Effects of Pioglitazone on Glucose-Dependent Insulinotropic Polypeptide–Mediated Insulin Secretion and Adipocyte Receptor Expression in Patients With Type 2 Diabetes Free

Obesity and type 2 diabetes mellitus (T2DM) are growing global public health crises. Worldwide, >650 million people are obese, and >422 million are affected by T2DM (1). The pathophysiology of T2DM is complex, with multiple metabolic abnormalities, including defects in incretin secretion and action, contributing to hyperglycemia in those with established disease. The incretin hormones glucagon-like peptide 1 (GLP-1) and glucose-dependent insulinotropic polypeptide (GIP) are secreted by enteroendocrine cells following nutrient ingestion and are responsible for stimulating >50% of postprandial insulin secretion (2). Both GLP-1 and GIP are rapidly degraded by dipeptidyl peptidase 4 (DPP-4) following secretion into the circulation. The secretion and clearance of GLP-1 and GIP are largely normal in those with T2DM; however, the insulinotropic effect to these hormones is markedly diminished (3,4). Blunted insulinotropic responses to GLP-1 can be overcome by infusions that achieve supraphysiologic levels; however, even supraphysiologic GIP levels fail to stimulate insulin secretion in most patients with T2DM (5–7). As a consequence, the development of incretin-based therapies for the treatment of T2DM has largely focused on the creation of GLP-1 analogs and inhibitors of DPP-4 action (8,9).

The incretin hormones act through specific G-protein–coupled membrane receptors that are widely expressed throughout the body (10). The GIP receptor (GIP-R) is a Gα_S-protein–coupled receptor found in many tissues, including pancreatic islets, adipose tissue, stomach, central nervous system, adrenal cortex, bone, heart, and endothelium (11,12). In the presence of glucose, activation of the β-cell GIP-R induces insulin secretion, increases proinsulin synthesis, and promotes proliferative and survival pathways (12–14). One theory supposes impairment of the insulinotropic effect of GIP in patients with T2DM to be due to decreased GIP-R levels in the pancreatic β-cells (15,16). The proposed reduction of GIP-R expression might occur through dysregulation of associated transcription factors and may be reversible with improved glycemic control (17–19). However, simple downregulation of the receptor would not explain preservation of early incretin responses with defective late-phase responses in obese patients with T2DM (3).

In addition to β-cells, adipocytes are also an important target of GIP signaling. Fat ingestion is a strong stimulator of GIP secretion (20). Activation of GIP-R in human adipocytes and cell lines in the presence of insulin increases lipoprotein lipase expression, promoting triglyceride storage (21,22). Global GIP-R knockout mice show resistance to weight gain and elevated rates of fat oxidation when fed high-fat diets, effects that can be reversed with rescue of adipose tissue–specific GIP-R expression (23–25). Similarly, adipose-specific GIP-R knockout mice gained less weight on high-fat diets but without changes in fat oxidation (26). GIP-R expression in human adipose tissue decreases with insulin resistance and with increasing BMI (27–29). In patients with T2DM, the attenuated insulinotropic effect of GIP appears to be balanced by increased lipogenic activity, suggesting tissue-specific alteration of GIP action (30). Dysregulation of GIP-R signaling in adipocytes may contribute to insulin resistance, altered lipid metabolism, and adipose tissue dysfunction in obesity and T2DM.

The GIP-R promoter regions contain functional peroxisome proliferator–activated receptor γ (PPARγ) response elements (PPREs), which have been shown to increase GIP-R expression following thiazolidinedione (TZD) treatment in rodent islets and adipocyte cell lines (18,31). Patients treated with TZDs manifest increased insulin sensitivity (S_i) and improved glucose control, but also demonstrate weight gain. The mechanisms underlying these effects are not well understood. We hypothesized that treatment with the TZD pioglitazone (PIO) would improve GIP resistance and increase the insulinotropic effect of GIP on β-cells. We also hypothesized that TZD treatment would increase GIP-R expression in human adipocytes and that this might be an important pathway mediating improved glycemic control, lipid metabolism, and weight gain. To test these hypotheses, we conducted a randomized, double-blind, placebo (PBO)–controlled trial of PIO in 24 subjects with well-controlled T2DM. We assessed glycemic control, insulin secretory responses, and lipid regulation using oral glucose tolerance tests (OGTTs), intravenous glucose tolerance tests (IVGTTs), a GIP infusion, mixed-meal tolerance (MMT) testing, body composition, and adipose tissue gene expression before and after 12 weeks of treatment.

