Table of Contents Author Guidelines Submit a Manuscript
PPAR Research
Volume 2010, Article ID 549101, 10 pages
Review Article

AMPK-Dependent Metabolic Regulation by PPAR Agonists

College of Pharmacy and Research Institute of Pharmaceutical Sciences, Seoul National University, Seoul 151-742, Republic of Korea

Received 22 April 2010; Accepted 25 June 2010

Academic Editor: Howard Glauert

Copyright © 2010 Woo Hyung Lee and Sang Geon Kim. This is an open access article distributed under the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


Comprehensive studies support the notion that the peroxisome proliferator-activated receptors, (PPARs), PPAR , PPAR , and PPAR , regulate cell growth, morphogenesis, differentiation, and homeostasis. Agonists of each PPAR subtype exert their effects similarly or distinctly in different tissues such as liver, muscle, fat, and vessels. It is noteworthy that PPAR or PPAR agonists have pharmacological effects by modulating the activity of AMPK, which is a key cellular energy sensor. However, the role of AMPK in the metabolic effects of PPAR agonists has not been thoroughly focused. Moreover, AMPK activation by PPAR agonists seems to be independent of the receptor activation. This intriguing action of PPAR agonists may account in part for the mechanistic basis of the therapeutics in the treatment of metabolic disease. In this paper, the effects of PPAR agonists on metabolic functions were summarized with particular reference to their AMPK activity regulation.

1. Introduction

The peroxisome proliferator-activated receptors (PPARs) are transcription factors that regulate diverse physiological and pathological processes including cell growth, morphogenesis, differentiation, and homeostasis [1]. PPARs are well-characterized receptors that belong to the nuclear hormone receptor superfamily: they were originally isolated as nuclear receptors that activate the proliferation of peroxisomes in the cell in 1990s [13]. PPARs consist of three isoforms (i.e., PPAR , PPAR / , and PPAR ), whose tissue distributions and functional roles are distinct [4]. After the discovery, intensive studies on the biology of PPARs and their modulation by synthetic ligands have been conducted. Thus, a myriad of research have identified natural or synthetic PPAR ligands as pharmaceutical agents in the treatment of metabolic disorders [1, 4] (Figure 1). Recently, it has been recognized that PPAR agonists have physiological effects by modulating the activity of AMP-activated protein kinase (AMPK), an important cellular energy sensor [59]. However, this action seems to be independent of PPAR receptor activation [5, 6, 9]. These findings suggest the concept that PPAR agonists exert their effects cooperatively or synergistically with their cellular partners or associated components. This paper focuses on the effects of PPAR agonists on metabolic functions with particular reference to AMPK activity modulation.

Figure 1: The chemical structures of PPAR activators.

2. Effects of PPAR Isoforms on Metabolic Functions

Like conventional nuclear receptors, each PPAR contains DNA-binding domain (DBD), ligand binding domain (LBD), and flexible hinge region. DBD has two zinc finger motifs whereas LBD contains 13 alpha helices and a small four-stranded beta sheet which composes a single domain [10]. Many nuclear receptors bind with retinoid X receptors (RXRs) as a modular heterodimeric partner, which also belong to members of the nuclear receptor superfamily; the RXRs possess a highly conserved central DNA-binding domain and less conserved ligand-binding domain [11]. Like other nuclear receptors, each PPAR forms a heterodimer with RXR as a permissive partner, and this complex transactivates target genes for physiological activity modulations. So, RXR acts as a supporting factor for strong DNA binding of PPAR. Ligand-mediated activation of the nuclear receptor homodimers or heterodimers involves their bindings to the DNA response element containing two core recognition sequences; this consensus DNA sequence present in the promoter regions of target genes is called “peroxisomal proliferator response element” (PPRE, 5 -AGGTCA-N-AGGTCA-3 ) [1]. Activation of PPARs with RXR contributes to diverse physiological processes through initiation of gene transcription, the protein products of which are required for glucose/fat metabolism, inflammation, vascular physiology, and muscle performance [14]. Hence, activated PPARs regulate genes involved in the adaptation of cells or organs to metabolic changes.

Three PPAR isoforms have different functional roles. PPAR (NR1C1), which is mainly expressed in the liver, heart, and kidney, shows high-catabolic rates of fatty acids and peroxisome-mediated activities [4]. Long-chain polyunsaturated fatty acids (especially, 20:5n-3 and 22:6n-3) and leukotriene B4 are natural ligands for PPAR whereas fibrates (i.e., hypolipidemics) and WY14643 (pirinixic acid, a nonsteroidal anti-inflammatory drug) are synthetic [12]. Activated PPAR promotes the expression of genes required for fatty acid and lipoprotein metabolism in mitochondria, peroxisomes, and endoplasmic reticulum [12]. Activation of PPAR by agonists or fatty acids induces peroxisomal proliferation, fatty acid oxidation, and the production of ketone bodies; PPAR stimulates the influx of fatty acids into the mitochondria via carnitine palmitoyltransferase 1 (CPT1) [12]. So, a deficiency of PPAR shows defects in hepatic fatty acid uptake and oxidation in an animal model [13]. PPAR activation increases plasma high-density lipoprotein (HDL) levels, transporting HDL particles from peripheral tissues to the liver with a decrease in plasma triglyceride (TG) level. Therefore, PPAR agonists suppress dyslipidemia observed in metabolic syndrome. Moreover, PPAR regulates energy balance in the body by modulating energy expenditure. Since uncoupling proteins (UCPs) contain PPREs in their gene promoter regions, PPAR activators induce UCP1 in brown adipose tissue, UCP2 in liver, and UCP3 in skeletal muscle [1416]. Recently, the role of PPAR in inflammation has emerged, implying that this receptor negatively regulates inflammatory responses [12].

Although PPAR / is ubiquitously expressed in most tissues, high level of PPAR / (NR1C2) is observed in skeletal muscle, suggesting that it is involved in energy metabolism [17]. PPAR / is engaged in membrane lipid synthesis/turnover, and cell proliferation and differentiation [17, 18]. Long-chain fatty acids and prostacyclin might serve the endogenous ligands of PPAR / . Erucic acid has a higher affinity for PPAR / than other PPAR subtypes [19]. Activated PPAR / promotes fatty acid oxidation in muscle and is thought to be engaged in the adaptation of muscle to fatty acid metabolism. Like PPAR , PPAR / regulates a series of genes involved in fatty acid catabolism and obesity (e.g., UCP3, CPT1 and malonyl-CoA decarboxylase) [1]. Wang et al. have shown that VP16-PPARδ transgenic mice with PPARδ activation in adipose tissue were resistant to high-fat diet-induced and genetically predisposed obesity and hyperlipidemia. By the same token, PPARδ knockout (KO) mice showed reduced energy uncoupling and were prone to obesity under high-fat diet feeding [20]. In addition, PPAR / is a sensor of very low-density lipoprotein (VLDL) in macrophages, and may play a role in fat storage [21]. In skeletal muscle, enforced expression of constitutively active PPAR / augmented oxidative muscle fibers and enhances running endurance [22, 23]. Consistently, PPAR / overexpression helps recover insulin resistance in obesity and enhance insulin action and glucose tolerance [22, 23].

