The mechanisms of the improvement of glucose homeostasis through angiotensin receptor blockers are not fully elucidated in hypertensive patients. We investigated the effects of telmisartan on insulin signaling and glucose uptake in cultured myotubes and skeletal muscle from wild-type and muscle-specific peroxisome proliferator–activated receptor (PPAR) δ knockout (MCK-PPARδ−/−) mice. Telmisartan increased PPARδ expression and activated PPARδ transcriptional activity in cultured C2C12 myotubes. In palmitate-induced insulin-resistant C2C12 myotubes, telmisartan enhanced insulin-stimulated Akt and Akt substrate of 160 kDa (AS160) phosphorylation as well as Glut4 translocation to the plasma membrane. These effects were inhibited by antagonizing PPARδ or phosphatidylinositol-3 kinase, but not by PPARγ and PPARα inhibition. Palmitate reducing the insulin-stimulated glucose uptake in C2C12 myotubes could be restored by telmisartan. In vivo experiments showed that telmisartan treatment reversed high-fat diet–induced insulin resistance and glucose intolerance in wild-type mice but not in MCK-PPARδ−/− mice. The protein levels of PPARδ, phospho-Akt, phospho-AS160, and Glut4 translocation to the plasma membrane in the skeletal muscle on insulin stimulation were reduced by high-fat diet and were restored by telmisartan administration in wild-type mice. These effects were absent in MCK-PPARδ−/− mice. These findings implicate PPARδ as a potential therapeutic target in the treatment of hypertensive subjects with insulin resistance.
The underlying metabolic causes of type 2 diabetes are the combination of insulin resistance and defective secretion of insulin by pancreatic β-cells. Insulin resistance typically precedes the onset of type 2 diabetes (1) and is commonly accompanied by other cardiovascular risk factors, such as dyslipidemia, hypertension, and metabolic syndrome (2). Several large clinical trials demonstrate that angiotensin-converting enzyme inhibitors or angiotensin II receptor blockers (ARBs) can significantly reduce the incidence of new cases of type 2 diabetes in patients at high risk compared with other antihypertensive therapies (3). However, the mechanisms involved in improved glucose homeostasis through ARBs are not completely understood.
Several recent studies show that ARBs exert beneficial effects on lipid and glucose metabolism that involve more than just their ability to block the angiotensin II receptor (2). These may include enhancing blood flow through the microcirculation of skeletal muscle (4) and increasing plasma adiponectin concentration (5). In addition, several ARBs, including telmisartan (TM), have been found to effectively activate the peroxisome proliferator–activated receptor (PPAR) γ (6,7). PPAR isoforms display tissue-specific expression and gene-regulatory profiles. PPARγ is a key regulator of adipocyte differentiation and adipose insulin sensitivity (8,9), but it is expressed at extremely low levels, if at all, in skeletal muscle. In contrast, PPARδ (also referred to as PPARβ) is expressed in a wide variety of tissues, with high levels in skeletal muscle (10). Recent studies show a crucial role of PPARδ in skeletal muscle glucose metabolism and insulin action. Krämer et al. (11) showed that activation of PPARδ results in a direct increase of fatty acid transport and glucose uptake and promotes lipid and glucose metabolism and gene expression in primary cultured human skeletal muscle cells (12,13). Muscle-specific PPARδ-transgenic mice were used to establish the role of PPARδ in whole-body glucose homeostasis. Schuler et al. (14) showed that mice in which PPARδ is selectively ablated in skeletal muscle myocytes exhibit fiber-type switching, obesity, and type 2 diabetes, demonstrating that PPARδ is instrumental for peripheral insulin sensitivity.