Twenty-four subjects with well-controlled T2DM (HbA_1c <7.0% [55 mmol/mol]) treated with diet and exercise (n = 9) or metformin (500–1,000 mg/day; n = 15) were enrolled in a 12-week, randomized, double-blinded, PIO- (45 mg/day; n = 12) or PBO-controlled (n = 12) trial (NCT00656864). Subject characteristics are presented in Table 1. Subjects were excluded from the study if they had acute or chronic health conditions, such as heart or kidney failure or cirrhosis, or if they were currently taking or had taken a TZD, sulfonylurea, DPP-4 inhibitor, GLP-1 analog, insulin, or weight-loss medication within the past 6 months. This study was approved by the University of Vermont Institutional Review Board. All subjects provided written informed consent prior to participating in any study procedures.

Subjects had a screening visit to verify inclusion criteria and provide informed consent. Prior to randomization, all subjects had a baseline OGTT, MMT test, frequently sampled IVGTT, graded glucose infusion and assessment of GIP-stimulated insulin secretion (GIP-SIS), measurement of body composition by whole-body DXA, and subcutaneous abdominal adipose tissue biopsies. All procedures were performed after an overnight fast and conducted with 1 week between each metabolic test. Following baseline testing, subjects were randomized to receive 45 mg/day PIO or PBO. After 12 weeks of treatment, subjects underwent a second round of metabolic testing and adipose tissue biopsies. All procedures and visits occurred at the University of Vermont Diabetes Research Center or University of Vermont General Clinical Research Center. HbA_1c and plasma lipids were measured at the Fletcher Allen Hospital clinical pathology laboratory according to standard methods.

For the OGTT, 75 g of glucose (300 kcal) was delivered as a flavored beverage. The MMT test consisted of a 300-kcal beverage containing 50% carbohydrate, 15% protein, and 35% fat (Boost Plus; Nestlé, Vevey, Switzerland). Blood samples were collected at baseline and every 30 min for 3 h for both tests. The IVGTT consisted of an intravenous bolus of 0.3 g dextrose/kg body weight followed by an insulin bolus (0.06 IU/kg body weight) 20 min later (32). All blood samples were collected on ice and aliquoted for measurement of glucose (K⁺ Oxalate tubes), insulin, and C-peptide (serum-separating tubes) and total incretin hormones (K⁺ EDTA with aprotinin; Bayer Pharmaceuticals, West Haven, CT). Glucose was measured by the glucose oxidase method (YSI Inc., Yellow Springs, OH), insulin by immunoassay (ALPCO, Salem, NH), C-peptide by immunoassay (Meso Scale Diagnostics, Rockville, MD), and nonesterified free fatty acids (FFAs) by an enzymatic colorimetric method (Wako Diagnostics, Mountain View, CA). Incretin hormones were measured using specific C-terminally directed assays detecting total GLP-1 and GIP (33). S_i was modeled from the OGTT (oral glucose S_i [OGIS]) according to the method of Mari et al. (34). S_i and the acute insulin response to glucose (AIR_g) were modeled from the IVGTT using the Bergman Minimal Model (35). Insulin secretion rates (ISRs) were calculated from C-peptide levels by population deconvolution (36).

