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Triglyceride:High-Density Lipoprotein Cholesterol Effects in Healthy Subjects Administered a Peroxisome Proliferator Activated Receptor δ Agonist

Originally published, Thrombosis, and Vascular Biology. 2007;27:359–365


Objectives— Exercise increases fatty acid oxidation (FAO), improves serum high density lipoprotein cholesterol (HDLc) and triglycerides (TG), and upregulates skeletal muscle peroxisome proliferator activated receptor (PPAR)δ expression. In parallel, PPARδ agonist-upregulated FAO would induce fatty-acid uptake (via peripheral lipolysis), and influence HDLc and TG-rich lipoprotein particle metabolism, as suggested in preclinical models.

Methods and Results— Healthy volunteers were allocated placebo (n=6) or PPARδ agonist (GW501516) at 2.5 mg (n=9) or 10 mg (n=9), orally, once-daily for 2 weeks while hospitalized and sedentary. Standard lipid/lipoproteins were measured and in vivo fat feeding studies were conducted. Human skeletal muscle cells were treated with GW501516 in vitro and evaluated for lipid-related gene expression and FAO. Serum TG trended downwards (P=0.08, 10 mg), whereas TG clearance post fat-feeding improved with drug (P=0.02). HDLc was enhanced in both treatment groups (2.5 mg P=0.004, 10 mg P<0.001) when compared with the decrease in the placebo group (−11.5±1.6%, P=0.002). These findings complimented in vitro cell culture results whereby GW501516 induced FAO and upregulated CPT1 and CD36 expression, in addition to a 2-fold increase in ABCA1 (P=0.002). However, LpL expression remained unchanged.

Conclusions— This is the first report of a PPARδ agonist administered to man. In this small study, GW501516 significantly influenced HDLc and TGs in healthy volunteers. Enhanced in vivo serum fat clearance, and the first demonstrated in vitro upregulation in human skeletal muscle fat utilization and ABCA1 expression, suggests peripheral fat utilization and lipidation as potential mechanisms toward these HDL:TG effects.

A specific PPARδ agonist was administered to human volunteers for the first time, revealing a decline in serum TG, an improvement in TG-clearance post-fat feeding, and an elevation in HDLc compared with placebo. Consistent with these findings, in vitro PPARδ-treated human skeletal muscle cells induced fatty acid oxidation, and upregulated ABCA1 expression.

Peroxisome proliferator-activated receptors (PPARs) are a family of nuclear receptors designated PPARα (NR1C1), PPARδ/β (NR1C2), and PPARγ (NR1C3) which serve as transcription factors.1,2 PPARs control the cascade of fatty acid intracellular metabolism. Although agonists of the PPARα (fibrates) and γ isoforms (thiazoladinediones) are currently marketed, a specific PPARδ agonist has not heretofore been administered to human subjects. Preclinical animal models have suggested that PPARγ stimulation promotes fat storage in cells, particularly in adipose tissue where this PPAR is especially abundant.3,4 In contrast, activation of PPARα and δ enhance fatty acid oxidation (FAO), leading to increased energy production and possible uncoupling. Both PPARα and δ activation have similar transcriptional effects on the genes associated with fat cellular uptake (eg, the fatty acid translocator, CD36/FAT) and metabolism (eg, carnitine palmitoyl transferase-1, CPT1), but their tissue specificity differs. PPARδ is ubiquitously expressed, in contrast to PPARα which has substantial activity in the liver. Muscle contains both PPAR α and δ, although PPARδ appears to predominate.5,6

It has been reported that exercise mediates upregulation of PPARδ7 and creates a requirement for an external (or serum-derived) triacylglycerol energy source. Lipoproteins are the major source of such combustible substrate, thereby suggesting the interplay between PPAR activity and lipoprotein modification. Mediation of this lipoprotein:cell transfer of lipid is processed through lipolytic activity, and influenced by scavenger receptor B1 (SR-B1) and ATP-binding cassette transporter protein-A1 (ABCA1) mechanisms. These mechanisms could be responsible for the changes in HDLc and TG concentrations observed with physical activity.8 HDLc is known to increase with exercise, and there is a tendency for HDLc to decrease with a drop in physical activity, the latter perhaps based on a similar mechanism.9 In support of this hypothesis, Tikkanen et al identified a striking correlation between increased carnitine palmitoyl transferase activity (CPT1; a protein that transports fats across the mitochondrial membrane) and elevated HDLc in the setting of exercise.10 Potentially, the administration of a synthetic PPARδ ligand can induce an “exercise-like” alteration in HDL. Consistent with this proposal, PPARδ-directed changes in the skeletal muscle of mice result in marked increases in exercise tolerance.11

The data presented here represent the first in vivo experimental human data reported with a PPARδ agonist. We demonstrate an enhanced HDLc response, as well as TG metabolism compared with placebo. Additionally, our findings suggest upregulated fat utilization/clearance as responsible, in part, for these changes.