The PPARδ-specific agonist GW501516 improves glucose tolerance and reduces plasma glucose and insulin levels in several animal models (15,16). Therefore, activation of PPARδ may offer an effective strategy to improve glucose homeostasis. However, the safety issues about this pharmacological agonist are still highly controversial (17,18). Thus, it is important to know whether ARBs, such as TM, affect PPARδ activity. Given the importance of skeletal muscle insulin resistance in the development of type 2 diabetes, we hypothesized that TM may affect glucose metabolism in skeletal muscle by activating PPARδ. Here, we present evidences supporting that TM as a bona fide ligand of PPARδ and its activation on phosphatidylinositol 3-kinase (PI3K) pathway are key mechanisms of enhancing insulin sensitivity and glucose uptake in skeletal muscle.
RESEARCH DESIGN AND METHODS
TM, palmitate, PPARγ inhibitor GW9662, PPARδ inhibitor GSK0660, PPARα inhibitor GW6471, and PI3K inhibitor LY294002 were all purchased from Sigma-Aldrich (St. Louis, MO).
Generation of muscle-specific PPARδ knockout mice.
Transgenic mice having the Cre recombinase gene driven by the muscle creatine kinase (MCK-Cre) promoter were purchased from The Jackson Laboratory (stock number 006475). Cre activity is observed in skeletal muscle. Mice possess loxP sites on either side of exon 4 of PPARδ gene (PPARδflox/flox) were also purchased from The Jackson Laboratory (stock number 005897). Mice with hemizygous MCK-Cre and homozygous PPARδflox allele are viable, fertile, and normal in size. Breeding of these two types of mice yielded Cre:PPARδflox/+ mice. Then, breeding of Cre:PPARδflox/+ mice with PPARδflox/flox mice yielded Cre:PPARδflox/flox mice, which have PPARδ-specific knockout in skeletal muscles (MCK-PPARδ−/−). The PPARδflox/flox littermates were used as control mice (wild-type [WT]) (19). DNA prepared from tail biopsy samples was used for genotyping by PCR using the following primers: for MCK-Cre, 5′-GTG AAA CAG CAT TGC TGT CAC TT-3′ (primer 1) and 5′-TAA GTC TGA ACC CGG TCT GC-3′ (primer 2) were used; and for PPARδflox, 5′-GAG CCG CCT CTC GCC ATC CTT TCA G-3′ (primer 1), 5′-GGC GTG GGG ATT TGC CTG CTT CA-3′ (primer 2), and 5′-GTC GAG AAG TAC TAG TGG CCA CGT GG-3′ (primer 3) were used.
Animals and experimental procedure.
Mice were housed in cages at a controlled temperature (22 ± 1°C) and relative humidity (55 ± 15%) in a 12-h light/12-h dark cycle. They were supplied with standard laboratory chow and tap water ad libitum until 6 weeks of age. Then, a high-fat diet (HFD) was supplied for 20 weeks to induce insulin resistance in the mice (20). After undergoing intraperitoneal glucose tolerance test (IPGTT) and intraperitoneal insulin tolerance test (IPITT), the mice were randomly given normal chow diet or HFD without (control or high-fat [HF] group, n = 6) or with TM (TM group or HF+TM group, n = 6) for another 10 weeks. TM (5 mg/kg body weight) was administrated by mixing into the normal chow diet or HFD. The mice kept on regular rodent chow were considered the control (control group, n = 6) throughout the experiment. The food intake was measured every day. Body weights were measured every 2 weeks. At the end of experiment, the mice were subjected to IPGTT and IPITT again. Then, the mice were killed after fasting for 14 h. The gastrocnemius muscle was removed and stored at −70°C for Western blot analysis. Blood was collected from the carotid arteries, and plasma was separated immediately. Plasma levels of cholesterol, triglycerides, and insulin were measured using commercially available assay kits (Applygen Technologies, Beijing, China). All of the experimental procedures were performed in accordance with protocols approved by the Institutional Animal Care and Research Advisory Committee.
IPGTT and IPITT.