PPAR (NR1C3) is expressed predominantly in the adipose tissue and, to a lesser extent, in the liver. PPAR exists as three forms ( 1, 2 and 3) by alternative splicing [1]. PPAR 1 is expressed in most tissues whereas PPAR 2 is present predominantly in adipose tissue. PPAR 3 shows high expression in macrophages, white adipose tissue, and large intestine. The results of gene KO study showed that the homozygous KO of PPAR impaired cardiac development, resulting in intrauterine death [24]. In addition, PPAR heterozygous KO mice exhibited impaired glucose homeostasis and adipocyte function, but showed increased leptin levels [25]. The PPAR activators such as free fatty acids (FFAs), eicosanoids and thiazolidinediones (TZD) induce physiological changes through target gene induction [1, 4]. PPAR activation in adipocytes sufficiently improved systemic insulin sensitization [26]. Adipose-specific PPAR activation by transgene expression and non-TZD PPAR agonist (AG035029) treatment prevented insulin resistance equivalent to TZD treatment [26]. These results suggest that fat-specific PPAR agonists may be novel candidates for diabetes. PPAR is significantly upregulated by the PPAR activators in the liver or under certain pathophysiological states (e.g., obesity) although its basal expression is rather low [27, 28]. Activation of PPAR improves insulin sensitivity in liver and muscle, decreases the intracellular lipid level in liver and muscle, and rescues insulin receptor signaling in type 2 diabetes [29]. Also, PPAR contributes to the balance between lipid influx and efflux in macrophages by upregulating target genes (e.g., fatty acid transporters and CD36) [28].

3. Regulatory Role of AMPK in Metabolic Functions

AMPK plays a critical role in sensing and regulating energy homeostasis in cells [30]. AMPK is a serine/threonine protein kinase which physiologically responds to the change in the cellular AMP to ATP ratio. AMPK activation regulates physiological and pathological responses in diverse tissues. AMPK activation induces fatty acid oxidation in liver and heart, inhibits hepatic lipogenesis and adipocyte differentiation, and stimulates glucose uptake in muscle [30, 31]. AMPK is composed of 3 subunits (α, β, and γ): a catalytic subunit ( 1 or 2) and two regulatory subunits ( 1 or 2 and 1, 2 or 3) [30]. AMPK activation is initiated with phosphorylation of threonine-172 in the catalytic domain of α subunit [31]. The γ subunit recognizes the AMP : ATP ratio because this has the AMP binding domain. Elevated level of AMP induces allosteric stimulation of AMPK. At present, two upstream kinases of AMPK have been discovered: LKB1 and Ca2+/calmodulin-dependent protein kinase kinase (CaMKK ) (Figure 2) [32, 33]. Transforming growth factor β-activated kinase-1 might be another one [34]. Both LKB1 and CaMKK directly phosphorylate thereonine-172 in AMPK subunit in an AMP-independent fashion. LKB1, also known as a tumor suppressor gene, is constitutively active. The upstream kinases of LKB1 may comprise protein kinase C (PKC)-ζ, protein kinase A, and ribosomal S6 kinase (RSK) [35, 36]. However, several studies also suggested that there may be LKB1-independent AMPK kinase based on the findings that constitutive AMPK activity could be detected even in LKB1-deficient mice or cells (e.g., HeLa and A549 cells) [37]. In addition, the result of CAMKK activity regulation by calcium/calmodulin (Ca2+/CaM) indicated that AMPK might be engaged in Ca2+ regulation in cells.

Figure 2: The regulatory role of AMPK in metabolic functions. CAMKK: CaM-dependent protein kinase kinase; eNOS: endothelial nitrogen oxide synthase; TSC1/2: tuberous sclerosis 1/2; mTOR: mammalian target of rapamycin; S6K1: p70 ribosomal S6 protein kinas; LXR: liver X receptor; ACC: acetyl-CoA carboxylase; CPT1: carnitine palmitoyltransferase 1.

The important role of AMPK in glucose metabolism has been investigated in cell or animal models. Insulin regulates glucose utilization in major organs that maintain serum glucose level, such as liver and muscle. When blood glucose content is elevated, insulin secreted from the pancreatic beta cell stimulates the storage of glucose in these organs [38]. The binding of insulin to insulin receptor in the plasma membrane transmits signals that induce diverse physiological responses. Autophosphorylation of insulin receptor triggered by insulin binding leads to activation of mammalian target of rapamycin (mTOR)-p70 ribosomal S6 kinase-1 (S6K1) via phosphatidylinositol 3-kinase (PI3K)/Akt pathway. This mTOR/S6K flow is linked to AMPK. Under the condition of starvation, AMPK phosphorylates tuberous sclerosis 2 (TSC2), which inhibits mTOR/S6K1 pathway [39]: phosphorylation of TSC2 by AMPK is critical in the process of mRNA translation and cell size regulation during energy deficiency (Figure 2). Thus, AMPK activation negatively regulates the mTOR/S6K1 pathway.

Glycogen synthase kinase-3 (GSK3 ), a Ser/Thr kinase, is constitutively activated in normal state. Phosphorylation at the serine 9 residue inactivates GSK3 , which may promote cell survival against ischemia/reperfusion injury by blocking mitochondrial permeability transition pore opening [40]. It has been shown that resveratrol treatment inhibited GSK3 activity downstream of AMP-activated protein kinase (AMPK) activation, and which was responsible for mitochondrial protection [41]. AMPK induces upregulation of the p53–p21 axis, which leads to G1 cell cycle arrest. It has been reported that AICAR treatment caused cell cycle arrest in various cell types (HepG2, mouse embryonic fibroblasts, and smooth muscle cells) [42]. AICAR promotes phosphorylation of tumor suppressor p53 (Ser15 in human), subsequently leading to p21 induction. In addition, the possibility that increases in p21, p27, and p53 by AICAR inhibit proliferation of several types of cancer cells has been reported [43]. In summary, these results demonstrated that AMPK plays a pivotal role in the regulation of cell cycle and survival by affecting several downstream signals.

