Institute for Molecular Bioscience, The University of Queensland, St. Lucia, Queensland 4072, Australia
Address all correspondence and requests for reprints to: George Muscat, Institute Molecular Bioscience, St. Lucia, Queensland 4072, Australia. E-mail: G.Muscat{at}imb.uq.edu.au.
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Research demonstrates the evolution of a multi-layered autoregulated system involving nuclear hormone receptors (NRs) for sensing and metabolizing biologically active lipids. NRs involved in control of lipid and cholesterol homeostasis, include the liver X receptors (LXRs), farnesoid X receptor, peroxisome proliferator-activated receptors (PPARs) , -ß/
, and -
[NR1C1, -2, -3, respectively (2)] liver receptor homolog-1 and the small heterodimeric partner (3, 4).
PPARs regulate the transcription of genes involved in lipid homeostasis, carbohydrate metabolism, energy expenditure and reverse cholesterol transport in a subtype- and tissue-specific manner. They are activated by a wide range of dietary factors, including saturated and unsaturated fatty acids (FAs), oxidized FA metabolites derived through the lipoxygenase and cyclo-oxygenase pathways, and selective synthetic compounds (e.g. hypolipidemic fibrates, and antidiabetic thiazolidinediones). From the viewpoint of PPARß/, the putative natural agonists are prostanoids, which are produced by the regulated conversion of poly-unsaturated FAs. In addition, PPAR
, -ß/
, and -
form obligate and permissive heterodimers with the retinoid X receptors (RXR) that can also be activated by the RXR agonists 9-cis-retinoic acid, and/or specific synthetic agonists called rexinoids (e.g. LG101305).
PPAR and -
are predominantly expressed in liver and adipose tissue, respectively. The expression of PPARß/
is ubiquitous. Moreover, it is very abundantly expressed in brain, intestine, skeletal muscle, spleen, macrophages, lung, and adrenals (5, 6, 7). Mouse transgenic, knockout, and knock-in studies coupled to pharmacological investigations have exposed the discrete physiological functions of the PPAR
and -
isoforms in lipid and carbohydrate metabolism. For example, PPAR
promotes adipogenesis and increases lipid storage. In contrast, PPAR
enhances the conflicting process of lipid catabolism/FA oxidation in the liver (5, 6). These physiological functions correlate with the hypolipidemic and antidiabetic (type II) effects of the synthetic and selective fibrate and glitazone drugs, which activate PPAR
and PPAR
, respectively.
Relatively less is known about PPARß/, which has been implicated in bone and fat metabolism (8, 9, 10). Recently, the potent, synthetic and selective PPARß/
agonist, GW501516, a phenoxyacetic acid derivative, has been reported (11). It was demonstrated that the triglyceride component of native very low-density lipoproteins (VLDLs) activate PPARß/
. GW501516 corrects hyperinsulinemia in insulin-resistant and obese primates. Furthermore, it raises ABCA1 mRNA expression, and serum HDL cholesterol, while lowering triglycerides. However, PPARß/
agonists also promote lipid absorption and storage in macrophages. Moreover, serum apolipoprotein (Apo) CIII levels and total cholesterol are raised. Hence, the overall effect of PPARß/
agonists on whole body cholesterol homeostasis, lipid metabolism, target tissues and mode of action remain unclear (10).
The PPAR-mediated FA oxidation in the liver plays a major role in ketosis that supports fuel requirements during fasting. Similar but distinct mechanisms must exist within peripheral tissues to implement localized responses to energy requirements and burdens in these tissues. For example, one would hypothesize that a PPAR
knockout would have major consequences on skeletal muscle fuel metabolism and gene expression. However, Muoio et al. (12) observed the skeletal muscle metabolic/ß-oxidation phenotype was not compromised in PPAR
-/- mice, in contrast to the dramatic deleterious effects in liver and heart tissue. A plausible hypothesis suggests that PPARß/
regulates fuel metabolism in skeletal muscle, a major mass peripheral tissue that accounts for 40% of the total body mass.
Skeletal muscle is one of the most metabolically demanding tissues that relies heavily on FAs as an energy source. PPARß/ is the most abundant PPAR in muscle tissue (12, 13, 14). It was first implicated in FA metabolism from studies using the knockout animals. Most PPARß/
-/- embryos die at an early stage due to a placental defect, the small number that survive exhibit a reduction in fat mass/adiposity (8, 15). However, this phenotype is absent in an adipocyte-specific PPARß/
knockout model, suggesting a complex autonomous action regulating systemic lipid metabolism (15, 16). This idea was further strengthened by the observation that treatment with the synthetic compound GW501516 in insulin resistant primates dramatically improves the serum lipid profile, and improves hyperinsulinemia. However, it is unclear which tissue is the major target for this activity. The classification of PPARß/
as sensor of dietary triglyceride in native VLDLs released by lipoprotein lipase (LPL) activity suggests skeletal muscle is a potential target tissue (17, 18). In addition, exercise and/or starvation induced up-regulation of FA oxidation genes in muscle remains intact in PPAR
-/- mice.