Clinical Study Design and Efficacy End Points

Healthy white normolipidemic male volunteers were randomly allocated placebo (n=6) or GW501516 in one of two doses: 2.5 mg (n=9) or 10 mg (n=9), orally, once-daily for 2 weeks during hospitalized care. GW501516 is a potent (EC50 of 1.2 nm ±0.1 nmol/L) and specific PPARδ agonist in vitro (>1200-fold selective for PPARδ relative to α and γ).12–14 Therefore, the doses of GW501516 used in this study did not crossover to PPARα or PPARγ. During this study, subjects were sedentary and prohibited from performing rigorous physical activity. They received a diet of approximately 2000 kcal and 62.6 g of fat (26% of total calories) per day. Safety and tolerability were evaluated through adverse events, vital signs, 12-lead ECG telemetry, and clinical laboratory measurements. Standard lipids/lipoproteins were measured and analyzed after an overnight fast before, during, and after the 14-day dosing period. GW501516 plasma levels were measured on Days 10, 12, and 14 (predose) and at designated times on day 14. The study was conducted at Northwick Park Hospital, Harlow, UK. Written informed consent was obtained before each subject could participate in the study in accordance with the relevant Institutional Ethics Committee. The study was conducted in accordance with “good clinical practice” (GCP) and all applicable regulatory requirements, including the 1996 version of the Declaration of Helsinki and the Institutional Ethics Committee.

Total plasma cholesterol and TG concentrations were determined by enzymatic methods using a Hitachi 917 Biochemical Analyzer (Hitachi Ltd, Japan). HDLc was measured enzymatically (Boehringer Mannheim). Low-density lipoprotein cholesterol (LDLc) was calculated by the Friedewald equation. Apolipoprotein AI (apoAI) concentrations were determined by immunonephelometry (Dade Behring BNII Nephelometer); interassay CVs were <4.3%.

A fat feeding study was conducted in which subjects were fed a high-fat (20 g) breakfast on Days 0 and 13 of the study at 1 hour post-dose. Subjects were fasted for more than 10 hours before receiving the meal. Subjects received a low fat lunch, dinner, and snack at 4, 10, and 14 hours post-dose, respectively. Blood samples were collected at 0, 1, 2, 3, 4, 6, 8, 10, 12, 15, and 24 hours post-fat feeding for TG measurements. TG analysis was performed as described above.

Fatty Acid Oxidation and Gene Expression Analyses in Human Skeletal Muscle Cells

Human skeletal muscle (HSKM) cells were purchased from Cambrex BioScience (Walkersville, Md). Cells were plated at 100 000 cells per well in a 12-well Costar plate and dosed in DMEM, 10% FBS with 10 nmol/L GW501516 or DMSO (vehicle control) for 2 days. This dose of GW501516 was chosen to provide the maximum response based on published dose-response experiments.14,15 The day before harvesting for gene expression or FAO analysis, cells were re-fed with the same media along with fresh compound. FAO assays were performed as described5,16 (see supplemental Methods, available online at

Gene expression analyses were performed as described.17 Total RNA was isolated using the Promega SV RNA isolation system according to the manufacturer’s instructions. Real-time quantitative polymerase chain reaction was performed using an ABI PRISM 7700 sequence detection system instrument and software (Applied Biosystems, Inc). Gene-specific primers and probes were designed using Primer Express Version 2.0.0 (Applied Biosystems, Inc) and synthesized by Keystone Laboratories (supplemental Table I and Methods).

Statistical Analyses

Lipid response data were evaluated for within group changes using paired t tests and between group differences using unpaired t tests. Change from baseline (CFB) values for the treatment groups were compared with placebo using mixed models in SAS (V9.1, SAS Institute) accounting for baseline lipid value as a covariate. For the fat feeding study, TG concentrations were measured over 24 hours after fat feeding at Days 0 and 13. Maximum TG values (TGmax) and area under the TG curve (TG-AUC) were calculated at each time period. Correlations between change in TG-AUC and serum TG were evaluated with simple Pearson correlation coefficients followed by analysis of variance models adjusting for baseline TG.


Clinical Results

The mean age of study subjects was 30±5 years, and mean body mass index was 24.0±3 with no significant between-group differences for either measure. GW501516 was safe and well-tolerated. Subjects treated with GW501516 had no significant adverse effects, including liver and muscle responses. We measured liver enzymes (total bilirubin, alkaline phosphatase, aspartate transaminase, alanine transaminase) and muscle proteins (CPK, aldolase, creatinine) for any changes associated with GW501516 treatment. None were noted. Steady-state was considered to have been achieved by two weeks, with GW501516 plasma concentration (Css min to Css max) ranging between 33 ng/mL to 88 ng/mL at the 2.5 mg dose and between 110 ng/mL to 327 ng/mL at the 10 mg dose. The Table shows lipid responses over the study period.