AMPK activators mimic the actions of insulin in terms of gluconeogenesis, repressing glucose production [44]. Therefore, the agents that activate AMPK are beneficial in the treatment of insulin resistance and diabetes [44]. Moreover, insulin induces glucose transporter 4 (GLUT4) expression and its translocation, which facilitates glucose uptake into cells [38]. Physiologically, exercise enables GLUT4 to translocate into plasma membrane from vesicles through AMPK. The role of AMPK in exercise-induced glucose utilization is supported by the finding that treatment with aminoimidazole carboxamide ribonucleotide (AICAR), a direct AMPK activator, promoted glucose uptake, and GLUT4 translocation in skeletal muscle [45]. Consequently, 4 weeks of AICAR treatment stimulated metabolic genes and enhanced running endurance by 44% even in sedentary mice [23].

AMPK is involved in the maintenance of lipid and cholesterol homeostasis; it stimulates the -oxidation of fatty acids in mitochondria for lipid utilization [44]. AMPK inhibits the activity of acetyl-CoA carboxylase (ACC) through phosphorylation (Figure 2). Under normal condition, ACC inhibits CPT1 that transports fatty acids into mitochondria and increases fatty acid oxidation. Inactivation of ACC by AMPK helps promote fatty acid utilization, leading to fat burning in liver and muscle. Liver X receptor (LXR ) is the lipid sensor that promotes fatty acid synthesis and leads to hypertriglyceridemia. AMPK activation by metformin or dithiolethiones represses LXR activity via phosphorylating threonine residue(s) of AMPK [46]. Moreover, AMPK inhibits hepatic cholesterol synthesis by inhibiting HMG-CoA reductase, a rate-controlling enzyme of the mevalonate pathway. AMPK inhibits HMG-CoA reductase activity by phosphorylation, which reduces cholesterol levels (Figure 2) [47]. Similarly, AMPK activation attenuates TG synthesis via the inhibition of LXR activity in the liver, and thus results in an antisteatotic effect [46]. Of note, S6K1 activation reverses this effect of AMPK on LXR -SREBP-1c pathway, as mediated by the phosphorylation of LXR at serine residue.

Vascular endothelium is usually exposed to physical stress (e.g., blood pressure or shear stress) even under normal conditions. Consequently, damaged vasculature causes blood coagulation and recruitment of immune cells. In particular, free radical stress contributes to the pathologic processes of cardiovascular diseases (e.g., atherosclerosis and coronary heart disease) [48, 49]. Apoptosis of endothelial cells by shear stress or FFAs causes injury of endothelial cell monolayer, provokes the migration of vascular smooth muscle cells (VSMCs) into the intima, and facilitates plaque formation [50]. Altered outer environment confers proliferation and migration of VSMCs, which may be stimulated by the cytokines and growth factors secreted from accumulated immune cells in the plaque. Under these conditions, uncontrolled growth of VSMCs in conjunction with endothelial cell death is critical for the development of atherosclerosis [50]. In addition, the production of reactive oxygen species (H2O2 and ) is amplified by the activation of NAD(P)H oxidase, peroxidase, and cyclooxygenase in these cells. AMPK may affect vascular physiology. AMPK activators including rosiglitazone and pioglitazone suppress high-glucose-induced hyperactivity of NAD(P)H oxidase in human umbilical vein endothelial cells [51, 52]. In addition, hypoxia-activated AMPK stimulates Akt that phosphorylates and activates endothelial nitrogen oxide synthase (eNOS) (Serine 1177) (Figure 2) [53, 54]. Thus, eNOS contributes to the survival and function of endothelial cells through nitric oxide (NO) production. Since AMPK responds to external stress and regulate cellular homeostasis, its activation enables endothelial cells to survive against severe stress [53, 54].

4. The Link between PPAR Agonists and AMPK-Dependent Metabolic Functions

4.1. Energy Metabolism

The metabolic disorder is a constellation of impaired glucose/lipid metabolism, hypertension, obesity, diabetes, and cardiovascular diseases [38]. Major causes of the metabolic disorder include overweight, physical inactivity, and high-carbohydrate diet that cause the disturbance of energy metabolism. A variety of metabolic diseases are highly associated with insulin resistance, as defined by the desensitization of target cells to insulin. Insulin-resistant diabetic patients are at high risk for developing hepatic diseases. Also, peripheral insulin resistance is monitored in most patients with liver cirrhosis. The major causes of insulin resistance are genetic (~50%) and environment factors including obesity (~25%), and physical fitness (~25%) [55]. Since treatment of insulin resistance has beneficial effects on diabetes, dyslipidemia, obesity, and atherosclerosis, AMPK emerges as a therapeutic target for metabolic disorders [44].

Endurance exercise is the treatment recommended for patients with metabolic disease. Intriguingly, PPAR / agonists serve exercise mimetics, as do AMPK activators [23] (Figure 3). Because PPAR / shows high expression in skeletal muscle, treatment with PPAR / agonist (GW1516) reprogrammed gene expression involved in oxidative metabolism in this tissue [23]. In addition, GW1516 administration and exercise training exert synergistic effects, as shown by the improvement of running endurance in exercise-trained mice [23]. In this study, GW1516 and AICAR synergistically increased transcription of several oxidative genes in mice quadriceps (i.e., Scd1, ATP citrate lyase, hormone sensitive lipase, muscle fatty acid binding protein, Lpl, and Pdk4). AMPK directly interacted with PPAR / although it did not phosphorylate PPAR / ; AMPK may form a transcriptional complex with PPAR / , which would strengthen the receptor activity [23]. In addition, PPAR / activation by GW1516 induced SIRT1 gene transcription, which regulates body physiology and metabolism [56]. These results suggest that PPAR / activates AMPK probably because SIRT1 contributes to AMPK activation.

Figure 3: The effects of PPAR / activators on AMPK-dependent functions. AICAR: aminoimidazole carboxamide ribonucleotide; TFs: transcription factors.

PPAR activators including fenofibrate and WY14643 activate AMPK signaling pathway in a receptor-independent manner (Figure 4). However, the mechanisms of these ligands on AMPK activation differ from each other. Fenofibrate induces the phosphorylation and activation of AMPK via the induction of small heterodimer partner (SHP, an orphan nuclear receptor) and its target genes [5]. On the other hand, WY14643 treatment increased the expression of AMPK 1 and 2 mRNA, leading to increase in AMPK subunit phosphorylation and its enzymatic activity. However, the mechanism for this activation remains elusive [6].

