610742X-dose treatment groups are shown (B). Data are presented as means ± SE.

 
There were no differences in the glutamate pool size following acetate administration in the vehicle vs. low- and high-dose groups (1.89 ± 0.39, 1.18 ± 0.17, and 1.30 ± 0.05 µmol/g, respectively). There were no differences in the calculated Vtca in the vehicle, low-, and high-dose groups (426 ± 128, 404 ± 136, and 396 ± 127 nmol·g–1·min–1, respectively; Fig. 4B). Additionally, there were no differences in the calculated anaplerosis in the vehicle, low-, and high-dose groups (19 ± 7, 17 ± 2, and 22 ± 13% of Vtca, respectively) as reflected by the ratio of [3-13C]glutamate enrichment to [4-13C]glutamate enrichment.

A 31P-NMR saturation-transfer experiment was performed to determine the kinetics of ATP synthesis. The set of spectra shown in Fig. 5A are the result of a saturation-transfer experiment performed in the hindlimb muscles of a rat. In the bottom spectrum, a continuous-wave RF pulse was used to saturate the {gamma}-ATP resonance (–2.4 ppm). In the top spectrum (no {gamma}-ATP saturation), the continuous-wave RF pulse was placed symmetrically to the downfield side of Pi (4.9 ppm). The resulting loss in magnetization of Pi ({Delta}M) was used to calculate the unidirectional ATP-synthesis flux. There were no differences in unidirectional ATP-synthesis flux in the vehicle, low-, and high-dose groups (47.8 ± 10.4, 29.1 ± 8.8, and 56.3 ± 14.2 nmol·g–1·s–1 respectively; Fig. 5B).


Figure 5
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  		Fig. 5. 31P-MRS saturation-transfer study. Spectra shown were obtained from a saturation-transfer experiment performed in rat hindlimb muscles (A). In lower spectrum, a continuous-wave radio frequency pulse was used to saturate {gamma}-ATP resonance (–2.4 ppm). In upper spectrum (no {gamma}-ATP saturation), continuous-wave pulse frequency was placed symmetrically to downfield side of Pi (4.9 ppm). Resulting loss in magnetization of Pi ({Delta}M) is due to exchange of saturated {gamma}-ATP nuclei with nonsaturated Pi nuclei. Calculated unidirectional ATP synthesis flux in vehicle, low-, and high-GW610742X-dose groups are shown (B). Data are presented as means ± SE.

 
The TCA cycle generates reducing equivalents (NADH, FADH2), which are necessary for mitochondrial respiration. Because the TCA cycle activity is coupled to O2 consumption via a stoichiometric relationship, the flux through the TCA cycle may be used as an index of substrate oxidation at steady state. Therefore, the ratio of the measured unidirectional ATP synthesis flux to Vtca may be used as a qualitative index of the degree of coupling between mitochondrial-substrate oxidation and ATP synthesis. Because the extent of basal mitochondrial uncoupling present as a result of combined proton transport and leaks across the inner mitochondrial membrane is unknown, this ratio was normalized to the vehicle-dose group. When analyzed in this manner, there were no differences in mitochondrial energy coupling between vehicle and low- and high-dose groups (1.00 ± 0.23, 0.75 ± 0.08, and 1.06 ± 0.21, respectively).

Gene expression. Expression of both glucose and lipid metabolism/thermogenesis PPAR{delta} target genes were examined in soleus muscle by Affymetrix gene-array analysis. Gene changes shown in Table 2 are presented as fold change in the GW610742X-treated groups vs. vehicle-only group. Key metabolic genes involved in the regulation of fat metabolism [i.e., carnitine palmitoyl transferase 1 (CPT1), acyl-CoA dehydrogrenase, hormone-sensitive lipase, peroxisomal D3,D2-enoyl-CoA isomerase, etc.] and potential thermogenesis (i.e., UCP2, UCP3) were significantly increased in the high-dose group but more moderately in the low-dose group. Glucose transporter GLUT4 (SLC2A4) was downregulated, and pyruvate dehydrogenase modulator gene (PDK4) was increased at the high GW610742X dose (Table 2). PPAR{gamma} coactivator-1{alpha} (PGC-1{alpha}) was not changed at either GW610742X dose (Table 2). These data support the metabolic shift from glucose to fat metabolism observed in the soleus muscle.