Muscle is a major site of glucose metabolism and FA oxidation. Furthermore, it is an important regulator of cholesterol homeostasis and HDL levels (19). Consequently, it has a significant role in insulin sensitivity, the blood lipid profile, and lipid metabolism. This underscores the need to define the contribution of this major mass tissue to PPARß/. Surprisingly, the fundamental role of PPARß/
in skeletal muscle cholesterol, lipid, glucose, and energy homeostasis has not been examined. Correspondingly, the objective of this study is to examine the functional role of PPARß/
in skeletal muscle, and to investigate the genes and regulatory genetic programs activated by PPARß/
involved in the control of lipid and energy homeostasis.
In summary, we demonstrate that PPARß/ directly and/or indirectly regulates genes involved in triglyceride-hydrolysis and FA oxidation [LPL, acyl-coenzyme A (CoA) synthetase 4 (ACS4), carnitine-palmitoyl-transferase (CPT1)], preferential lipid utilization (PDK4), energy expenditure [uncoupling protein (UCP)-1, -2, and -3], and lipid efflux (ABCA1/G1). Furthermore, we show that the muscle carnitine-palmitoyl-transferase-1 (M-CPT1) is directly regulated by PPARß/
in skeletal muscle, in a PPAR
coactivator-1 (PGC-1)-dependent manner. In summary, we show that PPARß/
activates the entire cascade of gene expression involved in lipid-uptake to FA oxidation, and in addition, activates the UCPs, thereby uncoupling oxidation from the production of energy, and increasing energy expenditure and thermogenesis. This provides the molecular basis for the lipid lowering effects of PPARß/
agonists previously described in obese primates (11), and we speculate that PPARß/
agonists would have therapeutic utility against a high-fat diet and obesity.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Subsequently, we examined the ability of this agonist to activate the different PPARß/-constructs in CV1 (data not shown) and C2C12 cells (Fig. 1B
). We cotransfected the various GAL4-PPARß/
constructs (full length, LBD, AF1) together with the G5E1b-LUC reporter into CV1 (data not shown), and C2C12-cells (Fig. 1B
), in the presence or absence of GW501516 (1 µM). The AF1-domain of PPARß/
inefficiently activated the LUC reporter, and did not respond to agonist treatment. In contrast, both GAL4-PPARß/
, and GAL4-PPARß/
-LBD-transactivated gene expression in an efficacious and agonist-dependent manner, in muscle (Fig. 1B
) and nonmuscle cells (data not shown). Similar transactivation patterns were observed when using another GAL4-dependent reporter construct, tkMH100-LUC, which utilizes the thymidine kinase (tk)-promoter backbone (20) instead of E1b (data not shown).
Subsequently, we examined the ability of the PPARß/ agonist GW501516 to activate a PPAR-dependent reporter (PPRE) in muscle cells. Moreover we examined the ability of the cofactors PGC-1, p300 and SRC-2/GRIP-1 to coactivate GW501516 dependent activation of gene expression in skeletal muscle C2C12 cells. We used the PPRE-tk-LUC reporter that contains three copies of a consensus binding site cloned upstream of the heterologous herpes simplex virus tk promoter linked to the LUC reporter gene. Furthermore, these experiments were performed in the absence of exogenous/ectopic PPARß/
expression vector (because these cells contain endogenous PPARß/
). As shown in Fig. 2A
the PPARß/
agonist GW501516 activated the expression of the PPRE-containing reporter approximately 2-fold in skeletal muscle C2C12 cells. No response was observed when the tk-LUC-backbone, lacking the PPRE, was used (data not shown). Furthermore, GW501516-dependent PPRE activation was enhanced when PGC-1, relative to p300 and SRC-2/GRIP-1, was cotransfected. For example, GW501516 activated the expression of the PPRE-reporter approximately 2.5-fold, and approximately 4-fold in the presence of the RXR agonist.
|
In addition, we further examined the ability of PPAR and PPAR
agonists (fenofibrate and rosiglitazone, respectively) to regulate PPAR-dependent PPRE-containing reporter in muscle cells (Fig. 2C
) and nonmuscle CV1 cells (data not shown) in the presence and absence of the coactivators PGC-1, p300, and SRC-2/GRIP-1. These experiments were performed to demonstrate these agonists regulate gene expression in skeletal muscle cells, as we subsequently wished to compare the relative effects of PPAR
, -ß/
, and -
agonists on the expression of the genes involved in skeletal muscle lipid and carbohydrate metabolism. Figure 2
, C and D, clearly demonstrates that PPAR
and -
agonists efficiently regulate gene expression in skeletal muscle cells. In addition, we show that SRC-2/GRIP-1 and PGC-1 selectively coactivate PPAR
and -
, respectively, in skeletal muscle cells (Fig. 2
, C and D). Finally, these experiments demonstrate that the C2C12 skeletal muscle cells express functional PPAR
, -ß/
, and -
receptors that support the activation of PPAR-dependent gene expression by selective agonists.