Figure 4: The effects of PPAR activators on AMPK-dependent functions. eNOS: endothelial nitrogen oxide synthase; NF-κB: nuclear factor-κB; PAI-1: plasminogen activator inhibitor-1; SHP, small heterodimer partner.

PPAR agonists (i.e., TZDs) activate AMPK by phosphorylating AMPK independently of PPAR activity [79] (Figure 5). Troglitazone (10 mg/kg, i.p.) increased the phosphorylations of AMPK and ACC rapidly (15 min after treatment) in skeletal muscle, liver, and adipose tissue of intact rats [9]. Consistently, troglitazone caused two-fold increases in 2-deoxy-d-glucose uptake in skeletal muscle through AMPK activation. Clinically, pioglitazone also activates AMPK signaling and intensifies mitochondrial function and fat oxidation in the muscles of diabetic patients [29]. Collectively, TZDs stimulate adiponectin signals, activating AMPK, which regulates glucose metabolism and fat catabolism [29, 57].

Figure 5: The effects of PPAR activators on AMPK-dependent functions. ACC: acetyl-CoA carboxylase; CPT1: carnitine palmitoyltransferase 1; ROS: reactive oxygen species; SIRT1: sirtuin; Ψm: change in mitochondria membrane potential.
4.2. Metabolic Diseases

Insulin resistance is characterized as the condition that higher level of insulin is required for normal metabolic responses because normal level of insulin fails to achieve these responses in peripheral organs. Hepatic insulin resistance causes defects in glycogen synthesis/storage and disables glucose production/release [38, 58]. Insulin resistance reduces glucose uptake in skeletal muscle, whereas it hampers the normal insulin actions and enhances hydrolysis of stored TG in fat tissue. Insulin resistance and the consequent hyperinsulinemia develop into several metabolic syndromes such as type 2 diabetes mellitus, fatty liver disease, and atherosclerosis [38, 58]. Several TZDs have been shown to recover insulin resistance via AMPK activation [5962]. Rosiglitazone promotes AMPK-mediated insulin secretion via the phosphorylation of Kir6.2 subunit of the potassium (ATP) channel in -cells [62]. However, Prentiki and his colleagues reported that pioglitazone inhibits glucose-induced insulin secretion although its antidiabetic effect depended on AMPK [60]. Therefore, the effect of pioglitazone may attribute to improve hyperinsulinemia and preserve -cell function. Like pioglitazone, troglitazone restrains insulin hypersecretion at the high level of glucose and fatty acids, leading to the rescue of -cells from glucolipotoxicity [59]. Recently, BLX-1002, a novel TZD with no PPAR affinity, activates AMPK in -cell. BLX-1002 raises cytoplasmic Ca2+ and enhances glucose-induced insulin secretion at high glucose [61]. These results suggest that certain moiety(s) of TZDs is (are) responsible for AMPK activation independently of PPAR activation. In the view of glucose uptake, rosiglitazone remarkably enhanced AMPK-mediated glucose uptake and glycogen synthesis in muscle and adipose tissues [63]. Likewise, AICAR induced whole body glucose disposal (27%) and glucose infusion rate (44%), which represents improvement of insulin resistance. In cardiac muscle, PPAR and PPAR activators stimulated glucose uptake via AMPK [64]. However, GW1506 (a PPAR / activator) had no effects on glucose uptake in rat L6 skeletal muscle cells [65].

Nonalcoholic fatty liver disease (NAFLD) is associated with metabolic syndrome and insulin resistance [58]. NAFLD represents the initiation step of hepatic metabolic syndrome such as steatosis, steatohepatitis, and fibrosis. Insulin resistance induced mostly by obesity may cause the development of hepatic steatosis; hyperinsulinemia augments hepatic lipogenesis of TGs and fatty acids. Several AMPK activators such as metformin and TZDs contribute to not only insulin sensitivity enhancement, but intervention of hepatic steatosis [46, 66], suggesting that signals downstream of the components activated by the drugs with different modes of action merge to the same pathway. Treatment with either rosiglitazone or pioglitazone attenuated hepatic steatosis and inflammation in patients with nonalcoholic steatohepatitis (Figure 5) [66, 67]. In addition, rosiglitazone ameliorated alcoholic fatty liver via adiponectin/SIRT1/AMPK pathway in mice [57]. Nonetheless, TZDs must be used cautiously as adjuvant therapy for nonalcoholic steatohepatitis treatment since they may provoke congestive heart failure. In recent studies, the activation of hypoxia inducible factor-1 (HIF1 )-PPAR axis promoted fatty acid uptake and glycerolipid biosynthesis genes, leading to cardiac hypertrophy [68]. In C57BL/6 mice fed a methionine-deficient and choline-deficient (MCD) diet, fenofibrate had an effect to prevent the progressive fibrosing steatohepatitis. In this model, fenofibrate induced AMPK-mediated SHP gene expression, but reduced plasminogen activator inhibitor-1 (PAI-1) mRNA and protein expression [5]. Another PPAR agonist, WY14643 also ameliorated steatohepatitis along with decrease in the gene expression involved in fatty acid synthesis [5].

eNOS plays a role in the endothelium homeostasis through NO production [50]. It has been shown that PPAR agonists including fenofibrate and WY14643 stimulate eNOS activity and NO production in human umbilical vein endothelial cells in association with AMPK activation [69, 70]. In mouse endothelial cells, AMPK activity modulation by fenofibrate contributes to inhibiting NF-κB activity, implying that the agent might attenuate atherosclerosis development (Figure 4) [69]. Rosiglitazone also reduced glucose-induced oxidative stress and increases eNOS enzyme stimulation via AMPK activation [52, 71].

5. Concluding Remarks

PPAR agonists have diverse metabolic effects both in vitro and in vivo. The mechanisms of action include nutrients (especially glucose and lipid) metabolism and maintenance of energy homeostasis. Since AMPK is an important regulator in energy metabolism, it may be a key downstream target of PPARs, as indicated by the convergence of PPAR agonists’ actions on AMPK. Overall, the effects of PPAR agonists on AMPK-mediated metabolic functions may contribute to the recovery of insulin sensitivity or treatment of metabolic syndrome (Table 1).

Table 1: The effects of PPAR agonists on AMPK-dependent metabolic functions.