IPGTT was performed in mice as previously described (21). After an overnight fast (14 h), glucose (2 g/kg body weight) was administered via injection into the peritoneal cavity, and blood was drawn from the tail vein at 0, 30, 60, and 120 min after glucose administration. Blood glucose levels were determined using the OneTouch Ultra blood glucose meter (LifeScan). IPITT was evaluated in fed mice on a different day. Humulin R (0.75 units/kg body weight; Eli Lilly and Co.) in sterile saline was administered intraperitoneally. Glucose levels were determined at 0, 15, 30, and 60 min after insulin injection.
Acute insulin-stimulation test.
Fasted mice were administered an injection of saline or insulin (5 units/kg of body weight). Gastrocnemius muscles were collected 5 min after the injection. The phosphorylation of Akt and AS160 was detected by Western blotting in freshly prepared muscle tissue homogenates. Glut4 protein was also detected in plasma membrane homogenates (22).
Glucose uptake assay.
The glucose uptake assay was conducted as described previously (23), with modifications. Briefly, myotubes were washed with Krebs-Ringer phosphate (KRP) buffer containing 0.05% BSA and incubated at 37°C for 30 min in KRP buffer. Then they were stimulated with insulin in KRP buffer for 10 min at 37°C or were left unstimulated. Cells were further incubated in KRP buffer containing [3H]-2-deoxyglucose (1 µCi/mL; PerkinElmer, Boston, MA) in 0.1 mmol/L of unlabeled 2-deoxyglucose for 15 min. Cells were washed in ice-cold PBS three times and lysed with 200 µL distilled water. A 100-µL aliquot of each sample was counted with liquid scintillation, and the rest of the sample was used to determine protein concentration by the Bradford assay. The radioactivity was normalized to the protein concentration.
C2C12 cell culture and treatment.
Mouse C2C12 myoblasts (ATCC, Manassas, VA) were maintained in Dulbecco's modified Eagle's medium (DMEM; Gibco, Grand Island, NY) with 10% FBS (Gibco) at 37°C in 5% CO2/95% O2 humidified air and differentiated in DMEM with 2% horse serum (Gibco) after reaching confluence. After 4 days, the myoblasts were differentiated into myotubes. Myotubes were incubated with 0.75 mmol/L palmitate for 18 h in DMEM containing 2% BSA and 10% FBS to induce insulin resistance, according to the method described by Wang et al. (24). Cells were then treated with TM dissolved in DMSO (dimethyl sulfoxide, 10 μmol/L) or DMSO alone in the presence or absence of indicated inhibitors for 24 h. At the end of treatment, cells were stimulated with 100 nmol/L insulin or PBS for 30 min before total protein or membrane protein were extracted.
Transfection and luciferase assay.
PPAR responsive element (PPRE) activity was determined by transactivation assays in C2C12 myotubes. Cells were plated in 48-well dishes. Lipofectamine LTX Reagent (Invitrogen, Carlsbad, CA) was used to transfect them with 300 ng luciferase reporter pTK-PPREx3-luc and 150 ng pRL-TK, a renilla luciferase reporter vector as an internal control, with or without cotransfection of 150 ng of the full-length coding region of the rat PPARδ expression plasmid pAdTrack-CMV-PPARδ, which was generated as described (25). An empty pAdTrack expression vector as the control for PPARδ expression vector. After 5 h, transfection medium was replaced by DMEM containing 10% FBS. Cells were then treated with the indicated concentrations of TM in the presence or absence of the indicated inhibitors for an additional 24 h. Cells were assayed for luciferase and renilla activity using the Dual-GloTM Luciferase Assay System (Promega, Madison, WI) and Varioskan Flash Type 3001 (Thermo Electron Corp., Waltham, MA). The luciferase activity was normalized to renilla activity in each sample.
Western blot analyses.
