Hepatocyte nuclear factor-1α (HNF1α) is a transcription factor expressed in tissues of endoderm origin. Mutations in HNF1A are associated with maturity-onset diabetes of the young 3 (MODY3). Mice deficient for Hnf1α are hyperglycemic, with their pancreatic β-cells being defective in glucose-sensing insulin secretion. The specific mechanisms involved in this defect are unclear. Gut hormones control glucose homeostasis. Our objective was to explore whether changes in these hormones play a role in glucose homeostasis in the absence of Hnf1α. An increase in ghrelin gene transcript and a decrease in glucose-dependent insulinotropic polypeptide (GIP) gene transcripts were observed in the gut of Hnf1α-null mice. These changes correlated with an increase of ghrelin and a decrease of GIP-labeled cells. Ghrelin serological levels were significantly induced in Hnf1α-null mice. Paradoxically, GIP levels were also induced in these mice. Treatment of Hnf1α-null mice with a ghrelin antagonist led to a recovery of the diabetic symptoms. We conclude that upregulation of ghrelin in the absence of Hnf1α impairs insulin secretion and can be reversed by pharmacological inhibition of ghrelin/GHS-R interaction. These observations open up on future strategies to counteract ghrelin action in a program that could become beneficial in controlling non–insulin-dependent diabetes.

Maturity-onset diabetes of the young (MODY) is a monogenic autosomal dominant form of diabetes that first occurs during early adulthood and is characterized by pancreatic β-cell dysfunction (1). Subtypes of MODY have been classified based on the specific nature of the mutated genes of which six have been identified (2). MODY3, the most common MODY mutation in the population, encodes the transcription factor hepatocyte nuclear factor-1α (HNF1α) (3,4) involved in regulation of a large subset of genes in the liver, pancreas, kidney, and intestine. Although some pancreatic HNF1α targets are suggested to impact the disease phenotypes, the exact nature of the molecular links between loss of HNF1α function and manifestation of the disease is still unclear.

Mouse models with deletion of Hnf1α functions display hepatic and renal dysfunction coupled to non–insulin-dependent diabetes and dwarfism (5,6). Although these mice still produce insulin, their pancreatic β-cells are defective in glucose-sensing insulin secretion (7,8). Simultaneous Hnf1α re-expression in both liver and endocrine pancreas of Hnf1α-null mice failed to restore normal blood glucose and insulin levels, suggesting that other tissues in which Hnf1α was deleted could be participating in the diabetic phenotype of these mice (9).

Gut hormones are produced by enteroendocrine cells and are crucial regulators of glucose homeostasis and pancreatic insulin secretion (10). Glucose-dependent insulinotropic polypeptide (GIP) and glucagon-like peptide 1 (GLP-1) are incretins that stimulate insulin secretion, whereas ghrelin targets pancreatic β-cells to limit insulin production (10). Hnf1α-null mice display intestinal epithelium dysfunctions, including altered enteroendocrine cell differentiation (11). Here, we aimed to explore if specific changes in gastrointestinal hormones could functionally relate to glucose homeostasis in Hnf1α-null mice.

Animals and Analytical Procedures

Hnf1α-null mice (5,11) and control littermates were treated in accordance with the Institutional Animal Research Review Committee of the Université de Sherbrooke (approval ID number 102-14B). Hnf1α-null mice were genotyped as previously described (11). Blood glucose values were determined from whole venous blood from mice fed ad libitum or 16 h fasted using a glucose monitor (FreeStyle Lite; Abbott Diabetes Care). [d-Lys-3]-GHRP-6 (Bachem), a classical but not highly selective GHS-R antagonist (1214), was freshly diluted in 100 µL of saline and intraperitoneal injections were performed every 12 h during five consecutive days followed by a 16-h fasting period before sacrifice or intraperitoneal glucose tolerance tests (IPGTTs) (2 g d-glucose/kg). The optimal dose of GHS-R antagonist (200 nmol/30 g) was determined according to previous published work (15,16). For metabolic analyses, mice were individually placed in metabolic cages, provided with the same quantities of food and water, and housed on a reverse light-dark cycle. All groups were fed ad libitum throughout the duration of the study. After a 5-day adaptation period after being transferred from group housed cages to single housed metabolic cages, mice were treated with GHS-R antagonist or saline during 5 days. Body weight (g), food intake (g), water intake (mL), urine (mL), and feces (g) were measured every morning before injections. Urine glucose content was determined with Chemstrip 10 urine test strips (Roche Diagnostics).