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  		Table 2. Summary of metabolic target-gene activation by GW610742X

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, we show for the first time, non-invasively, that IMCL content is decreased in soleus and tibialis anterior muscles on selective pharmacological activation of PPAR{delta} in rats. This change was also associated with a shift toward increased lipid vs. carbohydrate oxidation in soleus muscle, which was consistent with PPAR{delta} target gene regulation in this tissue. Although UCP2 and UCP3 gene expression were increased, there was neither an increase in energy expenditure nor a difference in mitochondrial energy coupling in the hindlimb of GW610742X-treated rats despite a decrease in body weight in the high-dose group animals. These data are in part consistent with previous studies (5, 17, 24, 26, 27) that show PPAR{delta}-mediated regulation of thermogenic and lipid beta-oxidation genes was associated with a phenotype (e.g., body weight loss, increased insulin sensitivity).

In the present study, skeletal muscle was targeted for metabolic measurements by in vivo MRS because PPAR{delta} is expressed at high levels, with PPAR{alpha} and PPAR{gamma} isoforms expressed at lower levels in skeletal muscle (4, 19, 22). IMCL and relative substrate-metabolism measurements were made in soleus (predominantly type I, oxidative/slow fiber) muscle because it was thought that this muscle bed would definitively show a PPAR{delta} phenotypic effect, given that expression of the PPAR{delta} receptor is higher in rodent type I vs. type II muscle fiber (26). However, the tibialis anterior muscle, which is mixed type I and type II muscle fiber, exhibited a similar reduction in IMCL. This result is consistent with the hypothesis that PPAR{delta} may regulate type I muscle-fiber content or total fiber number in this muscle (18). The hypothesis that selective PPAR{delta} agonist treatment could increase skeletal-muscle lipid oxidation and commensurately reduce IMCL content was based on data from previous studies reflecting increased expression of genes involved in lipid oxidation such as CPT1 (17, 24), fatty acid transporter (24), and acyl-CoA dehydrogenase, long chain (24) following GW501516 (selective PPAR{delta} agonist) activation. Although absolute IMCL content has not been measured in rodent skeletal muscle on selective PPAR{delta} activation to date, Tanaka et al. (24) have shown via electron-microscopic analysis of the skeletal muscle from GW501516-treated mice that mitochondrial density was increased and lipid droplets were significantly reduced. Although we were able to show in the same animals that there was an increase in relative lipid vs. carbohydrate oxidation in soleus muscle with GW610742X treatment, absolute substrate-oxidation rates could not be measured in this manner. Nevertheless, these in vivo results are consistent with published ex vivo data including absolute palmitate-oxidation measurements in quadriceps (24) or soleus (17) from GW501516-treated mice and oleate oxidation in GW610742X-treated primary human myotubes (19). Additionally, the downregulation of GLUT4 expression in soleus muscle in the present study was consistent with that found in GW501516-incubated mouse skeletal muscle cells (7) and supports the shift in substrate utilization observed.

Both Wang et al. (26) and Tanaka et al. (24) have shown that treatment with GW501516 resulted in a slower growth rate in high-fat-fed C57BL/6 mice, which was associated with increased glucose tolerance. The difference in weight gain was in large part due to a reduction in fat-pad weight because heart and skeletal muscle weights were unchanged. Additionally, although adipose tissue mass was reduced in these mice, skeletal muscle mitochondrial biogenesis was also implicated because cytochrome c expression was increased. In the same study, investigators showed that mice with skeletal muscle-targeted overexpression of PPAR{delta} exhibited increased exercise tolerance. Although we did not examine exercise tolerance in the present study, we were able to elicit a body-weight reduction in the high GW610742X dose-treated animals on normal diet. Anaplerosis was not increased on PPAR{delta} activation in our study, so a reduction in whole body adipose-tissue mass rather than increased protein catabolism may be responsible for the observed weight loss in the drug-treated group. However, food consumption was not monitored in the present study and as a result could explain the difference in weight gain between groups. Although obvious changes in the plasma lipid profile of PPAR{delta}-treated animals have been reported (17, 20, 25), we did not observe an effect on HDL raising. Perhaps species differences may have accounted for this lack of HDL increase as was previously observed in Sprague-Dawley rats treated for as long as 9 wk with GW610742X (11). Current use of PPAR{delta} agonist GW501516 in the clinic has shown benefits in raising HDL and increasing clearance of triglycerides in healthy volunteers following a high-fat meal (23). Therefore, a dyslipidemic patient population with cardiovascular risk may benefit from PPAR{delta} agonist treatment.