TABLE 1. Lipid Profiles and Percent Change From Baseline (CFB) Over the Course of Study by Treatment Group

Baseline (mmol/L)End of Study (mmol/L)Percent CFB
aP<0.01 for within group change; bP<0.01 vs Placebo; cP<0.05 for within group change; dP<0.05 vs Placebo.
2.5 mg3.74±0.173.71±0.200.7±3.4
10 mg4.39±0.234.04±0.18−7.1±3.4
2.5 mg2.38±0.142.45±0.182.5±0.02
10 mg2.96±0.222.68±0.18−7.6±5.8
2.5 mg1.04±0.061.06±0.081.8±2.6b
10 mg1.16±0.061.22±0.055.3±2.7b
2.5 mg0.93±0.080.77±0.07−15.4±6.5
10 mg1.04±0.150.76±0.09−23.4±7.0c
2.5 mg111±4107±4−3.2±1.6
10 mg116±4116±3−0.1±2.5d

TG levels declined significantly in the 2.5 mg (−15.4±6.5%, P=0.046) and 10 mg (−23.4±7.0%, P=0.01) treatment groups, while staying relatively stable in the placebo group (+6.1±10.2%, P=0.84). These TG responses in the treatment groups, however, were not significantly different than placebo after adjustment for baseline TG (P=0.16 for 2.5 mg; P=0.08 for 10 mg).

Contrary to TG, HDLc showed a significant decrease in the placebo group (−11.5±1.6%, P=0.002), while showing less movement in the two treatment groups (2.5 mg: 1.8±2.6%, P=0.43; 10 mg: 5.3±2.7%, P=0.12). However, compared with placebo, the HDLc response was significantly enhanced in both the 2.5 mg (P=0.004) and 10 mg (P<0.001) groups when adjusted for baseline HDL (Figure 1). The stability of ApoAI concentration for the 10 mg group represented a 7.7% improvement compared with placebo (P=0.03). Baseline systolic blood pressure values did not differ significantly among groups (Placebo=110±2 mm Hg, 2.5 mg=118±4 mm Hg, 10 mg=122±4 mmHG; P=0.16), nor did baseline diastolic blood pressure (Placebo=65±2 mmHG, 2.5 mg=63±2 mm Hg, 10 mg=68±2 mm Hg; P=0.17). There were no significant changes over the course of the study for systolic (Placebo P=0.10, 2.5 mg P=0.90, 10 mg P=0.81) or diastolic (Placebo P=0.48, 2.5 mg P=0.63, 10 mg P=0.70) blood pressures. Food intake and physical activity diaries were comparable in all groups of patients (data not shown).

Figure 1. Temporal changes in HDLc with GW501516 or placebo. Data are represented as the mean and 95% confidence interval (bars) for individuals in a treatment group on that day. Solid line: 2.5 mg; dotted line: 10 mg; dashed line: placebo.

Fat Feeding Study

Lipolytic activity was gauged with TG-AUC calculations after fat feeding (Figure 2). The placebo group showed an increase of 36.1±14.6% in TG-AUC from baseline to end, whereas the 2.5 mg group decreased 15.3±9.8% (P versus placebo=0.07), and the 10 mg group decreased 25.4±11.2% (P versus placebo=0.01). It should be noted that central tendency is difficult to estimate in the placebo group because 4 subjects experienced increases of greater than 40%, 1 had an increase of 13%, and 1 had a decrease of 24%. However, parametric and nonparametric statistical approaches yielded the same conclusions with regard to treatment effects. Change from baseline in TGmax showed similar patterns with an increase of 17.2±15.5% for the placebo group compared with −8.8±8.8% (P=0.09) for the 2.5 mg group and −13.4±7.8% (P=0.03) for the 10 mg group. Adjustments for baseline TG levels did not alter the conclusions.

Figure 2. Triglyceride response after high fat challenge at baseline and Day 13 by treatment. A, Placebo. B, 2.5 mg GW501516. C, 10 mg GW501516. ⋄ baseline; ▪ end point (Day 13). TG concentrations were measured over 24 hours after fat challenge at Days 0 and 13. TGmax and TG-AUC were calculated at each time. In the placebo group (A), upward arrows indicate increased TG values during a 24-hour period post-fat challenge at Days 0 and 13. In the two treatment groups (B and C), downward arrows indicate decreased TG values during a 24-hour period post-fat challenge at Days 0 and 13. Only the upper or lower halves of the standard error bars are shown to prevent obscuring the details of these figures.

The percentage of patients with decreases in TG-AUC follows a dose-response paradigm (P for trend=0.02). Moreover, the change in TG-AUC was significantly correlated to the change in serum TG for patients on active treatment (r=0.60, P=0.009). Although correlation analyses are prone to Type II error in small studies, neither baseline TG nor end TG correlated significantly with TG-AUC change in active treatment subjects. No significant correlations were noted in the placebo group.