RNA Isolation and RT-PCR

Total RNA from the jejunum and stomach was isolated, and qRT-PCR was performed as previously described (11). Results were calibrated with TATA box binding protein (TBP). Primer sequences are available upon request.

Immunofluorescence

Jejunum segments and pancreas were fixed in 4% paraformaldehyde overnight at 4°C, dehydrated, embedded in paraffin, and cut to 5-µm sections. Immunofluorescences were performed as previously described (17). The following affinity-purified antibodies (Santa Cruz Biotechnology) were used: goat anti-ghrelin (sc-10368; diluted 1/100), goat anti-GIP (sc-23554; diluted 1/50), and mouse anti-insulin (sc-8033; diluted 1/200).

ELISA

Blood was collected from the right heart ventricle of 16 h–fasted mice and pretreated with Pefabloc solution (ghrelin) or dipeptidyl peptidase 4 (DPP4) inhibitor (GIP and GLP-1). After 30 min at room temperature, samples were centrifuged at 3,000g for 15 min at 4°C. Acidification of the serum samples with HCl to a final concentration of 0.05 N was performed. Total ghrelin (EZRGRT-91K), active ghrelin (EZRGRA-90K), total GIP (EZRMGIP-55K), active GLP-1 (EGLP-35K), and insulin (EZRMI-13K) were measured using ELISA kits from EMD Millipore. Total glucagon was assessed with the ELISA kit DGCG0 (R&D Systems). The Ultra Sensitive Mouse Insulin ELISA Kit 90080 (Crystal Chem) was used during IPGTT procedures. Total DPP4 was measured with the DPP4 ELISA Kit SEA884Mu (USCN Life Science).

Statistical Analysis

Statistical analyses were performed using the GraphPad Prism 6 software. Statistics were calculated using the two-way, two-tailed Student t test or two-way nested ANOVA. Differences were considered significant with a P value of <0.05.

Assessment of circulating levels of glucose (Fig. 1A), insulin (Fig. 1B), and glucagon (Fig. 1C) in Hnf1α mutant and control mice confirmed the hyperglycemic state of the mutants with reduced circulating insulin levels without alteration of glucagon levels. Immunofluorescence detection of insulin in the pancreas of Hnf1α mutant (Fig. 1D) and control mice (Fig. 1E) suggested a comparable potential of pancreatic β-cells in expressing insulin peptide. Since the intestinal endocrine system plays a crucial role in regulating glucose metabolism, expression of relevant hormones in the intestine of Hnf1α mutant mice was monitored. Analysis of gene transcript expression for ghrelin, GIP, and GLP-1 was determined by RT-qPCR in the jejunum of newborn and adult Hnf1α mutant and control mice. Whereas ghrelin transcripts were significantly increased in the jejunum of mutant as compared with control mice (1.79-fold increase at day 1, P < 0.01; 4.30-fold increase at 4 months, P < 0.01) (Fig. 1F), GIP transcripts were significantly decreased (4.41-fold decrease at day 1, P < 0.05; 3.48-fold decrease at 4 months, P < 0.01) (Fig. 1G). GLP-1 transcripts were not affected under these conditions (Fig. 1H). Immunofluorescences were performed to monitor the distribution of corresponding enteroendocrine cells. The number of ghrelin-positive cells was significantly increased in the jejunum of adult Hnf1α mutant when compared with control mice (3.72-fold increase, P < 0.0001) (Fig. 1I and J), whereas the number of GIP-positive cells decreased (2.23-fold decrease, P < 0.0001) (Fig. 1K and L).

Figure 1

Loss of Hnf1α affects expression of gastrointestinal hormones. Blood glucose (A), insulin (B), and glucagon (C) levels were determined from 16 h–fasted adult control and Hnf1α-null mice (n = 5–9). Representative immunofluorescence for insulin that was performed on sections of pancreas of both Hnf1α mutant (D) and control (E) mice. qRT-PCR detection of ghrelin (F), GIP (G), and GLP-1 (H) mRNA was performed on total small intestinal RNA extracts from newborn and adult control and Hnf1α-null mice and calibrated in comparison with TBP mRNA detection (n = 4–7). The proximal small intestine of both control and Hnf1α-null mice was labeled for ghrelin (I) or GIP (K) by immunofluorescence. Total numbers of positively stained cells for ghrelin (J) and GIP (L) were calculated on an average of 40 crypt-villus axes per animal (n = 6). Data were analyzed with the unpaired Student t test and error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Figure 1