ACC:Acetyl-CoA carboxylase
AICAR:Aminoimidazole carboxamide ribonucleotide
AMP:Adenosine monophosphate
AMPK:AMP-activated protein kinase
ATP:Adenosine triphosphate
CAMKK:CaM-dependent protein kinase kinase
CPT1:Carnitine palmitoyltransferase 1
DBD:DNA biding domain
eNOS:Endothelial nitrogen oxide synthase
FFA:Free fatty acid
GLUT4:Glucose transporter 4
HDL:High-density lipoprotein
LBD:Ligand binding domain
LXR:Liver X receptor
mTOR:Mammalian target of rapamycin
NAFLD:Nonalcoholic fatty liver disease
NO:Nitric oxide
PAI-1:Plasminogen activator inhibitor-1
PI3K:Phosphoinositide 3-kinase
PPAR:Peroxisome proliferator-activated receptor
PPRE:Peroxisome proliferator response element
RXR:Retinoid X receptor
SHP:Small heterodimer partner
S6K:p70 ribosomal S6 protein kinas
TSC2:Tuberous sclerosis 2
UCP:Uncoupling protein
VLDL:Very low density lipoprotein
VSMC:Vascular smooth muscle cell.


This paper was supported by a Grant (10182KFDA992) from Korea Food & Drug Administration in 2010.