Effects of GW501516 on Gene Expression and Fatty Acid Oxidation in Human Skeletal Muscle Cells

The mRNA levels of FAO and lipid handling genes were determined in HSKM cells treated with 10 nmol/L GW501516 (Figure 3). GW501516 induced CD36 mRNA expression approximately 2-fold (P=0.049) above the vehicle (Figure 3A). Further, the genes encoding CPT1a and CPT1b are both expressed in HSKM cells and both respond to GW501516. CPT1a was induced nearly 5-fold (P=0.005; Figure 3B) and CPT1b nearly 3-fold (P=0.02; Figure 3C) above the vehicle. We determined that although the pyruvate dehydrogenase kinase-4 (PDK4) gene is barely detectable in vehicle-treated HSKM cells, it is induced approximately 200-fold (P=0.003) in response to GW501516 (Figure 3D). To determine whether GW501516 regulates fatty acid combustion, a FAO assay was performed, revealing that GW501516 induced oleate oxidation approximately 7-fold (P<0.001) compared with vehicle (Figure 4). We routinely checked that the HSKM cells remained viable throughout the studies and that cell toxicity did not occur. No reduction in cell viability was noted (data not shown).

Figure 3. Effects of GW501516 on mRNA expression in HSKM cells. Cells were incubated with 10 nmol/L GW501516 and analyzed for gene expression by RTQ-PCR. Data were normalized to 36B4. Normalized values were an average of 3 biological replicates and presented as copies of target RNA. A, CD36; P=0.049. B, CPT1a; P=0.005. C, CPT1b; P=0.021. D, PDK4; P=0.003. E, ABCA1; P=0.002.

Figure 4. Effect of GW501516 on fatty acid oxidation in HSKM cells. Cells were incubated with 10 nmol/L GW501516 and analyzed for FAO using [14C]oleate as described in the Methods. The assay was performed in triplicate and data presented as mean oleate oxidation with SEM. P<0.0001.

Further, to determine whether genes involved in HDL formation and flux are regulated by GW501516, we measured the quantity of LpL, SR-B1, ABCG1, and ABCA1 mRNA transcripts. Whereas LpL, SR-B1 and ABCG1 were expressed in HSKM cells, none were regulated by GW501516 (data not shown). Also, GW501516 had no effect on mRNA expression of PPARα, PPARγ, or PPARδ (data not shown). In contrast, ABCA1 was induced approximately 2-fold (P=0.002) compared with vehicle (Figure 3E). Finally, enhanced cholesterol efflux (30%, P=0.001) was observed when GW501516 (100 nM) was added to cholesterol-loaded HSKM cells and whole serum was used as an acceptor (supplemental Figure I).


The PPARδ agonist-induced lipoprotein effects (eg, an increase in HDLc of up to 16.8%) were observed as the dose increased to 10 mg of GW501516. Closer inspection of the placebo cohort within this small trial revealed the physiologically expected reduction in HDLc (−11.5%, P=0.002) when requiring the study participants to be in-house and sedentary. Previous preclinical studies demonstrated that administration of GW501516 in obese rhesus monkeys at doses of 0.6 and 2.0 mg/kg/d produced a significant increase in HDLc and a reduction in TG values14 at similar GW501516 plasma exposure levels observed in our human experiment at daily doses of 2.5 mg and 10.0 mg, respectively. The most dramatic results observed in these animals were at the 6.0 mg/kg/d, with exposure level considerably above those observed in humans at the 10.0 mg dose. These pharmacodynamic (PD) results were recently corroborated by similar findings in Vervet monkeys administered with 3.0 mg/kg/d dose of GW501516, leading to over 3-fold higher exposure level than observed in human at the 10.0 mg dose.15 However, the pathophysiologic mechanisms for these changes remain unclear. The comparable HDL:TG modifications with both exercise and PPARδ agonism are conducive to theories surrounding lipolytic activity.

When placebo subjects were administered a fat meal, a substantial increase in post-fat feeding TG-AUC was observed after the 2 weeks of inactivity, consistent with a reduction in lipolytic activity,18 and in turn a reduction in tissue uptake of fat. Such a clinical assay has been well correlated with LpL activity assessed after heparin infusion.19 As such, these kinetic assays for LPL activity have been associated with circulating TG and HDLc concentrations.20 In contrast to the placebo group, the 2.5 mg and, significantly, the 10 mg dose of GW501516 reduced the post-fat feeding TG-AUC and maximum TG levels, suggesting amelioration of the lipolytic deficiency induced by a sedentary status. Reduction in lipolysis has been reported to be associated with lack of muscle movement.21 In rodent studies, there is a dramatic (10- to 20-fold) and rapid (initiating within 4 hours) decrease in LpL activity (with no change in mRNA) when muscle contraction is severely limited, particularly related to red-oxidative muscle.21 Stabilization of the HDL particle, through such lipolytic processing, leading to a reduced fractional catabolic rate is consistent with a longer HDL half-life reported in physically active men, and progressively higher apoAI levels in exercise-trained individuals of advancing physical activity groups.10,22–24