Following each IVGTT, subjects underwent graded glucose infusion followed by a GIP infusion. After the 3-h IVGTT, 20% dextrose was administered at 300 mL/h for 30 min and at 450 mL/h for 30 min, and then the dextrose infusion was continued while GIP (2 pmol/kg/min) was coinfused for another 30 min. Glucose-stimulated insulin secretion (GSIS) was calculated as a linear insulin-to-glucose ratio during the first portion of the graded glucose infusion. GIP-SIS was calculated as the incremental difference between the insulin concentration for a given glucose concentration along the GSIS line and the plasma insulin level during the GIP infusion. GSIS rates (GS-ISRs) and GIP-SIS rates (GIP-SISRs) were calculated by the same method using ISRs derived from C-peptide data.

Subcutaneous periumbilical abdominal adipose tissue was obtained by percutaneous needle biopsy under local anesthesia. Tissue was aspirated into sterile PBS and washed twice to remove any blood. Whole tissue samples were then frozen in liquid nitrogen and stored at −80°C. Adipose tissue from needle biopsies was also incubated in saline with collagenase (Worthington Biochemical Corp, Lakewood, NJ) at 37°C for 80 min, passed through a 200-μm pore filter, and centrifuged to pellet stromal vascular cells. Following centrifugation, tubes were left for 5 min, allowing adipocytes to separate by floatation. Isolated adipocytes and stromal vascular cells were then snap frozen in liquid nitrogen and stored at −80°C.

Preadipocytes were cultured and differentiated into adipocytes using previously established protocols with slight modifications (37). Preadipocytes were expanded from stromal vascular fractions in growth media (minimum essential medium α with nucleosides; Gibco; Thermo Fisher Scientific, Waltham, MA), 2.5% FBS (Gibco), 10 ng/mL human epidermal growth factor (Gibco), 1 ng/mL human fibroblast growth factor-2 (Gibco), 5.5 mmol/L glucose, and 10 nmol/L human insulin (Humulin R; Eli Lilly and Company, Indianapolis, IN) at 37°C with 5% CO₂ for no more than two passages. Preadipocytes were subcultured at 4.5 × 10⁴/well in 6-well plates for 10 days. Media were changed after the first 24 h and then every 72 h. Preadipocyte plates were triggered to differentiate using minimum essential medium α/F12 with 17.5 mmol/L glucose (Gibco), 15 mmol/L HEPES (Gibco), 2 nmol/L triiodothyronine (Sigma-Aldrich, St. Louis, MO), 10 mg/L transferrin (Sigma-Aldrich), 17 μmol/L pantothenate (Sigma-Aldrich), 33 μmol/L biotin (Sigma-Aldrich), 100 nmol/L dexamethasone (Sigma-Aldrich), 1 μmol/L troglitazone (Sigma-Aldrich), 0.5 mmol/L 3-isobutyl-L-methylxanthine (Sigma-Aldrich), and 100 nmol/L human insulin. Cells were incubated in the trigger medium for 4 days and then changed to maintenance media (DMEM/F12 with 5.5 mmol/L glucose, 10 nmol/L insulin, 15 mmol/L HEPES, 1 nmol/L triiodothyronine, 10 mg/L transferrin, 17 μmol/L pantothenate, 33 μmol/L biotin, and 10 nmol/L dexamethasone). Adipocytes were allowed to mature for 12 days, and media were changed every 72 h. For the final 72 h, paired cultures were treated with 10 μmol/L troglitazone or vehicle to activate PPARγ for chromatin immunoprecipitation (ChIP) assays.

Adipose tissue and adipocyte RNA were extracted using RNeasy Lipid Tissue Mini Kits (Qiagen, Valencia, CA) with RNase-Free DNase treatment (Qiagen). A cDNA library was made using Advantage RT-for-PCR kits (Clontech Laboratories, Inc., Mountain View, CA) using 1 μg RNA template and oligo(dT) primers. Gene expression was measured by real-time PCR on an Applied Biosystems 7300 (Life Technologies Inc., Carlsbad, CA) using TaqMan primers for PPARγ (Hs01115510_m1) and GIP-R (Hs00609210_m1). All expression data were normalized to GAPDH (4326317E). Relative expression levels were calculated using the −ΔΔCt threshold cycle (Ct) method.