In summary, we demonstrate that PGC-1 expression in C2C12 cells selectively coactivates GW501516 and Rosiglitazone mediated activation of the PPAR-dependent PPRE. The selective coactivation of PPRE expression by cofactors in the presence of the selective PPAR agonists in the skeletal muscle cells was performed in the absence of exogenous/ectopic (high level) receptor expression, and provides an unbiased demonstration of cofactor selectivity, and receptor functionality in skeletal muscle cells. Clearly, PGC-1 expression in skeletal muscle cells increases GW501516 and Rosiglitazone inducibility, and the absolute level of PPRE-dependent expression. In contrast, SRC-2/GRIP1 expression preferentially increases Fenofibrate-mediated activation.
Regulation of Gene Expression in Skeletal Muscle Cells by PPAR, -ß/
, and -
Agonists
We investigated the expression of the genes involved in skeletal muscle lipid and carbohydrate metabolism (see Table 1) in the presence and absence of the PPAR
, -ß/
, and -
agonists. We undertook these studies in the C2C12 skeletal muscle cell culture model. In this system, proliferating C2C12 skeletal myoblasts differentiate into post-mitotic multinucleated myotubes that acquire a muscle-specific, contractile phenotype. This in vitro system has been used to investigate the regulation of cholesterol homeostasis and lipid metabolism by LXR agonists (19). Muscat et al. demonstrated that the selective and synthetic LXR agonist, T0901317 induced similar effects on mRNAs encoding ABCA1/G1, ApoE, stearoyl CoA desaturase (SCD-1), SREBP-1c, etc. in differentiated C2C12-myotubes and Mus musculus quadriceps skeletal muscle tissue. The physiological validation of the cell culture model in the mouse corroborates the utility of this model system. This evidence, coupled to the flexibility and utility of cell culture in terms of cost, agonist treatment, RNA extraction, and target validation provides an ideal platform to identify the PPARß/
-dependent regulation of metabolism. In addition, and more importantly this cell line (16, 21, 22, 23) and other rodent skeletal muscle cell lines (13, 14) have been demonstrated to express functional PPAR
, -ß/
, and -
receptors. Our quantitative real time analysis in Fig. 3C
verifies the published reports that the PPAR mRNAs are expressed in skeletal muscle C2C12 cells. Our analysis demonstrates PPAR
and ß/
are expressed at similar levels in 96 h differentiated myotube cells. PPAR
mRNA is abundantly expressed, however, the primers reflect mRNA expression from all three PPAR
isotypes (i.e.
1 +
2 +
3).
|
|
Subsequently, we used quantitative real-time PCR to investigate the expression pattern of genes involved in lipid/cholesterol absorption (CD36/FAT, FABP3; Fig. 4A), lipogenesis (SREBP-1c, FAS, SCD-1 and -2; Fig. 4B
), triglyceride hydrolysis, and FA oxidation (LPL, M-CPT1, ACS4; Fig. 4C
), glucose/fructose absorbtion and utilization (Glut-4, and -5; Fig. 4D
, PDK-2 and -4; Fig. 4E
), lipid efflux (ABCA1 and -G1, ApoE; Fig. 4F
), energy expenditure (UCP-1, -2, and -3; Fig. 4G
), and glucose and lipid storage (Glycogenin1/GYG1, adipophilin/ADRP; Fig. 4H
).
|
Other candidate mRNAs that showed a modest, but significant increase in expression (2-fold) upon treatment with the GW501516 were FABP3 (lipid uptake; Fig. 4A
), LPL and M-CPT1 (triglyceride-hydrolysis and FA oxidation, respectively; Fig. 4C
), UCP-3 (another member of the UCP family, involved in energy expenditure; Fig. 4G
), and ADRP (lipid storage; Fig. 4H
). All of these mRNAs also responded to treatment with the RXR agonist, LG101305, and were synergistically activated upon treatment with both agonists. Noteworthy, the level of mRNA encoding for UCP-2 was only marginally activated by LG101305, when compared with treatment with the PPARß/
agonist. ADRP mRNA expression was also induced by rosiglitazone treatment.
The increase in mRNA expression level subsequent to treatment with both agonists was also observed with a number of candidate target genes investigated. In the context of lipid and FA uptake, we observed a 6-fold increase in the mRNA encoding FABP3, and a 2-fold increase in CD36 (Fig. 4A). Some regulators and markers of lipogenesis (SREBP-1c, SCD-1 and -2; Fig. 4B
) were relatively refractory to treatment with one of the agonists, but showed significant induction after cotreatment (
2- to 3-fold). Interestingly, FAS was reproducibly repressed upon treatment with the PPARß/
agonist. In contrast, rosiglitazone increased FAS mRNA expression approximately 2-fold. Interestingly, SREBP-1c induction did not result in the induction of the downstream targets, FAS, SCD-1 and -2. In muscle, PPARß/
activation of SREB1c may be uncoupled from FA metabolism, similar to the uncoupling of LXR activity and FA metabolism observed in quadricep tissue (19).