Western blots of glyceraldehyde-3-phosphate dehydrogenase (GAPDH), PPARδ, PPARγ, PPARα, Akt, phospho-Akt, Akt substrate of 160 kDa (AS160), phospho-AS160, and Glut4 (Santa Cruz Inc., Santa Cruz, CA) were performed as reported (20). The positive control of Glut4 antibody (SC-2243) was provided by Santa Cruz Inc. (Supplementary Data).
Immunofluorescence was routinely performed using antibodies against Glut4 (Santa Cruz Inc.). The images were collected using a Nikon TE2000-U inverted fluorescence microscope (Nikon, Tokyo, Japan) and total internal reflection microscopy (Nikon) (26) (Supplementary Data).
The data are presented as the means ± SEM. All analyses were performed with SPSS (version 13.0). Statistical significance of differences between mean values was assessed by Student t test or one-way ANOVA with Bonferroni multiple comparison post hoc tests as appropriate. Two-tailed P < 0.05 was considered statistically significant.
Effect of TM on PPARδ expression and transcriptional activity in C2C12 myotubes.
PPRE activity induced by TM was examined in C2C12 myotubes transfected with a luciferase reporter construct containing three tandem repeats of PPRE (pTK-PPREx3-luc). In C2C12 myotubes cotransfected with PPARδ and pTK-PPREx3-luc, TM induced a time-dependent and dose-dependent enhancement of luciferase activity, which was associated with an upregulation of PPARδ protein expression (Fig. 1A and B). PPARδ activity and protein expression were slightly changed by acute exposure to TM (15 min or 1 h), whereas PPARδ activity and protein expression were significantly increased by chronic TM treatment for 12 h or 24 h (Fig. 1A). TM dose-dependently activated PPARδ, but TM at 30 μmol/L reduced the PPARδ transcriptional activity and protein expression compared with its low doses (Fig. 1B), which could be attributable to the cell toxic effect of high concentration of TM on transcriptional activity (27,28). As shown in Fig. 1C, all three PPAR isoform antagonists significantly inhibited the TM-induced activity. However, the combination of PPARδ and PPARγ antagonists (GSK0660 and GW9662) showed no additive effects. Treatment with TM (10 μmol/L) significantly increased the luciferase activity and PPARδ protein expressions in transfected C2C12 myotubes (Fig. 1D). Furthermore, it showed that incubation with 10 μmol/L TM for 24 h significantly upregulated PPARδ, PPARγ, and PPARα expression in C2C12 myotubes (Fig. 1E). These data strongly indicate that TM enhances the expression and transcriptional activity of PPARδ in C2C12 myotubes.
TM improves insulin signaling and glucose uptake through the PPARδ/PI3K pathway.
To examine the effect of TM on insulin resistance and its underlying mechanisms, we examined the insulin signaling pathway by measuring Akt, AS160 phosphorylation (29), and plasma membrane Glut4 expression (30) in insulin-sensitive (without palmitate) and palmitate-induced insulin-resistant C2C12 myotubes. In C2C12 myotubes not exposed to palmitate, the phosphorylation of Akt and AS160 and plasma membrane Glut4 expression were increased by ∼10-fold after acute insulin stimulation. However, in palmitate-treated insulin-resistant cells, the insulin-induced phosphorylation of Akt and AS160 and plasma membrane Glut4 expression were weak, which could be enhanced by TM treatment. Administration of TM did not restore the phosphorylated Akt or AS160, and it stimulate Glut4 translocation in C2C12 myotubes to the same levels compared with those not exposed to palmitate (Fig. 2A and B). Furthermore, these effects of TM on expression of phospho-Akt, phospho-AS160, and plasma membrane Glut4 were abolished in the presence of either PI3K inhibitor LY294002 or PPARδ inhibitor GSK0660. However, PPARγ inhibitor GW9662 and PPARα inhibitor GW6471 did not block the effects of TM (Fig. 2C and D). These results suggest that TM promotes Akt and AS160 phosphorylation and Glut4 translocation in insulin-resistant C2C12 myotubes in a PPARδ/PI3K–dependent, but not in a PPARγ-dependent or PPARα-dependent, manner.