There are, however, other explanations for these observations. The in-house-provided low fat meals may have played a role in reducing HDLc within the placebo group. The positive association between saturated fat intake and HDLc is well documented.25,26 Azrolan et al27 have proposed that the hepatic-lipid cellular milieu may sequester translatable mRNA, and/or modify translation initiation sites, thereby depressing apoAI synthesis when the environment is saturated-fat deficient. Further, there are 3 alternate ABCA1 transcripts, and the expression of these transcripts is tissue-specific and varies in response to diet.28 Preclinical models suggest modest PPARδ activity in liver to counter these changes, notwithstanding secondary effects imposed from the periphery.29 In addition, PPARδ is expressed in the intestine and may have influenced fat absorption. Preclinical models have suggested a reduction in cholesterol absorption.30,31 However, whereas long chain fatty acid intestinal uptake may be modified,30 it has been reported that PPARδ activation does not affect total fat absorption in rodent models.31 Lack of a weight loss trend and no reported stool abnormalities also lowers the probability of this option. Yet, although we propose lipolysis as contributory to the HDL/TG effects, we cannot exclude other PPARδ-mediated mechanisms.

The association between PPARδ-induced TG reductions and TG-AUC values after a fat feeding in our study with active treatment is consistent with the lipolysis concept. In contrast, there was no association between TG-AUC and the observed HDLc benefits. Although these cohorts are small, and cannot be used to exclude a particular mechanistic pathway, these data could suggest that the PPARδ-mediated HDLc change does not depend on lipolysis to the same extent as that of changes in TG or requires an alternative explanation.

In both the published obese rhesus monkey14 and HSKM cell gene analyses reported herein, ABCA1 expression was found to be upregulated by a PPARδ agonist. ABCA1 protein-deficient subjects cannot properly lipidate lipid-poor apoAI particles, and are noted to have severely low HDLc values.32 PPARα and γ agonists upregulate ABCA1 gene expression.33 The extent to which peripheral PPARδ, (rather than hepatic) activation plays a role in enhancing ABCA1-mediated cholesterol efflux to HDL remains unclear.

Finally, changes in glucose tolerance (admittedly unlikely in healthy volunteer subjects) could have contributed to the observed TG and HDL changes. Lee et al and Tanaka et al have noted an improvement in insulin sensitivity when a PPARδ agonist, including GW501516, is administered to rodents.34,35 This is hypothesized to be associated with reductions in the pool of skeletal muscle fat, a feature relevant to insulin resistance.36 In our 2-week clinical-safety trial presented here, insulin and glucose were not measured. However, if insulin sensitization did occur, an insulin-dependent impact on lipolysis18 as well as on hepatic synthesis37 might have contributed to the lipid trends observed.

Upregulation in FAO is observed in HSKM cell lines exposed to GW501516, demonstrated in this report, with induction of fatty acid catabolism genes, eg, CD36/FAT, CPT1a, and CPT1b. This is the first quantitative analysis of fatty acid catabolism genes in HSKM cells and corroborates similar changes in preclinical models provided PPARδ agonists.29,38,39 FAO upregulation was achieved in vitro with GW501516 concentrations far below the plasma levels observed in the clinical study. Nonetheless, free concentration circulating levels (available to the target tissues) are expected to be much lower because of the high binding of GW501516 to plasma proteins (data not shown). The parallel documented increase in FAO with human exercise40,41 and associated upregulation of the binding and transport fatty acid proteins (CD36/FAT, FABP),42 mitochondrial carrier protein CPT1,43 and the counter-regulatory PDK444,45 in human muscle biopsy tissue prompts speculation that exercise-induced FAO is mechanistically generated via a PPARδ physiological switch.11 Such a concept is consistent with upregulation of PPARδ shortly after exercise initiation in humans.46,47

There are caveats to the outcomes presented in this report. The increase in HDLc of 16.8% at the 10 mg dose is predicated on a reduction in placebo of 11%. Until larger studies and/or those of longer duration are performed with stable outpatient placebo populations, the actual HDLc benefits cannot be defined. Further, although nursing logs and protocol design requirements defined a sedentary bed-ridden existence for 2 weeks, we did not directly quantify physical activity. We do know, however, that these individuals did not leave the small research unit within the study time interval of two weeks. In addition, diet-log estimates of fat intake (26% of total calories) were assumed to be lower than baseline, even though the entry dietary patterns were not interrogated. Although this study does not perfectly support an exercise pathway, it does provide insight toward HDLc values in the resting state. To investigate thoroughly the impact of GW501516 treatment on HDL levels in the setting of exercise, both resting and nonresting subjects need to be evaluated in parallel and muscle biopsies evaluated for relevant genes of interest. However, this small study was designed to study the effects of a repeat-dose of GW501516 in healthy subjects. Additional studies that address these issues are planned.

In conclusion, the results of this study provide support for the influence of PPARδ on the enzymes critical to FAO in human cells, comparable to those observed in murine and monkey models, and similar to that observed with exercise. Further, HDLc and TG responses observed in this first use of a PPARδ agonist in human subjects were similar to lipid responses in preclinical models. We hypothesize that GW501516-induced enhancement of fatty-acid utilization contributes to these lipoprotein changes.

Original received June 26, 2006; final version accepted October 11, 2006.


All authors have employment and ownership interest in GlaxoSmithKline.