ChIP assay was performed on cultured human adipocytes treated with 10 μmol/L troglitazone or vehicle (DMSO) for 72 h using the ChIP-IT kit (Active Motif, Carlsbad, CA). Cells were fixed using formaldehyde solution and chromatin sheared by enzymatic digestion into DNA fragments averaging 300–500 bp. Rabbit polyclonal antibody against PPARγ (PA3–821A; Thermo Fisher Scientific) was incubated overnight with precleared chromatin, and parallel samples were incubated with negative control IgG provided with the kit. Protein G–agarose beads were then added and incubated for 1.5 h at 4°C. After reversing the cross-links, DNA was isolated, and real-time PCR (ABI 7300) was performed with target-validated SYBR Green primer pairs (EpiTect Human GIP-R ChIP assay, catalog no. GPH100697[-] 04 kb; Qiagen) for the flanking regions of human GIP-R PPRE promoter (GenBank: NM_000164.2, Chr19:46167667–46167684). PCR conditions were 1 cycle at 95°C for 10 min and 40 cycles at 95°C for 15 s and at 60°C for 1 min. The ChiP quantitative PCR DNA fractions for PPARγ and nonimmune serum (NIS) were normalized with input DNA to get ΔCt value, and ΔΔCt was obtained by normalizing NIS background. The fold enrichment of PPARγ on GIP-R promoter was calculated by 2^{(−ΔΔCt[PPAR/NIS])}. The ChIP was performed on three independent human adipocyte cultures for vehicle (DMSO) or troglitazone-treated samples.

Study data were collected using REDCap electronic data capture tools hosted at the University of Vermont (REDCap Consortium, Vanderbilt University, Nashville, TN). All data were loaded into STATA v11.2 (StataCorp LP, College Station, TX). Demographic, metabolic, and ChIP data were normally distributed. Gene expression data underwent ln transformation prior to analysis. Categorical data were analyzed by Fisher exact test. Continuous data were analyzed by t test, paired where appropriate. Prespecified primary outcomes were change in GIP-SIS and GIP-R expression. A priori power calculation indicated 80% power to detect 120% change in either parameter with 95% confidence. The study was not powered for multiple covariate analysis, but exploratory analysis with simple linear regression modeling was conducted using prestudy values as a covariate. All data are presented as mean ± SD. A P value <0.05 was considered significant.

At baseline, the age, weight, body composition, glycemic control, use of metformin and statins, and all measured parameters were similar between subjects that were randomized to the PIO and PBO group (Table 1). After 12 weeks of treatment, subjects on PIO gained more weight than those on PBO (P < 0.001), increased their BMI (P < 0.001), and increased DXA fat mass (P = 0.008) (Supplementary Table 1). Overall glycemic control, measured by HbA_1c, was significantly better in subjects on PIO compared with PBO (P = 0.04). Treatment with PIO reduced fasting glucose levels (P = 0.008), and fasting insulin concentrations trended lower compared with PBO (P = 0.09). The HOMA of insulin resistance was reduced in subjects treated with PIO (P = 0.03).

S_i derived from IVGTT was increased by treatment with PIO (P = 0.03), while first-phase insulin secretion (AIR_g) was unchanged compared with PBO (P = 0.7) (Fig. 1). Areas under the curve (AUCs) for glucose and insulin during the OGTT were significantly lower with PIO treatment compared with PBO (P = 0.002 and 0.003, respectively) (Fig. 2). S_i derived from the OGTT (OGIS) was significantly improved by treatment with PIO (P = 0.002). GLP-1 and GIP release following OGTT was not changed by treatment with PIO (P = 0.15 and P = 0.97, respectively) (Supplementary Fig. 1). Subjects treated with PBO did have a higher GLP-1 AUC at the end of the study compared with baseline (P = 0.008), but the poststudy levels did not differ between PBO and PIO treatment (P = 0.7). Subjects treated with PIO had decreased glucose AUC during the MMT test (P = 0.006) compared with PBO (Fig. 3). Insulin AUC trended lower with PIO treatment, but did not reach statistical significance (P = 0.09). GLP-1 and GIP levels in response to the MMT were not changed by treatment with PIO (P = 0.75 and P = 0.40, respectively) (Supplementary Fig. 2).