Loss of Hnf1α affects expression of gastrointestinal hormones. Blood glucose (A), insulin (B), and glucagon (C) levels were determined from 16 h–fasted adult control and Hnf1α-null mice (n = 5–9). Representative immunofluorescence for insulin that was performed on sections of pancreas of both Hnf1α mutant (D) and control (E) mice. qRT-PCR detection of ghrelin (F), GIP (G), and GLP-1 (H) mRNA was performed on total small intestinal RNA extracts from newborn and adult control and Hnf1α-null mice and calibrated in comparison with TBP mRNA detection (n = 4–7). The proximal small intestine of both control and Hnf1α-null mice was labeled for ghrelin (I) or GIP (K) by immunofluorescence. Total numbers of positively stained cells for ghrelin (J) and GIP (L) were calculated on an average of 40 crypt-villus axes per animal (n = 6). Data were analyzed with the unpaired Student t test and error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

ELISA was next performed to measure circulating levels of gastrointestinal hormones in Hnf1α mutant and control mice. Total circulating ghrelin was significantly increased in Hnf1α mutant as compared with control mice (6.57-fold increase at 1 month, P < 0.001; 4.16-fold increase at 4 months, P < 0.01) (Fig. 2A). These increases were also reflected at the level of active ghrelin form (4.92-fold increase at 1 month, P < 0.01; 4.72-fold increase at 4 months, P < 0.05) (Fig. 2B). As opposed to gene transcript level, total circulating GIP was significantly upregulated in Hnf1α mutant as compared with control mice (14.51-fold increase at 1 month, P < 0.05; 4.03-fold increase at 4 months, P < 0.01) (Fig. 2C). Basal active GLP-1 circulating levels were undetectable in both Hnf1α mutant and control mice under these conditions. Since GIP peptide stability is dependent on DPP4 activity (18) and Hnf1α activates transcription of DPP4 (19), DPP4 circulating levels were measured. A reduction of circulating DPP4 was observed in Hnf1α mutant as compared with control mice (5.07-fold decrease, P < 0.01) (Fig. 2D). Since ghrelin is mostly secreted from the stomach and the jejunum, the relative ratio of active ghrelin in each of these tissues was monitored in Hnf1α mutant and control mice. ELISA revealed a significant 2.55-fold increase (P < 0.05) in active ghrelin per gram of jejunum of Hnf1α mutant when compared with control mice, whereas no significant change was observed in the stomach of these animals (Fig. 2E). Coincidently, ghrelin transcripts were not significantly modulated in the stomach of Hnf1α mutant as compared with control mice (P = 0.49, n = 4).

Figure 2

Loss of Hnf1α impacts ghrelin and GIP circulating levels. Total ghrelin (A), active ghrelin (B), GIP (C), and DPP4 (D) circulating levels were assessed from 16 h–fasted adult control and Hnf1α-null mice by ELISAs (n = 3–6). E: Total protein extracts were isolated from whole stomach or jejunum of 16 h–fasted adult control and Hnf1α-null mice, and active ghrelin was assessed by ELISA (n = 5–6). Data were analyzed with the unpaired Student t test and error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001.

Figure 2

Loss of Hnf1α impacts ghrelin and GIP circulating levels. Total ghrelin (A), active ghrelin (B), GIP (C), and DPP4 (D) circulating levels were assessed from 16 h–fasted adult control and Hnf1α-null mice by ELISAs (n = 3–6). E: Total protein extracts were isolated from whole stomach or jejunum of 16 h–fasted adult control and Hnf1α-null mice, and active ghrelin was assessed by ELISA (n = 5–6). Data were analyzed with the unpaired Student t test and error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001.

Ghrelin can limit insulin release by interacting with the GHS-R1a receptor on β-pancreatic cells (20). To test whether increases in active ghrelin were functionally related to the hypoinsulinemia state of Hnf1α mutant mice, the GHS-R antagonist [d-Lys-3]-GHRP-6 was administrated intraperitoneally to mice. Single injections of the GHS-R antagonist every 12 h progressively led to a decrease in blood glucose level in Hnf1α mutant mice to reach statistically undistinguishable levels from the controls after 5 days of treatment (Fig. 3A). This effect was reversible with a progressive return to hyperglycemia steady state 1 week after stopping injections (Fig. 3B). Hypoinsulinemia of Hnf1α mutant mice was corrected after 5 days of ghrelin antagonist treatment (Fig. 3C). IPGTTs were further performed to monitor glucose clearance of treated and nontreated mice. Glucose levels in fasted nontreated Hnf1α mutant mice rose above 23 mmol/L after 15 min and failed to significantly decline at 120 min (Fig. 3D). Glucose levels in fasted Hnf1α mutant mice pretreated with GHS-R antagonist rose above nontreated and treated control mice at 15 min but rapidly declined to reach comparable values with the control groups at 120 min (Fig. 3D). Area under the curve (AUC) calculations revealed a significant recovery for Hnf1α mutant mice pretreated with the GHS-R antagonist in blood glucose clearance (Fig. 3E). Circulating insulin levels during IPGTT were significantly increased between Hnf1α mutant mice pretreated with GHS-R antagonist versus nontreated Hnf1α mutant mice (Fig. 3F), whereas GIP levels were significantly decreased with GHS-R antagonist pretreatment (Fig. 3G).

