PPAR� regulates glucose metabolism andinsulin sensitivityChih-Hao Lee*†, Peter Olson†‡§, Andrea Hevener¶, Isaac Mehl‡§, Ling-Wa Chong‡, Jerrold M. Olefsky¶, Frank J. Gonzalez�,Jungyeob Ham**, Heonjoong Kang**, Jeffrey M. Peters††, and Ronald M. Evans‡,‡‡

*Department of Genetics and Complex Diseases, Harvard School of Public Health, 665 Huntington Avenue, Boston, MA 02115; ‡Howard Hughes MedicalInstitute, Gene Expression Laboratory, The Salk Institute for Biological Studies, 10010 North Torrey Pines Road, La Jolla, CA 92037; Departments of §Biologyand ¶Medicine, University of California at San Diego, 9500 Gilman Drive, La Jolla, CA 92093; �Laboratory of Metabolism, Division of Basic Sciences, NationalCancer Institute, Bethesda, MD 20892; **Marine Biotechnology Laboratory, School of Earth and Environmental Sciences and Center for Marine NaturalProducts Drug Discovery, Seoul National University, Seoul 151-747, Korea; and ††Department of Veterinary Science and Center for Molecular Toxicologyand Carcinogenesis, Pennsylvania State University, University Park, PA 16802

Contributed by Ronald M. Evans, January 4, 2006

The metabolic syndrome is a collection of obesity-related disorders.The peroxisome proliferator-activated receptors (PPARs) regulatetranscription in response to fatty acids and, as such, are potentialtherapeutic targets for these diseases. We show that PPAR�(NR1C2) knockout mice are metabolically less active and glucose-intolerant, whereas receptor activation in db�db mice improvesinsulin sensitivity. Euglycemic–hyperinsulinemic-clamp experi-ments further demonstrate that a PPAR�-specific agonist sup-presses hepatic glucose output, increases glucose disposal, andinhibits free fatty acid release from adipocytes. Unexpectedly,gene array and functional analyses suggest that PPAR� ameliorateshyperglycemia by increasing glucose flux through the pentosephosphate pathway and enhancing fatty acid synthesis. Couplingincreased hepatic carbohydrate catabolism with its ability to pro-mote �-oxidation in muscle allows PPAR� to regulate metabolichomeostasis and enhance insulin action by complementary ef-fects in distinct tissues. The combined hepatic and peripheralactions of PPAR� suggest new therapeutic approaches to treattype II diabetes.

fatty acid and glucose metabolism � insulin sensitivity

Syndrome X, or metabolic syndrome, includes several disor-ders resulting from metabolic dysregulation (1). Among

these, insulin resistance alone affects millions of people world-wide. This disease is characterized by a compromised ability ofinsulin to control glucose and lipid metabolism, which results indecreased glucose disposal to muscle, increased hepatic glucoseproduction, and overt postprandial hyperglycemia. When leftuntreated, insulin resistance leads to the development of type IIdiabetes and related complications, including cardiovasculardisease, which are leading causes of death.

The PPARs are members of the nuclear receptor superfamily,which are ligand-modulated transcription factors that regulategene expression programs of many important biological pro-cesses (2). The three PPARs, � (NR1C1), ��� (NR1C2), and �(NR1C3), are activated by lipids and are targets for current andprospective drug therapies for components of the metabolicsyndrome (3, 4). PPAR�, a target for the fibrate class oflipid-lowering drugs (5–7), is primarily expressed in liver, whereit up-regulates genes involved in lipid oxidation in the fastedstate (8, 9). PPAR� is highly expressed in adipose tissue andregulates adipogenesis and insulin sensitivity (10–14). Thiazo-lidinediones are a class of drugs that increase insulin sensitivitythrough activating PPAR� (15). PPAR� is expressed in manytissues, including metabolically active sites such as liver, muscle,and fat, and its role in the metabolic syndrome is only now beingelucidated (16, 17). Treatment with a high-affinity PPAR�agonist GW501516 has been shown to increase high-densitylipoprotein cholesterol (18, 19), affect lesion progression in amouse model of atherosclerosis (20), cause weight loss, andregulate muscle fiber type-switching when constitutively acti-

vated (21, 22). In the latter two cases, the phenotypes appearedto be mediated through up-regulation of lipid catabolism andoxidative phosphorylation in fat and muscle. In addition, ligandsfor PPAR� have been proposed to be potential insulin sensitiz-ers, based on improvements in standard glucose-tolerance tests(21, 23). These studies, however, used long-term ligand-treatment regimens that resulted in significant weight loss and adecrease in fat mass. These effects alone enhance insulin sen-sitivity. Therefore, it remains unclear whether PPAR� candirectly regulate insulin sensitivity and, if so, through whichtissue and what mechanism.