The transcripts encoding LPL, M-CPT1, ACS4, and PDK4 that are involved in triglyceride-hydrolysis, FA oxidation and preferential fuel utilization were induced approximately 7-, 4-, 3-, and 7-fold by cotreatment, respectively (Fig. 4C). The significance of the synergistic activation of LPL and CPT1 by cotreatment with the PPARß/
and RXR agonists, are highlighted by the observation that the cotreatment with agonists for PPAR
and -
in the presence of an RXR agonist does not induce LPL and CPT1 expression (Fig. 4C
). Interestingly, PDK4 mRNA was activated by PPAR
, -ß/
, and -
agonists. The glucose, and fructose transporters (Glut-4 and -5) were induced by rexinoid treatment, but completely refractory to the PPARß/
agonist (Fig. 4D
). In contrast, we observed Glut-4, and -5 were activated by PPAR
and PPAR
agonists, respectively.
As mentioned earlier, cotreatment with agonists for PPARß/ and RXR led to a dramatic increase in the level of mRNAs encoding the UCPs that regulate energy expenditure. UCP-1, -2, and -3 were activated approximately 23-, 8-, and 16-fold, respectively (Fig. 4G
). The significance, and specificity of the UCP-1 to -3 response by cotreatment with PPARß/
and RXR agonists, are further highlighted by the observation that the cotreatment with the agonists for PPAR
and -
in the presence of an RXR agonist relatively poorly induced UCP-2 and -3 mRNA expression (Fig. 4G
).
We also examined the expression of genes involved in lipid efflux, lipid storage, and glycogen deposition (Fig. 4F). ABCA1 mRNA was synergistically induced approximately 11-fold by cotreatment with both agonists, ABCG1 approximately 4-fold. ApoE was induced by rexinoid treatment, but refractory to GW501516. Cotreatment did not lead to further activation. Finally ADRP/adipophilin was synergistically induced approximately 7-fold by PPARß/
and RXR agonist cotreatment. Furthermore, ADRP mRNA increased approximately 2- to 3-fold to treatment with PPARß/
and -
, and the RXR agonists alone (Fig. 4H
). The observed synergistic effect of cotreatment by both agonists is consistent with the fact that PPARs bind to DNA as heterodimers with RXR. Glycogenin was only induced approximately 2-fold by cotreatment. However, fenofibrate treatment induced glycogenin-1 mRNA levels approximately 4-fold.
To verify some of the results obtained from real-time PCR analysis, we performed Northern blot analysis using RNA extracted from C2C12-myotubes differentiated for 96 h and subsequently treated for 24 h with PPARß/ and/or RXR agonists (Fig. 5
, A and B). These results unconditionally confirm that the mRNAs of UCP-2/-3 and LPL are induced upon treatment with GW501516, and validate the real-time PCR analysis. Figure 5B
also demonstrates that the activation of UCP-2 occurs after a short time, such as 4 h.
|
|
|
In summary, we observed that the PPARß/ agonist GW101516 dramatically activates the mRNAs encoding the UCPs, suggesting that PPARß/
has an important role in energy uncoupling. Furthermore it activates the expression of genes involved in preferential lipid utilization, FA catabolism, and energy expenditure. Interestingly, fenofibrate induces genes involved in fructose uptake, and glycogen formation in skeletal muscle. In contrast, rosiglitazone-mediated activation of PPAR
induces gene expression associated with glucose uptake, FA synthesis and lipid storage. This demonstrates that PPARs have distinct, complementary, and opposing roles in skeletal muscle.
M-CPT1 Is a Primary Target of PPARß/ in Skeletal Muscle
We further explored the molecular basis of PPARß/-mediated gene activation in skeletal muscle cells by evaluating whether direct, or indirect mechanisms mediated the observed increase in mRNA levels. We investigated whether the promoters of selected target genes were active in skeletal muscle cells, and tested the responsiveness of the promoters to PPARß/
and RXR agonists in a cell-based reporter assay. Because the promoters were introduced into skeletal muscle cells in the absence of exogenous receptors, the ligand-dependent responses reflect the functional properties of the endogenous receptors.
We transiently transfected C2C12 cells with the regulatory sequences of selected target genes that were accessible to us, including ABCA1 (19), CD36/FAT (25), LPL (26), M-CPT1 (27), SREBP-1c (19), and UCP-2 (28), cloned in front of the pGL2/3-basic LUC backbone and examined the response after treatment with GW501516 and/or LG101305. Interestingly, only the M-CPT1 promoter responded to the PPARß/ agonist and was further activated by cotreatment with both agonists (Fig. 8A
). All other promoters tested, even though active in C2C12-skeletal muscle cells, did not respond to treatment with the PPARß/
agonist, GW501516 (data not shown).
|
We then investigated the regulation of the M-CPT1 promoter in CV1 cells, which do not endogenously express PPARß/. The M-CPT1 promoter was cotransfected into CV1 cells with PPARß/
, PGC-1, or both together. As seen in Fig. 8C
, PPARß/
activated the M-CPT1 promoter in nonmuscle CV1 cells significantly only in the presence of agonists and exogenous PGC-1. We also demonstrated that PGC-1, relative to SRC-2/GRIP-1 and p300 most efficiently coactivated the M-CPT1 promoter (Fig. 8D
). In summary, these results clearly demonstrate that M-CPT1 is a target for PPARß/
, and selectively coactivated by PGC-1. Cofactor expression increases GW501516 inducibility and the absolute levels of CPT1 expression.