  1. W. Ahmed, O. Ziouzenkova, J. Brown et al., “PPARs and their metabolic modulation: new mechanisms for transcriptional regulation?” Journal of Internal Medicine, vol. 262, no. 2, pp. 184–198, 2007. View at Publisher · View at Google Scholar · View at Scopus
  2. I. Issemann and S. Green, “Activation of a member of the steroid hormone receptor superfamily by peroxisome proliferators,” Nature, vol. 347, no. 6294, pp. 645–650, 1990. View at Publisher · View at Google Scholar · View at Scopus
  3. C. Dreyer, G. Krey, H. Keller, F. Givel, G. Helftenbein, and W. Wahli, “Control of the peroxisomal β-oxidation pathway by a novel family of nuclear hormone receptors,” Cell, vol. 68, no. 5, pp. 879–887, 1992. View at Publisher · View at Google Scholar · View at Scopus
  4. C. Diradourian, J. Girard, and J.-P. Pégorier, “Phosphorylation of PPARs: from molecular characterization to physiological relevance,” Biochimie, vol. 87, no. 1, pp. 33–38, 2005. View at Publisher · View at Google Scholar · View at Scopus
  5. D. Chanda, C. H. Lee, Y.-H. Kim et al., “Fenofibrate differentially regulates plasminogen activator inhibitor-1 gene expression via adenosine monophosphate-activated protein kinase-dependent induction of orphan nuclear receptor small heterodimer partner,” Hepatology, vol. 50, no. 3, pp. 880–892, 2009. View at Publisher · View at Google Scholar · View at Scopus
  6. S. Liangpunsakul, S.-E. Wou, K. D. Wineinger et al., “Effects of WY-14,643 on the phosphorylation and activation of AMP-dependent protein kinase,” Archives of Biochemistry and Biophysics, vol. 485, no. 1, pp. 10–15, 2009. View at Publisher · View at Google Scholar · View at Scopus
  7. L. G. D. Fryer, A. Parbu-Patel, and D. Carling, “The anti-diabetic drugs rosiglitazone and metformin stimulate AMP-activated protein kinase through distinct signaling pathways,” Journal of Biological Chemistry, vol. 277, no. 28, pp. 25226–25232, 2002. View at Publisher · View at Google Scholar · View at Scopus
  8. A. K. Saha, P. R. Avilucea, J.-M. Ye, M. M. Assifi, E. W. Kraegen, and N. B. Ruderman, “Pioglitazone treatment activates AMP-activated protein kinase in rat liver and adipose tissue in vivo,” Biochemical and Biophysical Research Communications, vol. 314, no. 2, pp. 580–585, 2004. View at Publisher · View at Google Scholar · View at Scopus
  9. N. K. LeBrasseur, M. Kelly, T.-S. Tsao et al., “Thiazolidinediones can rapidly activate AMP-activated protein kinase in mammalian tissues,” American Journal of Physiology—Endocrinology and Metabolism, vol. 291, no. 1, pp. E175–E181, 2006. View at Publisher · View at Google Scholar · View at Scopus
  10. V. Zoete, A. Grosdidier, and O. Michielin, “Peroxisome proliferator-activated receptor structures: ligand specificity, molecular switch and interactions with regulators,” Biochimica et Biophysica Acta, vol. 1771, no. 8, pp. 915–925, 2007. View at Publisher · View at Google Scholar · View at Scopus
  11. M. Gianní, A. Tarrade, E. A. Nigro, E. Garattini, and C. Rochette-Egly, “The AF-1 and AF-2 domains of RARγ2 and RXRα cooperate for triggering the transactivation and the degradation of RARγ2/RXRα heterodimers,” Journal of Biological Chemistry, vol. 278, no. 36, pp. 34458–34466, 2003. View at Publisher · View at Google Scholar · View at Scopus
  12. M. Yoon, “The role of PPARα in lipid metabolism and obesity: focusing on the effects of estrogen on PPARα actions,” Pharmacological Research, vol. 60, no. 3, pp. 151–159, 2009. View at Publisher · View at Google Scholar · View at Scopus
  13. P. Costet, C. Legendre, J. Moré, A. Edgar, P. Galtier, and T. Pineau, “Peroxisome proliferator-activated receptor α-isoform deficiency leads to progressive dyslipidemia with sexually dimorphic obesity and steatosis,” Journal of Biological Chemistry, vol. 273, no. 45, pp. 29577–29585, 1998. View at Publisher · View at Google Scholar · View at Scopus
  14. M. J. Barberá, A. Schlüter, N. Pedraza, R. Iglesias, F. Villarroya, and M. Giralt, “Peroxisome proliferator-activated receptor α activates transcription of the brown fat uncoupling protein-1 gene. A link between regulation of the thermogenic and lipid oxidation pathways in the brown fat cell,” Journal of Biological Chemistry, vol. 276, no. 2, pp. 1486–1493, 2001. View at Publisher · View at Google Scholar · View at Scopus
  15. T. Nakatani, N. Tsuboyama-Kasaoka, M. Takahashi, S. Miura, and O. Ezaki, “Mechanism for peroxisome proliferator-activated receptor-α activator-induced up-regulation of UCP2 mRNA in rodent hepatocytes,” Journal of Biological Chemistry, vol. 277, no. 11, pp. 9562–9569, 2002. View at Publisher · View at Google Scholar · View at Scopus
  16. S. Brun, M. C. Carmona, T. Mampel et al., “Activators of peroxisome proliferator-activated receptor-α induce the expression of the uncoupling protein-3 gene in skeletal muscle: a potential mechanism for the lipid intake-dependent activation of uncoupling protein-3 gene expression at birth,” Diabetes, vol. 48, no. 6, pp. 1217–1222, 1999. View at Publisher · View at Google Scholar · View at Scopus
  17. G. D. Barish, V. A. Narkar, and R. M. Evans, “PPARδ: a dagger in the heart of the metabolic syndrome,” Journal of Clinical Investigation, vol. 116, no. 3, pp. 590–597, 2006. View at Publisher · View at Google Scholar · View at Scopus
  18. R. M. Evans, G. D. Barish, and Y.-X. Wang, “PPARs and the complex journey to obesity,” Nature Medicine, vol. 10, no. 4, pp. 355–361, 2004. View at Publisher · View at Google Scholar · View at Scopus
  19. T. E. Johnson, M. K. Holloway, R. Vogel et al., “Structural requirements and cell-type specificity for ligand activation of peroxisome proliferator-activated receptors,” Journal of Steroid Biochemistry and Molecular Biology, vol. 63, no. 1–3, pp. 1–8, 1997. View at Publisher · View at Google Scholar · View at Scopus
  20. Y.-X. Wang, C.-H. Lee, S. Tiep et al., “Peroxisome-proliferator-activated receptor δ activates fat metabolism to prevent obesity,” Cell, vol. 113, no. 2, pp. 159–170, 2003. View at Publisher · View at Google Scholar · View at Scopus
  21. A. Chawla, C.-H. Lee, Y. Barak et al., “PPARδ is a very low-density lipoprotein sensor in macrophages,” Proceedings of the National Academy of Sciences of the United States of America, vol. 100, no. 3, pp. 1268–1273, 2003. View at Publisher · View at Google Scholar · View at Scopus
  22. Y.-X. Wang, C.-L. Zhang, R. T. Yu et al., “Regulation of muscle fiber type and running endurance by PPARδ,” PLoS Biology, vol. 2, no. 10, article e294, 2004. View at Publisher · View at Google Scholar · View at Scopus
  23. V. A. Narkar, M. Downes, R. T. Yu et al., “AMPK and PPARδ agonists are exercise mimetics,” Cell, vol. 134, no. 3, pp. 405–415, 2008. View at Publisher · View at Google Scholar · View at Scopus
  24. A. Chawla, W. A. Boisvert, C.-H. Lee et al., “A PPARγ-LXR-ABCA1 pathway in macrophages is involved in cholesterol efflux and atherogenesis,” Molecular Cell, vol. 7, no. 1, pp. 161–171, 2001. View at Publisher · View at Google Scholar · View at Scopus
  25. N. Kubota, Y. Terauchi, H. Miki et al., “PPARγ mediates high-fat diet-induced adipocyte hypertrophy and insulin resistance,” Molecular Cell, vol. 4, no. 4, pp. 597–609, 1999. View at Publisher · View at Google Scholar · View at Scopus