In this study, we sought to determine the effect of a PPAR�synthetic ligand in animals that have already developed insulinresistance but under conditions in which body weight, foodintake, and serum lipid profiles are minimally affected after drugintervention. We show that PPAR� knockout mice are glucoseintolerant, whereas treatment of diabetic db�db mice with ahigh-affinity PPAR� agonist improves insulin sensitivity in allmajor insulin-responsive tissues. Molecular and functional anal-yses suggest that PPAR� activation reduces hepatic glucoseoutput by increasing glycolysis and the pentose phosphate shuntand promoting fatty acid synthesis in the liver. This uncoveredhepatic activity thus constitutes the earliest component of theregulatory mechanism by which PPAR� regulates insulin sensi-tivity, in addition to its known function in fatty acid �-oxidation.

ResultsPPAR� Mutant Mice Are Metabolically Less Active and Glucose Intol-erant. To explore PPAR�’s role in the onset of the metabolicsyndrome, we examined a cohort of 6-month-old male PPAR�null mice along with age-matched wild-type controls in meta-bolic cages. Although PPAR��/� mice have been shown to gainless weight and exhibit defects in hepatic TG production at ayounger age (17, 24), the weight difference normalized in oldercohorts, and no significant changes were detected in levels ofcirculating Acrp30, insulin, glucose, and lipids between thecontrol and mutant group (see Table 3, which is published assupporting information on the PNAS web site). In addition,these weight-normalized null mice consumed less food, water,and oxygen and produced less carbon dioxide than controlanimals when fed a standard chow diet (Fig. 1 A and B).Metabolic rate (presented as heat) was also decreased, indicatingthat these animals expended less energy (Fig. 1C). The respira-tory exchange ratio [(RER) � VCO2�VO2], which indicates

Conflict of interest statement: No conflicts declared.

Abbreviations: GW, GW501516; Q-PCR, quantitative PCR; RER, respiratory exchange ratio;TG, triglyceride.

†C.-H.L. and P.O. contributed equally to this work.

‡‡To whom correspondence should be addressed. E-mail: evans@salk.edu.

© 2006 by The National Academy of Sciences of the USA

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whether lipids (RER � 0.7) or carbohydrates (RER � 1) arebeing oxidized, was then determined. Wild-type animals wentfrom burning �70% of their fuel from lipids during the fastedstate to nearly 100% from carbohydrates in the fed state (Fig.1D; and see Fig. 5A, which is published as supporting informa-tion of the PNAS web site, for statistical analysis). Null mice, onthe other hand, had significantly lower RER values, particularlyduring the fed state, suggesting a defect in switching their energysource. This observation prompted us to investigate glucosetolerance in the null animals, because one of the early signs ofdiabetes is a blunted response to insulin-stimulated glucoseutilization in the fed state. After a 6-hour fast, both genotypeshad similar baseline glucose concentrations (Fig. 1E). Whensubjected to a glucose-tolerance test, which evaluates the abilityof the body to adjust glucose levels after an acute glucoseinjection, glucose levels of wild-type mice peaked at 20 min andrapidly returned to the basal level. In contrast, null animalsshowed a delayed response, indicating glucose intolerance. Bothwild-type and null animals (n � 8) were then fed a high-fat,high-carbohydrate (HF) diet for 10 weeks to induce insulinresistance, followed by treatment with either GW501516 (GW),a high-affinity PPAR� agonist, at 4 mg�kg�day or DMSO alone(four mice from each genotype) for 2 weeks. Both groups gainedsimilar weight on this diet. Ligand treatment for 2 weeks also didnot affect weight gain in either genotype (see Fig. 6A, which is

published as supporting information on the PNAS web site).Because long-term treatment with this compound leads toweight loss, which affects the dynamics of metabolism (21, 23),this experimental design allowed us to directly study the acuteeffect of PPAR� activation on glucose tolerance.