To rigorously define that M-CPT1 was regulated by GW501516 in a PPARß/-PPRE-dependent manner, we mutated the previously defined PPAR
response element in the M-CPT1 promoter between 775 and 763 bp upstream of the initiator codon (27, 34). We showed that the wild-type M-CPT1 promoter and not the PPRE mutant M-CPT1m1 was specifically activated by the PPARß/
-specific agonist (Fig. 9A
). This demonstrated that GW501516 mediated activation is dependent on the M-CPT1 PPRE. This element was previously defined as a PPAR
-regulated motif in cardiac muscle and primary cardiomyocytes (27, 34).
|
In summary, M-CPT1 is an established target for PPAR in cardiac muscle (27, 34, 35). However, we clearly demonstrate by transfection in the presence of PPAR
, -ß/
, and -
agonists that the M-CPT1 promoter in skeletal muscle cells responds preferentially to PPARß/
agonist activation (and not a selective PPAR
agonist) (Fig. 9
, B and C). This is reminiscent of the differential cell specific regulation of the PPRE in the LPL promoter by PPAR
and -
agonists in adipose and liver (32).
Moreover, we showed that the wild type M-CPT1 promoter and not the PPRE mutant M-CPT1m1 was specifically activated by the PPARß/-specific agonist (Fig. 9
, AC). This demonstrated that GW501516-mediated activation was dependent on the M-CPT1 PPRE. This element was previously defined as a PPAR
-regulated motif in cardiac muscle (27, 34, 35).
Moreover, we demonstrate that the native LPL promoter responds to PPAR, not PPARß/
, agonists in muscle cells in the absence of exogenous PPAR
(Fig. 10A
), further validating the specificity of the PPARß/
response on the CPT1 promoter in skeletal muscle cells. The previous literature demonstrates that these cells express functional PPAR
, ß/
, and -
receptors (16, 21, 22, 23). In addition, our data demonstrate the multimerized DR-1 PPRE reporter is efficiently activated by PPAR
, -ß/
, and -
agonists in the absence of exogenous receptors. The LPL promoter data, transfection data in the absence of exogenous receptors and the previous reports above clearly demonstrate the selective and specific activation of M-CPT1 by PPARß/
and not -
agonists, is not due to lack of PPAR
expression, and/or nonfunctional PPAR
. Finally, to rigorously demonstrate that the selective activation of the M-CPT1 promoter is not due to the elevated expression of PPARß/
, relative to PPAR
mRNA, after agonist treatment we examined the expression of PPAR
and -ß/
mRNA expression after 24 h RXR agonist, PPARß/
agonist, and cotreatment (Fig. 10B
). As observed earlier, PPAR
and -ß/
mRNAs are similarly expressed relative to GAPDH mRNA before agonist treatment; however, RXR and PPARß/
agonist treatment preferentially induced PPAR
mRNA expression (Fig. 10B
). This definitively demonstrates that the selective activation of the M-CPT1 promoter by the PPARß/
agonist (and not the PPAR
agonist) in skeletal muscle cells is not due to lack of PPAR
expression.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Our investigation demonstrates that the PPARß/ agonist activates gene expression in skeletal muscle cells, which is involved in preferential lipid utilization, FA catabolism and energy uncoupling. Muoio et al. (12) observed that skeletal muscle metabolism/ß-oxidation and regulation of three well-characterized PPAR
target genes UCP-3, CPT1, and PDK4 (in other tissues) were almost identical in skeletal muscle from either wild type or PPAR
-/- mice (40, 41, 42, 43, 44, 45). Furthermore, metabolism and gene expression in skeletal muscle were not compromised in PPAR
-/- mice, in contrast to the dramatic deleterious effects in liver and heart tissue. Our data account for these observations and suggests PPARß/
in peripheral tissues functions in a complimentary manner to PPAR
in the liver and the heart.
In the context of the data from Evans and colleagues (16) in adipose tissue, we provide further data that suggest that PPARß/ targets skeletal muscle cells. Furthermore, our demonstration that M-CPT1 is preferentially regulated by PPARß/
, and not PPAR
in skeletal muscle cells illustrates distinct mechanisms exist within different cell types to implement localized responses to energy requirements and burdens in these tissues.