Insulin levels increased linearly during the glucose infusion, with the slope of the line defining GSIS (Fig. 4). Treatment with PIO reduced GSIS, but this change was not statistically significant (P = 0.07). The increase in circulating insulin above GSIS after GIP infusion (GIP-SIS) was reduced following treatment with PIO (P = 0.03) and unchanged in those receiving PBO (P = 0.9). Due to potential changes in hepatic insulin extraction, we also calculated ISRs from C-peptide levels (Fig. 5). The GS-ISR was reduced ∼50% in subjects treated with PIO (P < 0.001); however, the GIP-SISR was not changed by treatment with PIO (P = 0.5).

Circulating triglycerides and cholesterol/HDL ratios were significantly reduced by treatment with PIO (P = 0.04 and 0.03, respectively), while total cholesterol, HDL, and LDL were not (P > 0.3). FFA suppression during IVGTT was increased from 68 to 85% following treatment with PIO (P = 0.007) and increased from 57 to 72% during MMT testing (P = 0.04) (Fig. 6).

Expression of PPARγ in adipocytes was not different between subjects randomized to PIO or PBO at baseline or following the 12 weeks of treatment (Fig. 7A). Subjects treated with PIO had increased adipocyte GIP-R expression (P = 0.015), while those treated with PBO did not (P = 0.15) (Fig. 7B). Poststudy GIP-R expression was associated with increased FFA suppression during IVGTT (P = 0.02) and reductions in cholesterol/HDL ratio (P = 0.002) by simple linear regression, but had no association with alterations in glucose and insulin homeostasis, weight, or changes in body composition after controlling for prestudy gene expression. Treatment with PIO and metformin was associated with an attenuated increase in GIP-R expression compared with PIO alone (P = 0.009 for the interaction).

Because adipose GIP-R gene expression was increased in subjects treated with PIO, we asked if this was a direct effect of PPARγ binding. We treated differentiated human adipocyte cultures with 10 μmol/L troglitazone or vehicle in parallel (n = 3 paired cultures). PPARγ was bound to the GIP-R promoter in cultures treated with vehicle, and treatment with troglitazone increased the binding by more than twofold (P = 0.03) (Fig. 7C). These data show that PPARγ binds to the human GIP-R promoter and that treatment with a TZD increases PPARγ occupancy of the GIP-R PPREs.

Impairment of incretin function is a hallmark of T2DM, but mechanisms involved in the dysregulation of the incretin system are not fully delineated. PPARγ (NR1C3 gene) is a member of the nuclear hormone receptor superfamily of ligand-activated transcription factors. As a transcriptional regulator, PPARγ must be ligand activated, undergo heterodimerization with retinoid X receptors (NR2B), recruit cofactors, and recognize a response element (PPRE) in the promoter region of target genes (38–40). Transcriptional activation of PPARγ induces diverse biological actions, including lipid metabolism, cellular growth, and differentiation. TZDs are orally active agents that are high-affinity activators of PPAR-γ (41). Clinical trials of TZDs demonstrate improved glucose and lipid control, enhanced peripheral S_i, and preservation of β-cell function in prediabetes and early T2DM compared with other therapies (42,43).