In agreement with previous reports, RER values remained low(25) (�0.7) throughout the 24 h (Fig. 6B) because of the high fatcontent (35% by weight) in the diet. To test the efficacy of GWto improve glucose tolerance and whether the effect is receptor-dependent, all four groups were then subjected to a glucose-tolerance test (GTT). Both wild-type and null mice displayedhyperglycemia (220.6 � 14.0 in wild-type vs. 215.9 � 14.2 in nullmice, P � 0.8). Interestingly, both groups were similarly glucose-intolerant after high-fat, high-carbohydrate diet treatment (Fig.1F). Wild-type mice treated with GW, however, had a lowerfasting glucose level (225.0 � 11.4 in DMSO vs. 182.5 � 2.9 inGW, P � 0.03), and their response to the glucose challenge wasgreatly improved. Importantly, this improvement in the GTT byGW treatment was completely abolished in the null mice,demonstrating that this effect is receptor-dependent. These datasupport the proposal that PPAR� functions as a mediator ofinsulin sensitivity.

PPAR� Activation Increases Hepatic and Peripheral Insulin Sensitivity.To determine whether PPAR� acts as an insulin sensitizer, wenext examined the activity of GW on db�db mice, a wellestablished model to study diabetes. These animals becomeinsulin resistant on normal chow, which also allowed us toexamine RER. Ten 10-week-old male db�db mice were treatedwith either 4 mg�kg�day GW or DMSO for two weeks. Thisregimen did not cause changes in weight gain, food intake, orlevels of circulating free fatty acids, whereas total and high-density lipoprotein cholesterol was increased with GW treatment(Table 1). In line with the decreased TG level observed afterlong-term treatment (21, 23), serum TG levels were moderatelylower in the GW-treated group (Table 1 and data not shown).However, the difference was not statistically significant. Theliver-to-body-weight ratio was increased in the GW-treatedgroup, described in ref. 23. GW treatment significantly increasedRER, particularly during the fed state, indicating increasedcarbohydrate usage. Consistent with results from the HF study,GW treatment dramatically lowered fasting glucose and en-hanced glucose tolerance (Fig. 2B and Table 1). Similarly, GWalso increased the ability of insulin to lower glucose in theinsulin-tolerance test (Fig. 2C), demonstrating improved insulinsensitivity. Although there was a decrease in fasting serum

Fig. 1. Depressed metabolism and glucose intolerance in PPAR� null mice.Metabolic parameters of wild-type and PPAR� null mice were measured inmetabolic cages (n � 8). PPAR� null mice ate and drank less (A), consumed andproduced less O2 and CO2 (B), expended less energy (presented as heat) (C),and have lower RERs (RER � VCO2�VO2) (D), particularly in the fed state (6 p.m.to 6 a.m.). (E) The glucose-tolerance test (GTT), showing that PPAR� nullanimals were glucose-intolerant on normal chow. (F) The GTT, showing areceptor-dependent effect of a PPAR� agonist (GW) on improving glucosetolerance. Both wild-type and PPAR��/� mice were placed on a high-fathigh-carbohydrate diet for 10 weeks, followed by a 2-week ligand treatment(n � 4). Statistical differences were observed only between vehicle- and ligand(GW)-treated wild-type mice. *, P � 0.05.

Table 1. Metabolic parameters of db�db mice

Parameter Vehicle GW501516

Weight gain, g 5.91 � 0.56 6.06 � 0.46Liver�body weight 0.058 � 0.001 0.072 � 0.003**Fat�body weight 0.039 � 0.001 0.039 � 0.002Serum lipids

TG, mg�dl 300.42 � 37.58 272.61 � 34.17Free fatty acid, mEq�liter 0.48 � 0.05 0.46 � 0.04Cholesterol, mg�dl 123.58 � 6.69 164.23 � 9.69**HDL-c, mg�dl 68.20 � 3.97 94.21 � 6.05**

Fasting glucose, mg�dl 594.4 � 45.6 396.0 � 41.4*298.0 � 27.1† 229.3 � 17.0†*

Fasting insulin, ng�ml 27.24 � 4.48 20.91 � 2.9423.08 � 6.65† 7.38 � 1.82†*

Acrp30, mg�ml 7.20 � 0.73 9.00 � 1.08

Mice were 10 weeks old at the start of the 2-week drug treatment. HDL-c,high-density lipoprotein cholesterol.†8-week-old mice at the start of the 2-week drug treatment. *, P � 0.05;

**, P � 0.005.

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insulin (27.2 vs. 20.9 ng�ml) in the GW-treated group, it was notstatistically significant. However, ligand treatment decreasedinsulin levels in a younger cohort in which insulin values were lessvariable within the groups (see below).