Correspondence to Dennis L. Sprecher, MD, GlaxoSmithKline, Department of Discovery Medicine – Dyslipidemia, 709 Swedeland Road, UW2301, King of Prussia, PA 19406. E-mail


  • 1 Willson TM, Brown PJ, Sternbach DD, Henke BR. The PPARs: From orphan receptors to drug discovery [Review]. J Med Chem. 2000; 43: 527–550.CrossrefMedlineGoogle Scholar
  • 2 Berger J, Moller DE. The mechanisms of action of PPARs [Review]. Ann Rev Med. 2002; 53: 409–435.CrossrefMedlineGoogle Scholar
  • 3 Picard F, Kurtev M, Chung NJ, Topark-Ngarm A, Senawong T, de Oliveira RM, Leid M, McBurney MW, Guarente L. Sirt1 promotes fat mobilization in white adipocytes by repressing PPAR-gamma. Nature. 2004; 429: 771–776.CrossrefMedlineGoogle Scholar
  • 4 Fabris R, Nisoli E, Lombardi AM, Tonello C, Serra R, Granzotto M, Cusin I, Bohner-Jeanrenaud F, Federspil G, Carruba MO, Vettor R. Preferential channeling of energy fuels toward fat rather than muscle during high free fatty acid availability in rats. Diabetes. 2001; 50: 601–608.CrossrefMedlineGoogle Scholar
  • 5 Muoio DM, MacLean PS, Lang DB, Li S, Houmard JA, Way JM, Winegar DA, Corton JC, Dohm GL, Kraus WE. Fatty acid homeostasis and induction of lipid regulatory genes in skeletal muscles of peroxisome proliferator-activated receptor (PPAR) alpha knock-out mice - Evidence for compensatory regulation by PPAR delta. J Biol Chem. 2002; 277: 26089–26097.CrossrefMedlineGoogle Scholar
  • 6 Lee CH, Olson P, Evans RM. Minireview: Lipid metabolism, metabolic diseases, and peroxisome proliferator-activated receptors [Review]. Endocrinol. 2003; 144: 2201–2207.CrossrefMedlineGoogle Scholar
  • 7 Luquet S, Lopez-Soriano J, Holst D, Fredenrich A, Melki J, Rassoulzadegan M, Grimaldi P. Peroxisome proliferator-activated receptor delta controls muscle development and oxidative capability. FASEB J. 2003; 17: 2299–2301.CrossrefMedlineGoogle Scholar
  • 8 Varady KA, Jones PJH. Combination diet and exercise interventions for the treatment of dyslipidemia: an effective preliminary strategy to lower cholesterol levels? [Review]. J Nutr. 2005; 135: 1829–1835.CrossrefMedlineGoogle Scholar
  • 9 Suzuki Y, Kawakubo K, Makita Y, Goto S, Gunji A 20 Days Bed-rest on Insulin Action in Relation with Body Fat in Young Subjects. Physiologist. 1993; 36 [Supplement 1]: S153–S154.Abstract.MedlineGoogle Scholar
  • 10 Tikkanen HO, Hamalainen E, Harkonen M. Significance of skeletal muscle properties on fitness, long-term physical training and serum lipids. Atherosclerosis. 1999; 142: 367–378.CrossrefMedlineGoogle Scholar
  • 11 Wang YX, Zhang CL, Yu RT, Cho HK, Nelson MC. Regulation of muscle fiber type and running endurance by PPAR delta (vol 2, pg e61, 2004). Plos Biology. 2005; 3: 177.CrossrefGoogle Scholar
  • 12 Wei ZL, Kozikowski AP. A short and efficient synthesis of the pharmacological research tool GW501516 for the peroxisome proliferator-activated receptor delta. J Organ Chem. 2003; 68: 9116–9118.CrossrefMedlineGoogle Scholar
  • 13 Sznaidman ML, Haffner CD, Maloney PR, Fivush A, Chao E, Goreham D, Sierra ML, LeGrumelec C, Xu HE, Montana VG, Lambert MH, Willson TM, Oliver WR, Sternbach DD. Novel selective small molecule agonists for peroxisome proliferator-activated receptor delta (PPAR delta) - Synthesis and biological activity. Bioorg Medic Chem Lett. 2003; 13: 1517–1521.CrossrefMedlineGoogle Scholar
  • 14 Oliver WR, Shenk JL, Snaith MR, Russell CS, Plunket KD, Bodkin NL, Lewis MC, Winegar DA, Sznaidman ML, Lambert MH, Xu HE, Sternbach DD, Kliewer SA, Hansen BC, Willson TM. A selective peroxisome proliferator-activated receptor delta agonist promotes reverse cholesterol transport. Proc Natl Acad Sci U S A. 2001; 98: 5306–5311.CrossrefMedlineGoogle Scholar
  • 15 Wallace JM, Schwarz M, Coward P, Houze J, Sawyer JK, Kelley KL, Chai A, Rudel LL. Effects of peroxisome proliferator-activated receptor alpha/delta agonists on HDL-cholesterol in vervet monkeys. J Lipid Res. 2005; 46: 1009–1016.CrossrefMedlineGoogle Scholar