It is well-known that insulin stimulation induces Glut4 translocation to the cellular membrane and thus facilitates glucose uptake in skeletal muscle (30–32). In palmitate-induced insulin-resistant C2C12 myotubes, insulin had little effect on Glut4 translocation to the membrane, as shown by immunofluorescence (Fig. 3A and B). However, cotreatment with TM markedly enhanced the insulin-stimulated Glut4 translocation and expression, and this effect was attenuated in the presence of PPARδ inhibitor or PI3K inhibitor (Fig. 3A and B). We further examined the glucose uptake in C2C12 and palmitate-treated primary cultured myotubes from mice. Insulin stimulation significantly increased in glucose uptake of cells without palmitate treatment but had little effect on myotubes with palmitate (Fig. 3C and Supplementary Fig. 1). TM significantly increased the insulin-stimulated glucose uptake in C2C12 and mice myotubes in the presence of palmitate (Fig. 3C and Supplementary Fig. 1A).
TM treatment reverses insulin resistance of skeletal muscle in mice.
To verify the effect of TM on glucose homeostasis in vivo, we first established insulin resistance in mice fed a HFD for 20 weeks. The glucose tolerance and insulin sensitivity were both significantly impaired in HFD-fed mice, as detected by IPGTT and IPITT, respectively (Supplementary Fig. 2A and B). Next, these insulin-resistant mice were randomly divided into HF group and HF plus TM group for another 10 weeks. Both glucose tolerance and insulin sensitivity in the HF plus TM group were significantly improved, whereas those in the HF group were still significantly impaired compared with control mice fed a regular diet. However, neither the glucose intolerance nor the insulin resistance was improved in MCK-PPARδ−/− mice fed the HFD plus TM (Fig. 4A and B).
Mice fed a HFD had higher levels of plasma insulin, triglycerides, and cholesterol compared with the control mice fed the regular diet. TM significantly reduced these elevations of blood parameters in WT mice fed a HFD (Fig. 5A and B). In addition, chronic TM treatment prevented HFD-induced obesity in WT mice (Fig. 5C). The average food intake was similar between all groups of mice (Fig. 5D). Taken together, these findings indicate that long-term TM treatment significantly attenuates HFD-induced insulin resistance, hyperinsulinemia, hyperlipidemia, and obesity through stimulation of PPARδ activity and protein expression.
Chronic TM treatment enhances PPARδ-mediated insulin signaling proteins and Glut4 in skeletal muscle.
To further investigate the underlying mechanism of TM action in vivo, we detected the protein expression of PPARδ, PPARγ, PPARα, and insulin-signaling proteins in the gastrocnemius muscle tissue of WT and MCK-PPARδ−/− mice with or without acute insulin stimulation. As shown in Fig. 6A and C, WT mice fed a HFD had a significantly lower PPARδ level in skeletal muscle, whereas this reduction of PPARδ was markedly reversed by TM treatment for 10 weeks. However, the expressions of PPARγ and PPARα were slightly increased by TM compared with HFD group in WT and MCK-PPARδ−/− mice (Fig. 6A and B). Furthermore, the protein levels of phospho-Akt, phospho-AS160, and plasma membrane Glut4 in skeletal muscle were significantly decreased in HFD-fed mice, which were significantly increased by chronic TM treatment in WT mice but not in MCK-PPARδ−/− mice (Fig. 6C–F). This in vivo evidence indicates that long-term TM treatment enhances Akt and AS160 phosphorylation and Glut4 translocation through PPARδ activation, which could be responsible for the insulin-sensitizing effects of TM in skeletal muscle.