  26. S. Sugii, P. Olson, D. D. Sears et al., “PPARγ activation in adipocytes is sufficient for systemic insulin sensitization,” Proceedings of the National Academy of Sciences of the United States of America, vol. 106, no. 52, pp. 22504–22509, 2009. View at Publisher · View at Google Scholar · View at Scopus
  27. J.-G. Jiang, C. Johnson, and R. Zarnegar, “Peroxisome proliferator-activated receptor γ-mediated transcriptional up-regulation of the hepatocyte growth factor gene promoter via a novel composite cis-acting element,” Journal of Biological Chemistry, vol. 276, no. 27, pp. 25049–25056, 2001. View at Publisher · View at Google Scholar · View at Scopus
  28. J. Han, D. P. Hajjar, X. Zhou, A. M. Gotto Jr., and A. C. Nicholson, “Regulation of peroxisome proliferator-activated receptor-γ-mediated gene expression a new mechanism of action for high density lipoprotein,” Journal of Biological Chemistry, vol. 277, no. 26, pp. 23582–23586, 2002. View at Publisher · View at Google Scholar · View at Scopus
  29. D. K. Coletta, A. Sriwijitkamol, E. Wajcberg et al., “Pioglitazone stimulates AMP-activated protein kinase signalling and increases the expression of genes involved in adiponectin signalling, mitochondrial function and fat oxidation in human skeletal muscle in vivo: a randomised trial,” Diabetologia, vol. 52, no. 4, pp. 723–732, 2009. View at Publisher · View at Google Scholar · View at Scopus
  30. M. C. Towler and D. G. Hardie, “AMP-activated protein kinase in metabolic control and insulin signaling,” Circulation Research, vol. 100, no. 3, pp. 328–341, 2007. View at Publisher · View at Google Scholar · View at Scopus
  31. R. Lage, C. Diéguez, A. Vidal-Puig, and M. López, “AMPK: a metabolic gauge regulating whole-body energy homeostasis,” Trends in Molecular Medicine, vol. 14, no. 12, pp. 539–549, 2008. View at Publisher · View at Google Scholar · View at Scopus
  32. A. Woods, S. R. Johnstone, K. Dickerson et al., “LKB1 is the upstream kinase in the AMP-activated protein kinase cascade,” Current Biology, vol. 13, no. 22, pp. 2004–2008, 2003. View at Publisher · View at Google Scholar · View at Scopus
  33. S. A. Hawley, D. A. Pan, K. J. Mustard et al., “Calmodulin-dependent protein kinase kinase-β is an alternative upstream kinase for AMP-activated protein kinase,” Cell Metabolism, vol. 2, no. 1, pp. 9–19, 2005. View at Publisher · View at Google Scholar · View at Scopus
  34. M. Momcilovic, S.-P. Hong, and M. Carlson, “Mammalian TAK1 activates Snf1 protein kinase in yeast and phosphorylates AMP-activated protein kinase in vitro,” Journal of Biological Chemistry, vol. 281, no. 35, pp. 25336–25343, 2006. View at Publisher · View at Google Scholar · View at Scopus
  35. D. R. Alessi, K. Sakamoto, and J. R. Bayascas, “LKB1-dependent signaling pathways,” Annual Review of Biochemistry, vol. 75, pp. 137–163, 2006. View at Publisher · View at Google Scholar · View at Scopus
  36. Z. Xie, Y. Dong, R. Scholz, D. Neumann, and M.-H. Zou, “Phosphorylation of LKB1 at serine 428 by protein kinase C-ζ is required for metformin-enhanced activation of the AMP-activated protein kinase in endothelial cells,” Circulation, vol. 117, no. 7, pp. 952–962, 2008. View at Publisher · View at Google Scholar · View at Scopus
  37. R. L. Hurley, K. A. Anderson, J. M. Franzone, B. E. Kemp, A. R. Means, and L. A. Witters, “The Ca2+/calmodulin-dependent protein kinase kinases are AMP-activated protein kinase kinases,” Journal of Biological Chemistry, vol. 280, no. 32, pp. 29060–29066, 2005. View at Publisher · View at Google Scholar · View at Scopus
  38. E. Bugianesi, A. J. McCullough, and G. Marchesini, “Insulin resistance: a metabolic pathway to chronic liver disease,” Hepatology, vol. 42, no. 5, pp. 987–1000, 2005. View at Publisher · View at Google Scholar · View at Scopus
  39. K. Inoki, T. Zhu, and K.-L. Guan, “TSC2 mediates cellular energy response to control cell growth and survival,” Cell, vol. 115, no. 5, pp. 577–590, 2003. View at Publisher · View at Google Scholar · View at Scopus
  40. S.-S. Park, H. Zhao, R. A. Mueller, and Z. Xu, “Bradykinin prevents reperfusion injury bytargeting mitochondrial permeability transition pore through glycogen synthase kinase 3β,” Journal of Molecular and Cellular Cardiology, vol. 40, no. 5, pp. 708–716, 2006. View at Publisher · View at Google Scholar · View at Scopus
  41. S. M. Shin, I. J. Cho, and S. G. Kim, “Resveratrol protects mitochondria against oxidative stress through AMP-activated protein kinase-mediated glycogen synthase kinase-3β inhibition downstream of poly(ADP-ribose) polymerase-LKB1 pathway,” Molecular Pharmacology, vol. 76, no. 4, pp. 884–895, 2009. View at Publisher · View at Google Scholar · View at Scopus
  42. H. Motoshima, B. J. Goldstein, M. Igata, and E. Araki, “AMPK and cell proliferation–AMPK as a therapeutic target for atherosclerosis and cancer,” Journal of Physiology, vol. 574, no. 1, pp. 63–71, 2006. View at Publisher · View at Google Scholar · View at Scopus
  43. R. Rattan, S. Giri, A. K. Singh, and I. Singh, “5-aminoimidazole-4-carboxamide-1-β-D-ribofuranoside inhibits cancer cell proliferation in vitro and in vivo via AMP-activated protein kinase,” Journal of Biological Chemistry, vol. 280, no. 47, pp. 39582–39593, 2005. View at Publisher · View at Google Scholar · View at Scopus
  44. B. B. Zhang, G. Zhou, and C. Li, “AMPK: an emerging drug target for diabetes and the metabolic syndrome,” Cell Metabolism, vol. 9, no. 5, pp. 407–416, 2009. View at Publisher · View at Google Scholar · View at Scopus
  45. E. J. Kurth-Kraczek, M. F. Hirshman, L. J. Goodyear, and W. W. Winder, “5' AMP-activated protein kinase activation causes GLUT4 translocation in skeletal muscle,” Diabetes, vol. 48, no. 8, pp. 1667–1671, 1999. View at Publisher · View at Google Scholar · View at Scopus
  46. S. H. Hwahng, S. H. Ki, E. J. Bae, H. E. Kim, and S. G. Kim, “Role of adenosine monophosphate-activated protein kinase-p70 ribosomal S6 kinase-1 pathway in repression of liver X receptor-alpha-dependent lipogenic gene induction and hepatic steatosis by a novel class of dithiolethiones,” Hepatology, vol. 49, no. 6, pp. 1913–1925, 2009. View at Publisher · View at Google Scholar · View at Scopus
  47. D. Carling, P. R. Clarke, V. A. Zammit, and D. G. Hardie, “Purification and characterization of the AMP-activated protein kinase. Copurification of acetyl-CoA carboxylase kinase and 3-hydroxy-3-methylglutaryl-CoA reductase kinase activities,” European Journal of Biochemistry, vol. 186, no. 1-2, pp. 129–136, 1989. View at Publisher · View at Google Scholar · View at Scopus
  48. B. Fisslthaler and I. Fleming, “Activation and signaling by the AMP-activated protein kinase in endothelial cells,” Circulation Research, vol. 105, no. 2, pp. 114–127, 2009. View at Publisher · View at Google Scholar · View at Scopus
  49. D. Nagata and Y. Hirata, “The role of AMP-activated protein kinase in the cardiovascular system,” Hypertension Research, vol. 33, no. 1, pp. 22–28, 2010. View at Publisher · View at Google Scholar · View at Scopus