Figure 3

Impact of Hnf1α mutant mice treatment with the GHS-R antagonist on glucose homeostasis. A: Adult control and Hnf1α-null mice were intraperitoneally injected with saline (top panel) or GHS-R antagonist (bottom panel) for 5 days. Blood glucose levels were assessed every morning of each day (n = 6 for each group). B: Hnf1α-null mice were intraperitoneally injected with GHS-R antagonist for 5 days and left to recover. Blood glucose levels were assessed at each indicated day (n = 10). C: Adult control and Hnf1α-null mice were intraperitoneally injected with saline or GHS-R antagonist for 5 days. Mice were fasted for 16 h and blood insulin levels assessed by ELISA (n = 10 for each group). D: Adult control and Hnf1α-null mice were intraperitoneally injected with saline or GHS-R antagonist for 5 days. Mice were fasted for 16 h and IPGTT was performed. Blood glucose levels were measured at each indicated time (n = 5 for each group). Glucose AUC was calculated over the 120-min period (E) and insulin levels (F) and GIP levels (G) measured by ELISAs at 30 and 120 min. Data were analyzed with the unpaired Student t test except for AUC where ANOVA was performed. Error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Figure 3

Impact of Hnf1α mutant mice treatment with the GHS-R antagonist on glucose homeostasis. A: Adult control and Hnf1α-null mice were intraperitoneally injected with saline (top panel) or GHS-R antagonist (bottom panel) for 5 days. Blood glucose levels were assessed every morning of each day (n = 6 for each group). B: Hnf1α-null mice were intraperitoneally injected with GHS-R antagonist for 5 days and left to recover. Blood glucose levels were assessed at each indicated day (n = 10). C: Adult control and Hnf1α-null mice were intraperitoneally injected with saline or GHS-R antagonist for 5 days. Mice were fasted for 16 h and blood insulin levels assessed by ELISA (n = 10 for each group). D: Adult control and Hnf1α-null mice were intraperitoneally injected with saline or GHS-R antagonist for 5 days. Mice were fasted for 16 h and IPGTT was performed. Blood glucose levels were measured at each indicated time (n = 5 for each group). Glucose AUC was calculated over the 120-min period (E) and insulin levels (F) and GIP levels (G) measured by ELISAs at 30 and 120 min. Data were analyzed with the unpaired Student t test except for AUC where ANOVA was performed. Error bars represent SE. *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Since ghrelin can impact appetite and metabolism, solid and liquid metabolism was investigated among the various mouse groups using metabolic cages. Analysis of solid metabolism indicated that food intake ratios were increased in Hnf1α mutant compared with control mice (145%), and GHS-R antagonist treatment did not significantly influence this tendency (Fig. 4A). This observation was consistent with fecal ratios that were increased in Hnf1α mutant mice (178%) (Fig. 4B). Analysis of liquid metabolism revealed that water intake ratios were increased by 182% in Hnf1α mutant compared with control mice (Fig. 4C), and GHS-R antagonist treatment significantly reduced this ratio in Hnf1α mutant mice (Fig. 4C). Urine ratios were not significantly affected when Hnf1α mutant mice were compared with controls (Fig. 4D). However, GHS-R antagonist treatment significantly reduced this ratio in both Hnf1α mutant and control mice (Fig. 4D). Detection of glucose in the urine of these Hnf1α mutant mice revealed an important eradication of glucose content after GHS-R antagonist treatment (Fig. 4E), whereas control mice remained negative under these treatments (not shown).

Figure 4

Impact of Hnf1α mutant mice treatment with the GHS-R antagonist on liquid and solid metabolisms. Hnf1α mutant (n = 6) and control mice (n = 6) metabolism was evaluated at the beginning (day 0) and the end (day 5) of saline or GHS-R antagonist intraperitoneal treatment. Solid metabolism was measured by calculating food ratios (grams of chow per gram weight) (A) and fecal excretion (grams of feces per gram weight) (B). Liquid metabolism was measured by calculating water ratios (milliliters of water per gram weight) (C) and urine ratios (milliliters of urine per gram weight) (D). E: Urine glucose content was determined in Hnf1α mutant mice at the beginning (day 0) and the end (day 5) of GHS-R antagonist intraperitoneal treatment. Data were analyzed with the unpaired Student t test, and error bars represent SE. ns, not significant. *P < 0.05; **P < 0.01.