  • 16 Kim JY, Hickner RC, Cortright RL, Dohm GL, Houmard JA. Lipid oxidation is reduced in obese human skeletal muscle. Am J Physiol. 2000; 279: E1039–E1044.CrossrefMedlineGoogle Scholar
  • 17 Maglich JM, Parks DJ, Moore LB, Collins JL, Goodwin B, Billin AN, Stoltz CA, Kliewer SA, Lambert MH, Willson TM, Moore JT. Identification of a novel human constitutive androstane receptor (CAR) agonist and its use in the identification of CAR target genes. J Biol Chem. 2003; 278: 17277–17283.CrossrefMedlineGoogle Scholar
  • 18 Seip RL, Angelopoulos TJ, Semenkovich CF Exercise induces human lipoprotein lipase gene expression in skeletal muscle but not adipose tissue. Am J Physiol. 1995; 31: E.Google Scholar
  • 19 Nilsson-Ehle P, Schotz M. A stable, radioactive, substrate emulsion for assay of lipoprotein lipase. J Lipid Res. 1976; 17: 536–541.CrossrefMedlineGoogle Scholar
  • 20 Goldberg IJ, Vanni-Reyes T, Ramakrishnan S, Holleran S, Ginsberg HN. Circulating lipoprotein profiles are modulated differently by lipoprotein lipase in obese humans. J Cardiovasc Risk. 2000; 7: 41–47.CrossrefMedlineGoogle Scholar
  • 21 Hamilton M, Hamilton D, Zderic T. Exercise physiology versus inactivity physiology: an essential concept for understanding lipoprotein lipase regulation. Exerc Sport Sci Revs. 2004; 32: 161–166.CrossrefMedlineGoogle Scholar
  • 22 Otarod J, Goldberg I. Lipoprotein lipase and its role in regulation of plasma lipoproteins and cardiac risk. Curr Atheroscler Rep. 2004; 6: 335–342.CrossrefMedlineGoogle Scholar
  • 23 Barter PJ, Rye KA. High density lipoproteins and coronary heart disease [Review]. Atherosclerosis. 1996; 121: 1–12.CrossrefMedlineGoogle Scholar
  • 24 Nofer JR, Kehrel B, Fobker M, Levkau B, Assmann G, von Eckardstein A. HDL and arteriosclerosis: beyond reverse cholesterol transport [Review]. Atherosclerosis. 2002; 161: 1–16.CrossrefMedlineGoogle Scholar
  • 25 Jones PJH, Lichtenstein AH, Schaefer EJ, Namchuk GL. Effect of dietary fat selection on plasma cholesterol synthesis in older, moderately hypercholesterolemic humans. Atheroscler Thromb. 1994; 14: 542–548.CrossrefMedlineGoogle Scholar
  • 26 Cox CMA, Sutherland WHF, Ball MJ, Mann JI. Response of plasma lathosterol concentration to change in the quality of dietary fat in men and women. Eur J Clin Nutr. 1996; 50: 358–363.MedlineGoogle Scholar
  • 27 Azrolan N, Odaka H, Breslow JL, Fisher EA. Dietary fat elevates hepatic apoA-I production by increasing the fraction of apolipoprotein a-i mrna in the translating pool. J Biol Chem. 1995; 270: 19833–19838.CrossrefMedlineGoogle Scholar
  • 28 Singaraja RR, James ER, Crim J, Visscher H, Chatterjee A, Hayden MR. Alternate transcripts expressed in response to diet reflect tissue-specific regulation of ABCA1. J Lipid Res. 2005; 46: 2061–2071.CrossrefMedlineGoogle Scholar
  • 29 Holst D, Luquet S, Nogueira V, Kristiansen K, Leverve X, Grimaldi PA. Nutritional regulation and role of peroxisome proliferator-activated receptor delta in fatty acid catabolism in skeletal muscle. Biochim Biophys Acta. 2003; 1633: 43–50.CrossrefMedlineGoogle Scholar
  • 30 Poirer H, Niot I, Monnot MC, Braissant O, Meunier-Durmort C, Costet P, Pineau T, Wahl W, Willson TM, Besnard P. Differential involvement of peroxisome-proliferator-activated receptors alpha and delta in fibrate and fatty-acid-mediated inductions of the gene encoding liver fatty-acid-binding protein in the liver and the small intestine. Biochem J. 2001; 355: 481–488.CrossrefMedlineGoogle Scholar
  • 31 van der Veen JN, Kruit JK, Havinga R, Baller JFW, Chimini G, Lestavel S, Staels B, Groot PHE, Groen AK, Kuipers F. Reduced cholesterol absorption upon PPAR delta activation coincides with decreased intestinal expression of NPC1L1. J Lipid Res. 2005; 46: 526–534.CrossrefMedlineGoogle Scholar