  50. J. Gutierrez, S. W. Ballinger, V. M. Darley-Usmar, and A. Landar, “Free radicals, mitochondria, and oxidized lipids: the emerging role in signal transduction in vascular cells,” Circulation Research, vol. 99, no. 9, pp. 924–932, 2006. View at Publisher · View at Google Scholar · View at Scopus
  51. J. C. Choy, D. J. Granville, D. W. C. Hunt, and B. M. McManus, “Endothelial cell apoptosis: biochemical characteristics and potential implications for atherosclerosis,” Journal of Molecular and Cellular Cardiology, vol. 33, no. 9, pp. 1673–1690, 2001. View at Publisher · View at Google Scholar · View at Scopus
  52. G. Ceolotto, A. Gallo, I. Papparella et al., “Rosiglitazone reduces glucose-induced oxidative stress mediated by NAD(P)H oxidase via AMPK-dependent mechanism,” Arteriosclerosis, Thrombosis, and Vascular Biology, vol. 27, no. 12, pp. 2627–2633, 2007. View at Publisher · View at Google Scholar · View at Scopus
  53. K. Fujisawa, T. Nishikawa, D. Kukidome et al., “TZDs reduce mitochondrial ROS production and enhance mitochondrial biogenesis,” Biochemical and Biophysical Research Communications, vol. 379, no. 1, pp. 43–48, 2009. View at Publisher · View at Google Scholar · View at Scopus
  54. D. Nagata, M. Mogi, and K. Walsh, “AMP-activated protein kinase (AMPK) signaling in endothelial cells is essential for angiogenesis in response to hypoxic stress,” Journal of Biological Chemistry, vol. 278, no. 33, pp. 31000–31006, 2003. View at Publisher · View at Google Scholar · View at Scopus
  55. C. Bogardus, S. Lillioja, D. M. Mott, C. Hollenbeck, and G. Reaven, “Relationship between degree of obesity and in vivo insulin action in man,” American journal of physiology, vol. 248, no. 3, part 1, pp. E286–E291, 1985. View at Google Scholar · View at Scopus
  56. M. Okazaki, Y. Iwasaki, M. Nishiyama et al., “PPARβ/δ regulates the human SIRT1 gene transcription via Sp1,” Endocrine Journal, vol. 57, no. 5, pp. 403–413, 2010. View at Publisher · View at Google Scholar · View at Scopus
  57. Z. Shen, X. Liang, C. Q. Rogers, D. Rideout, and M. You, “Involvement of adiponectin-SIRT1-AMPK signaling in the protective action of rosiglitazone against alcoholic fatty liver in mice,” American Journal of Physiology—Gastrointestinal and Liver Physiology, vol. 298, no. 3, pp. G364–G374, 2010. View at Publisher · View at Google Scholar · View at Scopus
  58. K. Qureshi and G. A. Abrams, “Metabolic liver disease of obesity and role of adipose tissue in the pathogenesis of nonalcoholic fatty liver disease,” World Journal of Gastroenterology, vol. 13, no. 26, pp. 3540–3553, 2007. View at Google Scholar · View at Scopus
  59. X. Wang, L. Zhou, L. Shao et al., “Troglitazone acutely activates AMP-activated protein kinase and inhibits insulin secretion from beta cells,” Life Sciences, vol. 81, no. 2, pp. 160–165, 2007. View at Publisher · View at Google Scholar · View at Scopus
  60. J. Lamontagne, É. Pepin, M.-L. Peyot et al., “Pioglitazone acutely reduces insulin secretion and causes metabolic deceleration of the pancreatic β-cell at submaximal glucose concentrations,” Endocrinology, vol. 150, no. 8, pp. 3465–3474, 2009. View at Publisher · View at Google Scholar · View at Scopus
  61. F. Zhang, D. Dey, R. Bränstrom et al., “BLX-1002, a novel thiazolidinedione with no PPAR affinity, stimulates AMP- Activated protein kinase activity, raises cytosolic Ca2+, and enhances glucose- Stimulated insulin secretion in a PI3K-dependent manner,” American Journal of Physiology - Cell Physiology, vol. 296, no. 2, pp. C346–C354, 2009. View at Publisher · View at Google Scholar · View at Scopus
  62. T.-J. Chang, W.-P. Chen, C. Yang et al., “Serine-385 phosphorylation of inwardly rectifying K+ channel subunit (Kir6.2) by AMP-dependent protein kinase plays a key role in rosiglitazone-induced closure of the KATP channel and insulin secretion in rats,” Diabetologia, vol. 52, no. 6, pp. 1112–1121, 2009. View at Publisher · View at Google Scholar · View at Scopus
  63. J.-M. Ye, N. Dzamko, A. J. Hoy, M. A. Iglesias, B. Kemp, and E. Kraegen, “Rosiglitazone treatment enhances acute AMP-activated protein kinase-mediated muscle and adipose tissue glucose uptake in high-fat-fed rats,” Diabetes, vol. 55, no. 10, pp. 2797–2804, 2006. View at Publisher · View at Google Scholar · View at Scopus
  64. X. Xiao, G. Su, S. N. Brown, L. Chen, J. Ren, and P. Zhao, “Peroxisome proliferator-activated receptors γ and α agonists stimulate cardiac glucose uptake via activation of AMP-activated protein kinase,” Journal of Nutritional Biochemistry, vol. 21, no. 7, pp. 621–626, 2010. View at Publisher · View at Google Scholar · View at Scopus
  65. N. Dimopoulos, M. Watson, C. Green, and H. S. Hundal, “The PPARδ agonist, GW501516, promotes fatty acid oxidation but has no direct effect on glucose utilisation or insulin sensitivity in rat L6 skeletal muscle cells,” FEBS Letters, vol. 581, no. 24, pp. 4743–4748, 2007. View at Publisher · View at Google Scholar · View at Scopus
  66. V. Ratziu, F. Charlotte, C. Bernhardt et al., “Long-term efficacy of rosiglitazone in nonalcoholic steatohepatitis: results of the Fatty Liver Improvement by Rosiglitazone Therapy (FLIRT 2) extension trial,” Hepatology, vol. 51, no. 2, pp. 445–453, 2010. View at Publisher · View at Google Scholar · View at Scopus
  67. A. Gastaldelli, S. A. Harrison, R. Belfort-Aguilar et al., “Importance of changes in adipose tissue insulin resistance to histological response during thiazolidinedione treatment of patients with nonalcoholic steatohepatitis,” Hepatology, vol. 50, no. 4, pp. 1087–1093, 2009. View at Publisher · View at Google Scholar · View at Scopus
  68. J. Krishnan, M. Suter, R. Windak et al., “Activation of a HIF1α-PPARγ axis underlies the integration of glycolytic and lipid anabolic pathways in pathologic cardiac hypertrophy,” Cell Metabolism, vol. 9, no. 6, pp. 512–524, 2009. View at Publisher · View at Google Scholar · View at Scopus
  69. H. Murakami, R. Murakami, F. Kambe et al., “Fenofibrate activates AMPK and increases eNOS phosphorylation in HUVEC,” Biochemical and Biophysical Research Communications, vol. 341, no. 4, pp. 973–978, 2006. View at Publisher · View at Google Scholar · View at Scopus
  70. T. Okayasu, A. Tomizawa, K. Suzuki, K.-I. Manaka, and Y. Hattori, “PPARα activators upregulate eNOS activity and inhibit cytokine-induced NF-κB activation through AMP-activated protein kinase activation,” Life Sciences, vol. 82, no. 15-16, pp. 884–891, 2008. View at Publisher · View at Google Scholar · View at Scopus
  71. J. G. Boyle, P. J. Logan, M.-A. Ewart et al., “Rosiglitazone stimulates nitric oxide synthesis in human aortic endothelial cells via AMP-activated protein kinase,” Journal of Biological Chemistry, vol. 283, no. 17, pp. 11210–11217, 2008. View at Publisher · View at Google Scholar · View at Scopus
  72. S. H. Caldwell, E. E. Hespenheide, J. A. Redick, J. C. Iezzoni, E. H. Battle, and B. L. Sheppard, “A pilot study of a thiazolidinedione, troglitazone, in nonalcoholic steatohepatitis,” American Journal of Gastroenterology, vol. 96, no. 2, pp. 519–525, 2001. View at Publisher · View at Google Scholar · View at Scopus