To further define which tissues have improved insulin sensi-tivity and to what extent, we performed euglycemic–hyperinsulinemic clamp. The animals used for this study were 8weeks old. After a 2-week treatment, GW had no effect on bodyweight (35.9 � 0.13 g vs. 34.6 � 0.2 g; P � 0.613) but significantlylowered fasting blood glucose and insulin levels (23.0 vs. 7.4ng�ml) (Table 1). As predicted from these reductions, basalglucose turnover was decreased by 35% (P � 0.028, Fig. 2E,vehicle vs. GW at the basal state), and the exogenous glucoseinfusion rate (GIR) required to maintain euglycemia was mark-edly elevated by 57% (P � 0.04) in the GW-treated mice (Fig.2D). A good portion of the difference in GIR could be accountedfor by the insulin-stimulated reduction in hepatic glucose pro-duction (P � 0.016; Fig. 2E, vehicle vs. GW in the clampedstate), which was improved by 47% and represents improvedhepatic insulin sensitivity. Insulin-stimulated glucose disposalwas increased �2-fold by the drug (Fig. 2F), demonstrating thatthis compound also improved peripheral insulin sensitivity. Inaddition, GW treatment also led to an enhanced insulin responsein adipocytes, as demonstrated by a 33% reduction in serum freefatty acids after the clamp (Fig. 2G).

PPAR� Regulates Critical Transcriptional Programs of Glucose andFatty Acid Metabolism in Liver. To determine the molecular mech-anism underlying the insulin-sensitizing activity of PPAR� ac-tivation, we conducted DNA array analysis to compare geneexpression profiles of liver, adipocytes, and muscle betweencontrol and GW-treated db�db mice. The results obtained were

confirmed by either quantitative-PCR (Q-PCR) or Northernblot (Table 2; and see Fig. 7, which is published as supportinginformation on the PNAS web site). Several key genes were alsosimilarly regulated in samples from high-fat-diet-induced insu-lin-resistant mice, further confirming the importance of thesetarget genes (Fig. 7B). Unexpectedly, there were no significantdifferences in the expression of relevant genes in adipocytes,whereas carnitine palmitoyltransferase 1 (CPT1), a key gene infatty acid �-oxidation, was identified as a target in muscle asdescribed in refs. 21 and 23. The most responsive tissue was theliver, where clusters of genes involved in fatty acid synthesis wereup-regulated by ligand treatment (Table 2). These genes can befurther divided into three groups. The first group is comprisedof genes that are directly responsible for fatty acid synthesis,including acetyl-CoA carboxylase � (ACC�), fatty acid synthase(FAS), acyl-CoA thioesterase 1, and ATP citrate lyase. Thesecond group consists of genes involved in elongation andmodification of fatty acids including ELOVL family member 6(ELOVL6), stearoyl-CoA desaturase 2 (SCD2), and glycerol-3-phosphate acyltransferase (GPAT). The third group includesgenes such as malic enzyme in the pyruvate�malate cycle andphosphogluconate dehydrogenase (PGD) in the pentose phos-phate pathway that mediates NADPH generation to providereducing power for lipid synthesis. It is interesting to note thatboth fatty acid synthesis and the pentose phosphate pathway useglucose or its metabolites as substrates. It has been demonstratedthat overexpression of glucokinase in liver increases glycolysis byactivating the pentose phosphate shunt with concomitant up-regulation of lipogenic genes, including ACC1 and FAS, resultingin reduced hepatic glucose production, increased fatty acidoxidation in muscle, and improved peripheral insulin sensitivity(26). This suggests that PPAR� may increase glucose catabolismthrough these processes.