The current study demonstrates that TM can improve insulin sensitivity and increase insulin-stimulated glucose uptake in both cultured myotubes and skeletal muscle from mice. Importantly, this effect was PPARδ-dependent, because the genetic deletion of PPARδ and the presence of PPARδ antagonist abolished the insulin-sensitizing effect of TM. Furthermore, the beneficial effect of TM on glucose homeostasis is associated with increased phosphorylation of Akt and AS160, and Glut4 translocation, which were absent in MCK-PPARδ knockout mice. Thus, in addition to previously reported effects on PPARγ activation, we show for the first time that TM enhances glucose uptake and insulin sensitivity through activation of PPARδ-mediated PI3K signaling in skeletal muscle.
Most hypertensive patients are insulin resistant, and the risk of cardiovascular events is remarkably higher when complicated with diabetes (1,2). Several clinical trials have demonstrated that angiotensin-converting enzyme inhibitors and ARBs may prevent the new onset of diabetes in patients with hypertension (3). Mori et al. (33) and de Luis et al. (34) reported TM improves insulin sensitivity in patients with obesity-related hypertension. Krämer et al. (11) showed that activation of PPARδ results in a direct increase of fatty acid transport and glucose uptake and promotes lipid and glucose metabolism and gene expression in primary cultured human skeletal muscle cells. At 40 mg TM and 160 mg TM, the bioavailability was 42% and 58%, respectively, in humans. Maximal concentrations of TM in human plasma were 159 ± 104 ng/mL for 40 mg and 693 ± 606 ng/mL for 80 mg (35). Plasma concentration was 142.86 ± 14.85 ng/mL in adult C57BL/6J male mice after 1 mg/kg per day administration of TM for 7 days (36). In this study, we treated adult C57BL/6J mice with TM (5 mg/kg per day) for the long-term (37).
The underlying mechanisms of TM that improve insulin sensitivity are not fully elucidated. Previous studies show that TM can increase adiponectin levels, which improves insulin sensitivity (5). Additionally, other beneficial effects of ARBs were reported, including improvement of microcirculation in skeletal muscle (4), reduction of body weight (38), protection of pancreatic islets from glucotoxicity, and oxidative impairment (39). In particular, some ARBs, including TM, can act as a partial agonist of PPARγ (40). Activation of PPARγ consequently might facilitate insulin signaling and improve insulin secretion in β cells (41).
Clemenz et al. (42) recently found that TM also acts as a partial PPARα agonist, at least in the liver, indicating that the various actions of TM might include activation of different PPAR subtypes. Compared with PPARγ, which is rarely expressed in skeletal muscle, PPARδ is highly expressed in skeletal muscle (10). PPARδ activation in adipose tissue leads to enhanced fatty acid oxidation, improved lipid profile, and increased CB1 expression (25), as well as improved glucose metabolism (43). PPARδ activation in cultured human skeletal muscle cells promotes gene-regulatory responses (11). Muscle-specific PPARδ overexpression results in a profound remodeling of myofibers attributable to hyperplasia and a shift to more oxidative fibers, and it increases both the activity and the expression of enzymes involved in oxidative metabolism (12,16).
It was unknown whether TM had an effect on PPARδ. Our study shows that TM acts through PPARδ in skeletal muscle. First, we demonstrated that TM activates PPARδ transcriptional activity and upregulates PPARδ expression in C2C12 myotubes. Second, long-term administration of TM markedly increases PPARδ expression in the skeletal muscle from insulin-resistant mice. Third, no TM stimulatory effect is observed in skeletal muscle from MCK-PPARδ knockout mice or myotubes in the presence of a PPARδ antagonist. These observations support the notion that TM activates PPARδ in skeletal muscle. Skeletal muscle is one of the major insulin target tissues responsible for the maintenance of whole-body glucose homeostasis and accounts for the bulk of insulin-stimulated glucose utilization (30). Glucose transport in skeletal muscle is mainly regulated by glucose transporter Glut4 (30). Insulin stimulates glucose uptake by recruiting Glut4 from an intracellular pool to the cell surface through a mechanism that is dependent on PI3K (30). Insulin resistance leads to defective PI3K/Akt signaling, reduced Glut4 expression, and impaired insulin-stimulated glucose uptake (30). The activation of the PI3K/Akt pathway by PPARδ agonists has been reported previously in several cell types, such as hepatocytes, epithelial cells, and lung cancer cell lines (44–46). In addition, PPARδ agonists also are reported to increase glucose uptake in an insulin-independent manner and lead to the phosphorylation of AMP-activated protein kinase and p38 mitogen-activated protein kinase in cultured primary human skeletal muscle cells (47).