The Oliver et al. 2001 study (11) also demonstrated that GW501516 raised cholesterol efflux in macrophages and serum HDL cholesterol. We demonstrate that GW501516 activated ABCA1 mRNA expression with the subsequent metabolic consequence of increased cholesterol efflux from skeletal muscle cells. Therefore, the effects of PPARß/ agonists on skeletal muscle cell gene expression is entirely consistent with the beneficial impact of GW501516 on dyslipidemia and hyperinsulinemia in obese primates, especially when one considers that muscle is a metabolically demanding tissue that accounts for 40% of the total body mass.
Interestingly, in contrast to the role of PPAR in the liver and the heart, fenofibrate induces genes involved in fructose uptake, and glycogen formation in skeletal muscle. Furthermore, we observed a repression in SREBP-1c expression, similar to the effect observed in hepatic cells (46). However, gene expression involved in preferential FA catabolism was not activated by the PPAR
agonist. In congruence with the observations that ß-oxidation in skeletal muscle was not compromised in PPAR
-/- mice. Furthermore, the induction of fructose uptake by fenofibrates is in accordance with the amelioration of high fructose induced insulin resistance, fat accumulation and hyperlipidemia in rats by fenofibrate treatment (47). This suggests that PPARß/
, not PPAR
, regulates lipid catabolism in skeletal muscle cells. We speculate the ß/
isoform also activates FA oxidation in skeletal muscle, in vivo.
Rosiglitazone-mediated activation of PPAR induces gene expression associated with glucose uptake, FA synthesis, and lipid storage, consistent with previous studies. We did not observe robust changes in gene expression after agonist treatment. Thiazolidinediones induce dramatic changes in diseased, not normal healthy animals (48).
We demonstrate that PPARß/ agonists have a significant role in the regulation of the mRNAs encoding the UCPs (UCP-1 to -3, mitochondrial proton carriers) that control metabolic efficiency, energy expenditure, adaptation to nutrient (i.e. preferential lipid utilization) and thermogenesis by uncoupling oxidation/respiration from ATP synthesis (see Fig. 11
). These data are consistent with the observations that mRNA expression associated with selective utilization of lipid substrates is augmented during exercise, starvation, and physiological states that are associated with increased systemic delivery and utilization of FAs. In adult organisms, UCP-1 is almost exclusively expressed in brown adipose tissue, whereas UCP-2 and UCP-3 are expressed in adipose and skeletal muscle tissue, although UCP-3 is predominantly found in muscle. However, UCP-1 mRNA expression has been observed previously in muscle cells (24) and may be an artifact of the myogenesis (muscle differentiation) program in cell culture, and/or reflect an expression profile associated with differentiation during embryogenesis.
A number of studies have investigated ectopic and muscle specific overexpression of UCP-2 and -3 in transgenic mice, and in cell culture. These investigations have reported: 1) reduced metabolic efficiency and increased rates of energy expenditure; 2) preferential FA oxidation vs. glucose utilization/oxidation; 3) resistance to high fat diet induced weight gain, and obesity in the context of hyperphagic behavior; 4) lower fasting plasma glucose and insulin levels, and increased glucose tolerance and clearance rate; and 5) adaptive thermogenesis. These studies emphasize the regulatory role of UCP-2 and -3 in metabolic efficiency/energy expenditure, thermogenesis, and in preferential substrate utilization (49, 50, 51, 52, 53, 54, 55). Similarly, when UCP-1 (normally expressed in brown adipose tissue) is overexpressed in the muscle, the transgenic mice have a lower body weight, increased food intake accompanied by energy expenditure (56). We hypothesize that the effects of PPARß/ agonists on skeletal muscle, a major mass peripheral tissue would have utility and protect against diet-induced obesity and glucose intolerance.
In addition, we confirm that M-CPT1 (and the PPRE), an established target for PPAR in cardiac muscle (27, 34, 35) responds preferentially to PPARß/
agonist activation (and not selective PPAR
agonist) in skeletal muscle cells. This is reminiscent of the differential cell specific regulation of the LPL promoter by PPAR
and -
agonists in adipose and liver. This observation highlights that PPARs have distinct tissue specific functions, and that a single DNA motif can mediate a cell-specific transcriptional phenotype. Furthermore, it suggests that PPAR
and PPARß/
function in a complementary but tissue-specific manner.
Furthermore, we demonstrate the PGC-1-dependent nature of muscle-CPT1 transcriptional activation, and PPRE trans-activation by GW501516-PPARß/ in skeletal muscle cells. The studies by Spiegelman and colleagues (31, 57) exquisitely demonstrate that PGC-1 regulates adaptive thermogenesis, and mitochondrial biogenesis. Moreover, PGC-1 is regulated by exercise (57, 58). Our studies in cells are in consonance with these in vivo studies.