Other genes that are involved in lipid transport (CD36 andphospholipid transfer protein, PLTP) and cholesterol synthesis(mevalonate decarboxylase) were also identified from thearray. However, it appeared that cholesterol synthesis was notaltered, because the expression of the rate-limiting enzymeHMG-CoA reductase remained unchanged (Fig. 3A), andthere was no difference in liver cholesterol content (Fig. 4C).GW treatment may also inhibit glycogenolysis through sup-pressing protein phosphatase 1 regulatory subunit (Ppp1r3C)expression. We also examined the expression of phosphoenol-pyruvate carboxykinase (PEPCK), a rate-limiting enzyme forgluconeogenesis, which was not included in the array collec-tion, and found a moderate reduction in GW-treated samples(Fig. 7A). Interestingly, we did not detect expression changesin known PPAR� targets, such as acetyl-coA oxidase andCPT1 in liver (Figs. 4 and 7A), indicating that these tworeceptors regulate distinct transcriptional programs.

Recently, it has been shown that liver X receptors (LXR� and-�) regulate lipogenesis through transcriptional induction ofsterol regulatory element-binding protein (SREBP)-1c (27, 28),a master regulator of hepatic lipid metabolism. To rule out thepossibility that the observed regulation was indirectly mediatedby SREBP-1c, we analyzed its expression in liver by Q-PCR andfound no changes between control and GW-treated groups (Fig.3A). We next examined the activity of the ACC� promoter in areporter transient transfection assay. The human ACC� gene iscontrolled by two promoters (29) (Fig. 3B). SREBP-1c is knownto induce its expression through promoter II. However, PPAR�did not affect the activity of a reporter driven by this promoter[PII (1.3 kb)] when cotransfected into HepG2 cells (Fig. 3C). Incontrast, we found that promoter I [PI (0.58 kb)] could beactivated by PPAR�. Subsequently, a putative PPAR responseelement was identified at �480 to �468 (relative to the tran-scriptional start site). Mutation [PI (0.58 kb�M)] or deletion ofthis element completely abolished the ability of PPAR� to

Fig. 2. Treatment with a PPAR� agonist increases insulin sensitivity in db�dbmice. PPAR� ligand treatment increased RER (A) and improved performance inthe glucose-tolerance test (B) and insulin-tolerance test (C) in db�db mice (n �10). In the insulin-tolerance test, glucose levels after insulin injection werepresented as the percentage of initial glucose concentrations. (D–G) Euglyce-mic–hyperinsulinemic clamp in db�db mice, showing improved insulin sensi-tivity in multiple tissues after ligand treatment. PPAR� ligand-treated micehave a higher glucose infusion rate (GIR) (D), decreased hepatic glucoseproduction (HGP) (F), increased insulin stimulated-glucose disposal rate (IS-GDR) (G), and lower free fatty acid levels after clamp (F). Vehicle (�);GW501516 (■ ); *, P � 0.05

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induce the promoter activity (Fig. 3C and data not shown). Theseresults suggest that ACC� is a target gene of PPAR� and indicatethat PPAR� controls hepatic fatty acid synthesis through directtranscriptional control.

PPAR� Increases Glucose Consumption, Activates the Pentose Phos-phate Shunt, and Promotes Fatty Acid Synthesis. Previous studiesand our array data have demonstrated a critical role of PPAR�in peripheral fatty acid oxidation (21, 23). However, this bene-ficial activity alone does not explain how PPAR� improvesinsulin sensitivity. Therefore, the regulation of the glycolyticpathways mediated by the pentose phosphate shunt and lipo-genesis appears to constitute the second component of theinsulin-sensitizing action of PPAR�. To test this hypothesis, weperformed a series of in vitro functional assays. First, we mea-sured the activity of hepatic glucose-6-phosphate dehydrogenase(G-6-PDH), the rate-limiting enzyme in the pentose phosphatepathway, and found a 2-fold (100%) increase in livers of GW-treated db�db mice compared with the control group (Fig. 4A).The flux from glucose to fatty acid synthesis was then directlydetermined in primary hepatocytes isolated from both vehicle-and GW-treated db�db mice by [14C]glucose labeling. Indeed,there was also an �2-fold increase in the conversion rate of14C-labeled glucose into organically extractable lipids (TGs�fatty acids) in GW-treated cells (Fig. 4B). Consistent with thefunctional analyses, we detected a 20% increase in liver TGcontent in GW-treated animals, whereas cholesterol and glyco-gen content was not affected (Fig. 4C and data not shown).Lastly, we examined fatty acid catabolism in isolated soleusmuscle strips and found GW treatment significantly increasedthe �-oxidation rate (Fig. 4D), which is consistent with resultsfrom previous reports (21, 23) and the up-regulation of CPT-1in muscle. In summary, these data reveal a previously unrecog-nized role of PPAR� in the liver and suggest that PPAR�regulates metabolic homeostasis and insulin sensitivity througha unique mechanism, which consists of a fat-production com-

ponent in liver to consume glucose and a counterbalancing fatburning component in muscle to reduce lipid burden.