Mitochondrial uncoupling proteins play an integral role in regulating cellular energy consumption via nonshivering thermogenesis (10). This regulation is accomplished by diminishing the proton-motive force across the inner mitochondrial membrane, which results in uncoupling of respiration from ATP synthesis. Because quiescent skeletal muscle accounts for 33% of whole body oxygen consumption (9), much attention has been given to the control and function of UCP3 as a means of regulating energy expenditure and body weight. A number of groups have shown that both UCP2 (7, 24) and UCP3 (7, 19, 22, 24) are upregulated in either cultured skeletal muscle cells treated with PPAR{delta} agonist or in skeletal muscle derived from animals treated with an agonist. These results are consistent with our findings that UCP2 and UCP3 expression were increased over twofold in soleus muscle. Whereas thermogenic-program genes have been shown to be upregulated in myotubes with GW501516 treatment, both PPAR{alpha}- and PPAR{gamma}-specific agonists did not have this effect (24). The activation of the mitochondrial biogenesis program could in part be due to activation of PGC-1{alpha}, which contains a conserved PPAR response element. However, in the present study as well as in PPAR{delta}-overexpressing mice (18, 26), PGC-1{alpha} expression was found not to be increased. Although there is clear evidence for whole body thermogenic activation on PPAR{delta} agonist treatment, there is no definitive evidence that skeletal muscle represents the target tissue. In fact, not only were the in vivo mitochondrial energy-coupling measurements similar between vehicle- and GW610742X-treated animals in our study, but the TCA cycle flux was similar as well, indicative of similar oxidative rates between groups.

More recently, Constantin et al. (6) have shown that activation of lipid metabolism, but not mitochondrial dysfunction, occurred in isolated mitochondria from soleus muscle in GW610742-treated rats. It is plausible that UCP3 may not function in a regulatory thermogenic capacity following PPAR{delta} activation. It has been shown by Beinengraeber et al. (1) that simultaneous mutation of histidine residues His145 and His147 results in loss of the proton-transport capacity of UCP1, and both of these residues are absent in mouse and rat UCP3. One of the limitations of the present study is that the mitochondrial-coupling MRS measurements are performed in the entire hindlimb skeletal musculature. Because of the relatively low sensitivity of the 13C-NMR measurement, it was necessary to perform the measurement over the bulk muscles of the hindlimb, comprised of both type I and type II muscle fibers. As a result, metabolic differentiation in the soleus alone would be extremely difficult to achieve. However, it is plausible that a metabolic perturbation such as mitochondrial uncoupling may be realized in the entire hindlimb given that PPAR{delta} skeletal muscle-overexpressing mice exhibit increased oxidative muscle-fiber formation in the entire hindlimb, including gastrocnemius (26), plantaris (26), and tibialis anterior (18) muscle beds.

In summary, these are the first noninvasive measurements illustrating a selective PPAR{delta}-mediated decrease in soleus muscle lipid content that was consistent with a shift in metabolic substrate utilization from carbohydrate to lipid. This metabolic shift in soleus muscle was supported by changes in target gene expression in this muscle. Additionally, the thermogenic status of the hindlimb muscles was evaluated. However, the mitochondrial energy-coupling efficiency was not altered in the presence of increased uncoupling protein expression. These data support the notion that the target tissue for PPAR{delta}-mediated thermogenic regulation may be the adipose tissue rather than the skeletal muscle. The metabolic activation in muscle may serve to preserve metabolic requirements necessary for periods of increased physical load (i.e., exercise).


   ACKNOWLEDGMENTS
 
We acknowledge Thomas Schaeffer and Raymond White for expert care and handling of the animals in support of the imaging studies. We also acknowledge Dr. Lea Sarov-Blat and Dr. Mark Hurle for support in genomic pathway analysis.


   FOOTNOTES
 

Address for reprint requests and other correspondence: B. M. Jucker, GlaxoSmithKline, UW2510, 709 Swedeland Rd., King of Prussia, PA, 19406 (e-mail: beat.m.jucker{at}gsk.com)

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
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 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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