Against this background, we examined the effect of TM on glucose transport and its underlying mechanism in the skeletal muscle. First, using a common in vitro cell model for insulin resistance, we first showed that TM enhanced insulin-stimulated glucose uptake in cultured myotubes. Second, TM promoted the insulin-stimulated translocation of Glut4 in cultured C2C12 myotubes. Third, TM increased phospho-Akt, phospho-AS160, and plasma membrane Glut4 expression in the presence of insulin; however, this effect was inhibited by a PPARδ antagonist and a PI3K inhibitor, but not by PPARγ and PPARα antagonists in cultured myotubes. Experimental evidences in vivo further showed that WT mice fed a HFD and development of glucose intolerance and insulin resistance. Chronic TM treatment reversed the HFD-induced insulin resistance and glucose intolerance in WT mice but not in MCK-PPARδ−/− mice. Meanwhile, TM treatment promoted phosphorylation of Akt and AS160, and increased the expression of PPARδ and plasma membrane Glut4 in the skeletal muscle of WT mice but not in MCK-PPARδ−/− mice.
These findings strongly imply a previously unrecognized role for TM in promoting glucose uptake and improving insulin sensitivity through activation of the PPARδ/PI3K pathway, which may have additional benefits during diabetes therapy. However, TM is not able to eliminate the insulin resistance induced; it only partly improves the defects induced by palmitate. Although TM treatment can increase insulin sensitivity through activation of PPARδ/PI3K pathway in insulin-resistant myotubes induced by palmitate and in mice fed a HFD, the pathophysiology of human diabetes is somewhat different from that obtained in rodents. Further studies are worthy to clarify this point. From a clinical perspective, our findings highlight an important role for TM in the improvement of insulin resistance in the skeletal muscle and further implicate PPARδ as a potential therapeutic target in the treatment of hypertensive subjects with type 2 diabetes.
In summary, we demonstrate that TM has a profound role in the improvement of glucose homeostasis in skeletal muscle, which is associated with activation of the PPARδ/PI3K pathway. It is tempting to speculate that our data provide the molecular basis for the use of ARBs in the treatment and prevention of diabetes in hypertensive patients. A randomized clinical trial is needed to verify whether ARBs could provide additional improvements of skeletal muscle function to hypertensive patients.
This study was funded by the Natural Science Foundation of China (grant no. 30890042) and by National Basic Research Program (973 Program) grants from China (grant no. 2012CB517806 and 2011CB503902).
No potential conflicts of interest relevant to this article were reported.
L.L. and Z.L. performed most of the experiments, analyzed data, and wrote the manuscript. H.Y. performed some experiments and contributed to the discussion. X.F. reviewed and edited the manuscript. P.W., J.C., Y.P., Y.Z., H.H., and J.Z. performed some experiments. D.L. contributed to the discussion and edited the manuscript. Z.Z. designed the experiments and wrote and edited the manuscript. Z.Z. is the guarantor of this work and, as such, had full access to all of the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.
The authors thank Tingbing Cao and Lijuan Wang (Chongqing Institute of Hypertension, China) for technical assistance. The authors thank Bin Tan (Institute of Pediatrics, Children’s Hospital of Chongqing Medical University) for immunofluorescence images from total internal reflection microscopy. The authors thank Prof. Yu Huang and Dr. Wing Tak Wong (Chinese University of Hong Kong, China) for critical review of the manuscript.