DiscussionThis study utilizes metabolic, genetic, and pharmacologic ap-proaches to examine the role of PPAR� in regulating insulinsensitivity. We identify liver as a major PPAR�-responsive tissueand reveal an unexpected glucose-burning pathway, which con-tributes to the ability of PPAR� agonists to alleviate hypergly-cemia and improve insulin sensitivity.

Based on the glucose–fatty acid cycle proposed by Randle(30), an induction in fatty acid oxidation should result indecreased carbohydrate usage. This hypothesis raises an intrigu-ing question as to how activation of PPAR�, a fatty acid receptorknown to activate lipid oxidation, enhances glucose utilization.The answer seems to reside in the ability of PPAR� to promoteglucose flux through the pentose phosphate shunt and stimulatefatty acid synthesis in liver. The former generates reducing power(NADPH) needed for lipid synthesis and can consume up to 20%of hepatic glucose (31, 32). The importance of the pentosephosphate pathway and lipogenesis in regulating hepatic glucoseoutput and systemic metabolic homeostasis has emerged inrecent studies. It has been demonstrated that deletion of theGLUT4 gene leads to activation of hepatic fatty acid synthesis�pentose phosphate pathway and increased skeletal-muscle �-oxidation, an energy substrate-switching phenotype, because ofthe inability of the body to use glucose. Thus, fats were made atthe expense of glucose in the liver to be consumed in muscle,providing a mechanism how GLUT4 knockout mice maintainnormal glucose levels and insulin sensitivity (33). Importantly,adenoviral-mediated glucokinase overexpression drives glycol-ysis through the fatty acid synthesis�pentose phosphate pathway,resulting in reduced hepatic glucose production, increased �-oxidation in muscle, and improved systemic insulin sensitivity(26). It appears that PPAR� activation exerts an ‘‘energy sub-strate-switching’’ phenotype, suggesting that this receptor is

Table 2. PPAR� target genes identified by DNA array analysis

Gene Array Q-PCR Northern blot

LiverFatty acid�TG synthesis

Fatty acid synthase (Mm.268690*) 2.1** 3.0 3.0Acetyl-CoA carboxylase � (Mm.81793) 2.2 3.5Stearoyl-CoA desaturase 2 (Mm.193096) 3.7 3.1ELOVL family member 6 (Mm.26171) 2.9 4.5Acyl-CoA thioesterase 1 (Mm.1978) 1.7Glycerol-3-phos acyltransferase (Mm.275758) 1.7 2.5

Pyruvate�malate cycleMalic enzyme (Mm.148155) 2.2 2.6ATP citrate lyase (Mm.25316) 2.4 2.3Carnitine acetyltransferase (Mm.20396) 1.5

Pentose phosphate pathwayPhosphogluconate dehydrogenase (Mm.252080) 1.8 2.8

GlycogenolysisProtein phos 1, reg (inhib) subunit C (Mm.24724) �2.3 �1.8

Cholesterol synthesisMevalonate decarboxylase (Mm.28146) 2.3 3.3

Lipid transport�metabolismCD36 (Mm.18628) 1.4 1.9Phospholipid transfer protein (Mm.6105) 3.2 4.4

MuscleLipid oxidation

Carnitine palmitoyltransferase 1 (Mm.18522) 1.6 1.8

*UniGene number.**Fold-changes comparing ligand-treated to the control group. Minus sign indicates down-regulation.

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capable of controlling the substrate utilization through regulat-ing distinct transcriptional programs in different tissues, whichrepresents a unique mechanism for metabolic regulation. Theincrease in TG raises the concern as to whether long-term drugtreatment will cause hepatic steatosis. However, we did notobserve signs of fatty liver with treatment up to 6 months inwild-type C57BL�6 mice (data not shown). It is likely that theincreased �-oxidation in muscle results in a net decrease in thefat content. Indeed, long-term treatment has been shown toreduce body weight and levels of circulating and liver TG(21, 23).