  • 32 Attie AD, Kastelein JP, Hayden MR. Pivotal role of ABCA1 in reverse cholesterol transport influencing HDL levels and susceptibility to atherosclerosis [Review]. J Lipid Res. 2001; 42: 1717–1726.CrossrefMedlineGoogle Scholar
  • 33 Chinetti G, Lestavel S, Bocher V, Remaley AT, Neve B, Torra IP, Teissier E, Minnich A, Jaye M, Duverger N, Brewer HB, Fruchart JC, Clavey V, Staels B. PPAR-alpha and PPAR-gamma activators induce cholesterol removal from human macrophage foam cells through stimulation of the ABCA1 pathway. Nat Med. 2001; 7: 53–58.CrossrefMedlineGoogle Scholar
  • 34 Lee CH, Olson P, Hevener A, Mehl I, Chong LW, Olefsky JM, Gonzalez FJ, Ham J, Kang H, Peters JM, Evans RM. PPAR delta regulates glucose metabolism and insulin sensitivity. Proc Natl Acad Sci U S A. 2006; 103: 3444–3449.CrossrefMedlineGoogle Scholar
  • 35 Tanaka T, Yamamoto J, Iwasaki S, Asaba H, Hamura H, Ikeda Y, Watanabe M, Magoori K, Ioka RX, Tachibana K, Watanabe Y, Uchiyama Y, Sumi K, Iguchi H, Ito S, Doi T, Hamakubo T, Naito M, Auwerx J, Yanagisawa M, Kodama T, Sakai J. Activation of peroxisome proliferator-activated receptor delta induces fatty acid beta-oxidation in skeletal muscle and attenuates metabolic syndrome. Proc Natl Acad Sci U S A. 2003; 100: 15924–15929.CrossrefMedlineGoogle Scholar
  • 36 Petersen KF, Shulman GI Etiology of insulin resistance. Am J Med. 2006; 119: 10S–16S.CrossrefMedlineGoogle Scholar
  • 37 Lewis GF, Rader DJ. New insights into the regulation of HDL metabolism and reverse cholesterol transport [Review]. Circ Res. 2005; 96: 1221–1232.LinkGoogle Scholar
  • 38 Dressel U, Allen TL, Pippal JB, Rohde PR, Lau P, Muscat GEO. The peroxisome proliferator-activated receptor beta/delta agonist, GW501516, regulates the expression of genes involved in lipid catabolism and energy uncoupling in skeletal muscle cells. Mol Endocrinol. 2003; 17: 2477–2493.CrossrefMedlineGoogle Scholar
  • 39 Kramer DK, Al Khalili L, Perrini S, Skogsberg J, Wretenberg P, Kannisto K, Wallberg-Henriksson H, Ehrenborg E, Zierath JR, Krook A. Direct activation of glucose transport in primary human myotubes after activation of peroxisome proliferator-activated receptor delta. Diabetes. 2005; 54: 1157–1163.CrossrefMedlineGoogle Scholar
  • 40 Spriet LL. Regulation of skeletal muscle fat oxidation during exercise in humans. Med Sci Sports Exerc. 2002; 34: 1477–1484.CrossrefMedlineGoogle Scholar
  • 41 Turcotte L. Muscle fatty acid uptake during exercise: possible mechanisms. Exerc Sports Sci Revs. 2000; 28: 4–9.MedlineGoogle Scholar
  • 42 Kiens B, Kristiansen S, Jensen P, Richter EA, Turcotte LP. Membrane associated fatty acid binding protein (Fabppm) In human skeletal muscle is increased by endurance training. Biochem Biophys Res Commun. 1997; 231: 463–465.CrossrefMedlineGoogle Scholar
  • 43 Kim JK, Hickner RC, Dohm GL, Houmard JA. Long- and medium-chain fatty acid oxidation is increased in exercise-trained human skeletal muscle. Metabol Clin Exper. 2002; 51: 460–464.CrossrefMedlineGoogle Scholar
  • 44 Pilegaard H, Ordway GA, Saltin B, Neufer PD. Transcriptional regulation of gene expression in human skeletal muscle during recovery from exercise. Am J Phyisol. 2000; 279: E806–E814.CrossrefMedlineGoogle Scholar
  • 45 Tunstall RJ, Mehan KA, Wadley GD, Collier GR, Bonen A, Hargreaves M, Cameron-Smith D. Exercise training increases lipid metabolism gene expression in human skeletal muscle. Am J Physiol. 2002; 283: E66–E72.CrossrefGoogle Scholar
  • 46 Russell AP, Hesselink MKC, Lo SK, Schrauwen P Regulation of metabolic transcriptional co-activators and transcription factors with acute exercise. FASEB J. 2005; 19: NIL.Google Scholar
  • 47 Watt MJ, Southgate RJ, Holmes AG, Febbraio MA. Suppression of plasma free fatty acids upregulates peroxisome proliferator-activated receptor (PPAR) alpha and delta and PPAR coactivator 1 alpha in human skeletal muscle, but not lipid regulatory genes. J Mol Endocrinol. 2004; 33: 533–544.CrossrefMedlineGoogle Scholar