Contrary to our hypothesis, we found that treatment with PIO did not increase the insulinotropic effect of GIP in subjects with well-controlled T2DM. The lack of effect likely reflects the relatively good glycemic control of these study subjects. It is possible that β-cell expression of GIP-R was not downregulated and therefore could not be enhanced with PIO. This notion is supported by robust prestudy insulin secretion in response to GIP infusion. These subjects had well-controlled T2DM and, possibly, a preserved insulinotropic incretin response. Conversely, the effects of PIO treatment along other pathways or in other tissues may predominate, resulting in a lack of change in response to incretins. Subjects treated with PIO did have significant improvement of S_i with increased S_i and OGIS in concert with reduced GS-ISRs. In addition to the lack of improvement in GIP-SISR, we did not observe any alteration in first-phase insulin secretion or GSIS by PIO treatment. These results are consistent with an overall improvement in peripheral glucose disposal without an increase in β-cell function, suggesting islet incretin function was not impaired at baseline.

PIO treatment caused increases in fat mass, reduced circulating triglycerides, and the cholesterol/HDL ratio and increased FFA suppression during IVGTT and MMT testing, effects consistent with amplified lipid storage. While we do not have direct evidence of a functional effect of increased adipocyte GIP-R expression, the change in adipose GIP-R could contribute to the well-described weight gain and increases in subcutaneous adipose tissue that occur with TZD treatment (44). Acute GIP infusions in obese subjects with untreated T2DM resulted in more FFA suppression and subcutaneous adipose triglyceride storage during a hyperglycemic clamp compared with PBO (30). This lipogenic effect was not seen in metabolically normal lean or obese subjects or in obese subjects with impaired glucose tolerance, causing the authors to conclude that there is a dissociation of the lipogenic and insulinotropic actions of GIP in T2DM, which may increase obesity and insulin resistance. We found associations between increased adipocyte GIP-R expression and improved lipid metabolism, but whether elevated adipocyte GIP-R contributes to fat accumulation or improved lipid metabolism during PPARγ agonist treatment requires further investigation.

The PPAR family has a wide cistrome driving a range of gene promoters; thus, our observed effects may represent contributions from other pathways promoted by PPAR binding (38). Our results suggest dysregulated adipocyte GIP-R may be a key feature of the altered incretin response in T2DM, but the functional role of adipose GIP-R in T2DM is not fully understood (45). Diminished GIP or discordant action in clinical and experimental models of T2DM may be directly related to tissue-specific downregulation of GIP-R or reduced recycling of the receptor to the cell surface (10,30,45). Obesity may be associated with downregulation of GIP-R in adipose tissue and adipocytes that could contribute to insulin resistance (27–29). We speculate that reduced adipose GIP-R expression could result from attenuated adipose PPARγ signaling and may be one mechanism causing incretin dysfunction in T2DM. A clearer understanding of the functional changes of GIP-R signaling induced by T2DM is especially important, as dual incretin agonists have a more profound effect on weight loss than GLP-1 receptor agonists (46). Emerging therapies using direct GLP-1 and GIP agonists will benefit greatly from a more thorough understanding of extrapancreatic incretin biology. Our results suggest the potential for synergy of TZD and incretin agonist therapy (47).

Duality of Interest. This research was funded by an investigator-initiated grant from Takeda Pharmaceuticals North America. The funding source had no input on data analyses or interpretation. No other potential conflicts of interest relevant to this article were reported.

Author Contributions. W.G.T. assisted with conduct of the clinical study, assisted with ChIP experiment design, performed adipose culture, analyzed the data, and wrote the manuscript. D.G. designed and performed the ChIP experiments, assisted with adipose culture, and edited the manuscript. O.S. processed adipose tissue biopsies and conducted gene expression analyses. C.F.D. and J.J.H. performed and interpreted incretin hormone measurements and edited the manuscript. D.E. provided the GIP hormone, assisted with study analysis and interpretation, and edited the manuscript. R.E.P. designed and conducted the clinical study, oversaw data analyses, and edited the manuscript. W.G.T. and R.E.P. are the guarantors of this work and, as such, had full access to all of the data in the study and take responsibility for the integrity of the data and the accuracy of the data analysis.

Prior Presentation. This study was presented at the 71st Scientific Sessions of the American Diabetes Association, San Diego, CA, 24–28 June 2011.