Together with previous findings, this study expands our viewof the differential roles of the three PPARs in lipid metabolism,in which PPAR� is essential for hepatic fatty acid oxidationduring fasting, PPAR� is important for lipid synthesis�storage inadipocytes, and PPAR� increase glycolysis�lipogenesis in liverwhile activating fat burning in muscle. Consistent with this idea,pharmacological activation of PPAR� and PPAR� improvesinsulin sensitivity through regulating different metabolic path-ways. Unlike thiazolidinedione treatment, which increases bodyweight, reduces serum fatty acid, and increases levels of Acrp30(3), PPAR� agonists drastically improve insulin sensitivity with-out affecting these parameters (Table 1). In addition, Q-PCRanalyses demonstrated that the expression of PPAR� targetgenes in adipocytes, such as TNF�, GLUT4, and CD36 wasunaffected by GW treatment (see Fig. 8, which is published as

supporting information on the PNAS web site). Our datasuggests that activation of PPAR� promotes a metabolic shift toreduce hepatic glucose output, enabling an enhancement ofinsulin sensitivity. It is likely, however, that PPAR� would exerta more profound effect on muscle and adipocyte with prolongedligand treatment as proposed in refs. 21 and 22. Together withthe PPAR� loss-of-function data, our study implicates PPAR� asa bona fide target to treat insulin resistance and as a candidatefor population-based screening for genetic polymorphisms thatpredispose to this disease.

Materials and MethodsAnimal Experiments. PPAR� knockout mice were generated intwo laboratories in different genetic backgrounds (16, 17).PPAR� null animals in the C57BL�6 background were used formetabolic studies (eight age-matched 6-month-old male PPAR�null and wild-type mice) (17). These mice were then challengedwith a high-fat, high-carbohydrate diet (F3282, Bio-Serv,Frenchtown, NJ) for 10 weeks to induce insulin resistance. Fourmice from each group were given either vehicle or ligand for anadditional 2 weeks. GW501516 compound was dissolved inDMSO and suspended in 0.5% carboxymethylcellulose. Micewere gavaged at the dose of 4 mg�kg�day based on a previousreport (18). Db�db mice were purchased from The JacksonLaboratory. Ten db�db male mice (10 weeks old) were used forboth control and ligand-treated groups for metabolic studies,whereas six mice (8 weeks old) for each group were included inthe clamp study. These mice were also given the ligand at 4mg�kg�day for 2 weeks.

Fig. 3. PPAR� regulates hepatic glucose and fatty acid homeostasis throughdirect transcriptional regulation. (A) Q-PCR analyses demonstrating increasedACC� expression in livers of GW-treated db�db mice (Left). GW had no effectson levels of SREBP-1c (Center) or HMG-CoA reductase (Right). (B) The 5�regulatory regions of the human ACC� gene. ACC� is controlled by twopromoters. A putative PPAR� response element (PPRE) was identified in thepromoter I (�480 to �468, relative to the transcriptional starting site). Resi-dues that were mutated in the reporter construct [PI (0.58 kb�M)] are indi-cated. (C) ACC� is a direct target gene of PPAR�. HepG2 cells were cotrans-fected with a luciferase reporter driven by the promoter I or II of the ACC�

gene, together with control vectors or expression vectors for PPAR� and theheterodimer partner RXR�, as well as a �-galactosidase internal control. Cellswere treated with GW at 0.1 �M for 16 h. PPAR� increases the activity of thepromoter I in a PPRE-dependent manner. PI (0.58 kb), the luciferase reporterdriven by 0.58-kb 5� upstream region of the promoter; I, PI (0.58 kb�M);0.58-kb promoter I with PPRE mutations; PII (1.3 kb), 1.3-kb promoter II.