During the preparation of this manuscript, a number of manuscripts appeared in the literature describing the role of PPARß/ agonists in cardiac myocytes and macrophages. For example, Gilde et al. (59) published work describing the role of PPARß/
in cardiac lipid metabolism. They demonstrated that the long chain FA induced regulation of gene expression in primary cardiomyocytes is controlled by PPAR
and PPARß/
. Lipid catabolism was activated in response to PPAR
and -ß/
agonists, concluding that PPAR
and PPARß/
have overlapping functions in the control of cardiac lipid homeostasis. Our investigation, and the study by Wang et al. (16) suggest that PPAR
and -ß/
have distinct functions in adipose and muscle. Moreover, work from Vosper et al. (1) demonstrated that PPARß/
agonists induce lipid absorption and storage in macrophages. Paradoxically, ApoE and cyp27 mRNA expression is repressed, and in contrast, ABCA1 mRNA expression and ApoA1-dependent cholesterol efflux is induced. This is entirely consistent with the observations reported in this study. Furthermore, increased lipid absorption and ApoA1-dependent cholesterol efflux in macrophages and skeletal muscle cells are entirely consistent with the profound effects observed by Oliver et al. (11) in lowering circulating levels of triglycerides and LDL, with a corresponding increase in HDL cholesterol. We comment that this compound is in clinical trials, and the Oliver et al. 2001 manuscript reported that the beneficial effects of GW501516 on HDL cholesterol and triglycerides were observed at 1 and 3 mg/kg with corresponding circulating serum concentrations of approximately 265 and 700 ng/ml, or 0.5 and 1.5 µM, respectively. This is consistent with the concentration used in this study. Finally, Chawla et al. (17) state that triglyceride-enriched VLDLs activate PPARß/
in macrophages and lead to the induction of ADRP/adipophilin (lipid storage droplets), which is consistent with the induction of ADRP mRNA expression we observed in skeletal muscle cells after agonist treatment. In conclusion, we suggest the activation of PPARß/
in skeletal muscle cells programs a cascade of gene expression designed to activate catabolism, and energy expenditure.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
RNA Extraction and Northern Hybridization
Total RNA used for RT-PCR cloning and Northern blot analysis was extracted using TRI-Reagent (Sigma Aldrich Australia Pty Ltd, Castle Hill, New South Wales, Australia), according to manufacturers protocol. For quantitative real-time RT-PCR, RNA was further purified using RNeasy (QIAGEN, Clifton Hill, Victoria, Australia), after manufacturers instructions. Northern blot hybridization was carried out as described previously (61).
Cell Culture, Transient Transfections, and Cholesterol Efflux Assay
Mouse myogenic C2C12 cells were cultured in growth medium [DMEM supplemented with 10% Serum Supreme (BioWhittaker, Edward Keller Pty Ltd, Hallam, Victoria, Australia)] in 6% CO2. For differentiation assays, cells were grown to confluency, at which point media was changed into differentiation medium (DMEM supplemented with 2% horse serum). Cells were harvested at indicated time points. For drug assays, cells were differentiated into myotubes for 4 d (MT4s), and the medium was changed into phenol red-free differentiation medium supplemented with the agonists for PPARß/ (GW501516, 1 µM), RXR (LG101305, 100 nM), PPAR
(Fenofibrate, 100 µM or Wyeth14643, 10 µM), PPAR
(Rosiglitazone, 10 µM), or the vehicle (DMSO) as control. Cells were harvested at the indicated time points (usually 24 h, if not indicated differently). African green monkey kidney CV1 cells were grown in DMEM supplemented with 10% heat-inactivated fetal calf serum.
Transfections were carried out using a DOTAP (N-[1-(2,3-dioleoyloxy)propyl]-N,N,N-trimethylammonium methylsulfate) (Biontex Laboratories GmbH, Munich, Germany)/DOSPER (1,3-di-oleoyloxy-2-(6-carboxy-spermyl)-propylamid) (Roche Diagnostics Pty Ltd, Castle Hill, New South Wales, Australia) 3:1 liposome mixture in HEPES buffered saline [42 mM HEPES, 275 mM NaCl, 10 mM KCl, 0.4 mM Na2HPO4, 11 mM dextrose (pH 7.1)], with 1µg of total DNA per well. Medium was replaced after 16 h with the respective fresh medium, supplemented with agonists for PPARs and/RXR (as described above), harvested after 24 h and assayed for LUC activity using the Luclite kit (PerkinElmer Life Science, Knoxfield, Victoria, Australia) according to manufacturers protocol.
ApoA1-dependent cholesterol efflux was performed as described previously (19).