Figure 4

Impact of Hnf1α mutant mice treatment with the GHS-R antagonist on liquid and solid metabolisms. Hnf1α mutant (n = 6) and control mice (n = 6) metabolism was evaluated at the beginning (day 0) and the end (day 5) of saline or GHS-R antagonist intraperitoneal treatment. Solid metabolism was measured by calculating food ratios (grams of chow per gram weight) (A) and fecal excretion (grams of feces per gram weight) (B). Liquid metabolism was measured by calculating water ratios (milliliters of water per gram weight) (C) and urine ratios (milliliters of urine per gram weight) (D). E: Urine glucose content was determined in Hnf1α mutant mice at the beginning (day 0) and the end (day 5) of GHS-R antagonist intraperitoneal treatment. Data were analyzed with the unpaired Student t test, and error bars represent SE. ns, not significant. *P < 0.05; **P < 0.01.

MODY3 is characterized by a loss of insulin secretory capacity. Past efforts to better define molecular links between HNF1α function and disease phenotypes have focused on the pancreas (21). Using Hnf1α mutant mice, we identified a novel functional regulatory loop between deregulated production of intestinal ghrelin, restricted potential of insulin secretion, and control of blood glucose homeostasis.

Our data suggest that sustained increases of circulating ghrelin in Hnf1α mutants are dependent on defects from the intestine. This assumption is reasonable given that intestine size is larger than stomach and that Hnf1α mutants display intestinalomegaly (11). Although studies support that pancreatic ε-cells can produce ghrelin (22), attempts to detect ghrelin in the pancreas of Hnf1α mutants was unsuccessful. These observations suggest that specific regulatory mechanisms must occur to differentially regulate expression and/or ghrelin cell commitment in the intestine as compared with other tissues.

The regulatory mechanisms connecting Hnf1α with ghrelin and GIP expression are likely to be complex. Loss of Hnf1α could mechanistically impact enteroendocrine cell fate, including GIP and ghrelin cells. It is also possible that Hnf1α regulates ghrelin and GIP transcription. Bioinformatic analysis of murine ghrelin and GIP gene promoters predicted several Hnf1α elements. In contrast to GIP, this transcriptional connection would imply a negative regulatory loop for ghrelin as it has been suggested for Pax4 transcriptional regulator (23). However, assessment of such mechanisms remains challenging since the population of intestinal ghrelin and GIP cells represents a tiny portion of this epithelium and no normal enteroendocrine cellular models are yet available for such studies.

Although the number of GIP-positive cells and gene transcripts is reduced in the small intestine of Hnf1α mutants, GIP circulating levels are paradoxically increased. Similar observations were reported in patients with type 2 diabetes with exaggerated GIP secretion and dissociated insulin response (24,25). GIP peptide stability could be increased due to the reduction of circulating DPP4 in Hnf1α mutant mice.

In conclusion, pharmacological blockade of ghrelin/GHS-R interaction corrected diabetic features in a MODY3 mouse model. This opens up on preclinical studies targeting MODY3 patients in a program designed to limit ghrelin action and better control blood glucose homeostasis.

Acknowledgments. The authors thank the Electron Microscopy & Histology Research Core of the Faculty of Medicine and Health Sciences at the Université de Sherbrooke for their histology and phenotyping services.

Funding. C.R.L. was a recipient of a Natural Sciences and Engineering Research Council of Canada fellowship. This study was supported by a grant from the Canadian Institutes of Health Research (MOP-126147 to F.Bo.). P.S. and F.Bo. are members of the Fonds de recherche du Québec–Santé–funded Centre de Recherche du CHUS.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. F.Br., C.R.L., and K.B. designed, researched data, and reviewed, edited, and approved the final version of the manuscript. P.S. designed and approved the final version of the manuscript. F.Bo. designed, researched data, wrote the manuscript, and reviewed, edited, and approved the final version of the manuscript. F.Bo. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.

Prior Presentation. Data from this study were presented in part at the 2013 Canadian Digestive Diseases Week, Victoria, Canada, 1–5 March 2013, and at the Digestive Disease Week, Orlando, FL, 18–21 May 2013.

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