Fig. 4. PPAR� promotes glucose flux to the pentose phosphate pathway andfatty acid synthesis in liver. (A) PPAR� ligand activates the pentose phosphatepathway. GW treatment increases the activity of glucose-6-phosphate dehy-drogenase, the first enzyme in the pentose phosphate pathway, by 2-fold. Theactivities were determined by NADPH produced (OD 340 nm) by using liverlysates from vehicle or GW-treated db�db mice (n � 10). (B) PPAR� increaseshepatic glucose-to-fatty acid conversion. Primary hepatocytes were isolatedfrom vehicle or GW-treated db�db mice and loaded with [14C]glucose. Theamount of glucose-derived fatty acid was determined by the radioactivityrecovered from the organic phase after lipid extraction and was normalized toprotein concentration. (C) Increased hepatic TG, but not cholesterol, contentin ligand-treated db�db mice. (D) PPAR� increases �-oxidation in muscle.Soleus muscle strips were isolated from vehicle or GW-treated db�db mice andloaded with [3H]palmitic acid. The catabolic rate was determined by measur-ing the fatty acid �-oxidation product 3H2O and normalized to tissue weight.Vehicle (�); GW501516 (■ ); *, P � 0.05.

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Metabolic Studies. Metabolic cage studies were conducted in aComprehensive Lab Animal Monitoring System (12-chamberCLAMS system, Columbus Instruments, Columbus, Ohio). Se-rum and plasma were collected after 6-h fasting. Total choles-terol, TG, and free fatty acids were determined by using enzy-matic reactions (Thermo DMA, Arlington, TX; free fatty acid,Wako Chemicals, Richmond, VA). The activity of G-6-PDH inliver lysates from vehicle or GW-treated db�db mice was deter-mined by NADPH produced (OD340) within a period of 5 minby using a commercial kit (Trinity Biotech, St. Louis). Aglucose-tolerance test was preformed by injecting 1.5 mg ofglucose per g of body weight into the peritoneum after 6-hfasting. Blood glucose was measured through the tail tip beforeand after injection at the time course indicated by using theOneTouch (Lifescan, Milpitas, CA) glucose-monitoring system.ITT was conducted similarly by injecting 2 units of insulin per kgof body weight. Insulin and Acrp30 were measured by usingradioimmunoassay kits (Linco, St. Charles, MO). Methods for invitro functional assays and hyperinsulinemic–euglycemic clampare described in Supporting Methods, which is published assupporting information on the PNAS web site. Statistics wereperformed by using Student’s t test. Values were presented asmeans � SEMs. Significance was established at P � 0.05.

Gene Expression Analysis. Tissue RNAs were isolated by usingTRIzol reagent (Invitrogen). For DNA array analysis, RNAsamples from three mice of each group were amplified andhybridized to GeneChip 430A 2.0 (Affymetrix). Target geneswere determined by using both ANOVA (P � 0.05) and

fold-change (�1.4-fold) with the assist of GENESPRING soft-ware (Silicon Genetics, Redwood City, CA) and the BULLFROGprogram (34). The results were confirmed by either Northernblot analysis or Q-PCR. For Q-PCR, 1 �g of RNA wasDNA-free treated, reverse-transcribed by using oligo-dT (Su-perscript II kit, Invitrogen), and then treated with RNase.Samples were run in triplicate by using SYBR green (AppliedBiosystems) and compared with levels of 36B4 as a control.The 0.58-kb (promoter I) and 1.3-kb (promoter II) humanACC� promoters were generated by PCR according to thepublished sequence (29) and cloned into the pGL-3 basicvector (Promega). These constructs, together with expressionvectors of PPAR� and RXR�, were transfected into HepG2cells in the presence of 0.1 �M dexamethasone.

We thank M. Nelson, Y. Zou, and H. Juguilon for technical help;Dr. M. Montminy and J. Sonoda for valuable comments; and E. Stevensand E. Ong for administrative assistance. C.H.L. was supported by theMilton Fund (Harvard University). P.O. was supported by Cellular andMolecular Genetics training grant [Department of Biology, University ofCalifornia at San Diego (UCSD)], the Chapman Fellowship, the Mo-lecular Neurobiology Grant at The Salk Institute, and the AchievementRewards for College Scientists Foundation. H.K. is supported byMarineBio 21 (Ministry of Maritime Affairs and Fisheries, Korea).R.M.E. is an investigator of the Howard Hughes Medical Institute at TheSalk Institute for Biological Studies and March of Dimes Chair inMolecular and Developmental Biology. This work was supported by theHoward Hughes Medical Institute, National Institutes of Health (NIH)Grant 5R37DK057978, the UCSD Specialized Center of Research(5P50HL56989), NIH Nuclear Receptor Signaling Axis Orphan Recep-tor Program Grant U19DK62434-01, and the Hilblom Foundation.

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