Quantitative RT-PCR
Target cDNA levels were quantitated by real-time RT-PCR using an ABI Prism 7700 Sequence Detector system utilizing SYBRE green I (Molecular Probes, Eugene, OR; catalog no. S-7562, used at 0.8x) as a nonspecific PCR product fluorescence label. Quantitation was over 45 cycles of 95 C for 15 sec and 60 C for 1 min two-step thermal cycling preceded by an initial 95 C for 2 min for activation of 0.75 U Platinum Taq DNA polymerase (Invitrogen Australia Pty Ltd, Mulgrave, Victoria, Australia). The 25-µl reaction also contained 20 mM Tris-HCl (pH 8.4), 50 mM KCl, 5 mM MgCl2, 200 µM each of deoxy (d) GTP, dATP, dCTP, 400 µM deoxyuridine triphosphate, 0.5 U uracil-N-glycosylase, 500 nM ROX reference dye (Invitrogen) and 200 nM each forward and reverse primers. Mus musculus primer sequences (forward and reverse, respectively): ABCA1: GCTCTCAGGTGGGATGCAG, GGCTCGTCCAGAATGACAAC;
ABCG1: CTGAGGGATCTGGGTCTGA, CCTGATGCCACTTCCATGA;
ACS4: GGTTTGGTAACAGATGCCTTCAA, CCCATACATTCGCTCAATGTCTT; ADRP: CCCTGGTTCTAAGAAGCTGCTTT, GGCCAGATGACCCCTTTTG;
ApoE: GCTGTTGGTCACATTGCTGA, TGCCACTCGAGCTGATCTG;
CD36: GGCCAAGCTATTGCGACAT, CAGATCCGAACACAGCGTAGA;
CPT1: ATCATGTATCGCCGCAAACT, CCATCTGGTAGGAGCACATGG;
FABP3: CCCCTCAGCTCAGCACCAT, CAGAAAAATCCCAACCCAAGAAT;
FAS: CGGAAACTTCAGGAAATGTCC, TCAGAGACGTGTCACTCCTGG;
GAPDH: GTGTCCGTCGTGGATCTGA, CCTGCTTCACCACCTTCTTG;
Glut4: ATGGCTGTCGCTGGTTTCTC, ACCCATACGATCCGCAACAT;
Glut5: CTTGCCTTTACCGGGTTGAC, CATCTGGTCTTGCAGCAACTCT;
GYG1: CCCAAACCCCTCATCTGATG, GCACGTTTCCATACATAGTATGTGAA;
LPL: CCAATGGAGGCACTTTCCA, TGGTCCACGTCTCCGAGTC;
PDK2: TGCTCCGGCTTGCCTTAT, CACTCCATCCTTCTTAACATTGACA;
PDK4: AAAGGACAGGATGGAAGGAATCA, TTTTCCTCTGGGTTTGCACAT;
PPAR: TCTTCACGATGCTGTCCTCCT, GGAACTCGCCTGTGATAAAGC
PPARß/: TCCAGAAGAAGAACCGCAACA, GGATAGCGTTGTGCGACATG;
PPAR: CAGGCCGAGAAGGAGAAGCT, GGCTCGCAGATCAGCAGACT
SCD1: TGTACGGGATCATACTGGTTCC, CCCGGCTGTGATGCC;
SCD2: ACTGTGACTCAAGTTCAACTCTTGAAA, TGCCCACAAATTGAGGATAGC;
SREBP-1c: CGTCTGCACGCCCTAGG, CTGGAGCATGTCTTCAAATGTG;
UCP-1: ACAGAAGGATTGCCGAAAC, AGCTGATTTGCCTCTGAATG;
UCP-2: GTTCCTCTGTCTCGTCTTGC, GGCCTTGAAACCAACCA;
UCP-3: TGACCTGCGCCCAGC, CCCAGGCGTATCATGGCT.
Amplification specificity was verified by visualizing PCR products on an ethidium bromide-stained 2.5% agarose gel. GAPDH was used for normalization between samples for quantitation.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
FOOTNOTES |
---|
This work was supported by the National Health and Medical Research Council (NHMRC) of Australia. G.E.O.M. is an NHMRC Principal Research Fellow, and U.D. is a University of Queensland Postoctoral Research Fellow.
Abbreviations: ABC, ATP binding cassette; ACS, acyl-CoA synthetase; ADRP, adipocyte differentiation-related protein; ApoE, apolipoprotein; CD36/FAT, FA translocase; CMB, confluent myoblasts; CoA, coenzyme A; CPT, carnitine palmitoyl transferase; DMSO, dimethylsulfoxide; FA, fatty acid; FABP, FA binding protein; FAS, fatty acid synthase; FFA, free fatty acids; G-6-P, glucose-6-phosphate; GLUT, glucose transporters; GYG1, glycogenin; HDL, high-density lipoprotein; LDL, low-density lipoprotein; LPL, lipoprotein lipase; LUC, luciferase; LXR, liver X receptor; M-CPT1, muscle carnitine-palmitoyl-transferase-1 (M-CPT1); MUFAs, monounsaturated fatty acids; NR, nuclear hormone receptor; PDC, pyruvate dehydroxygenase complex; PDK, pyruvate dehydroxygenase kinases; PGC-1, PPAR coactivator-1; PMB, proliferating C2C12 myoblasts; PPAR, peroxisome proliferator-activated receptor; PPRE, PPAR-dependent reporter; RXR, retinoid X receptor; SCD, stearoyl CoA desaturase; SFAs, saturated fatty acids; TCA, tricarboxylic acid; tk, thymidine kinase; UCPs, uncoupling proteins; VLDL, very low-density lipoprotein.
Received for publication April 22, 2003. Accepted for publication September 29, 2003.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|