Twenty-five male Sprague- Dawley rats (6 weeks old, weighing approximately 150–180 g) were obtained from the Experimental Animal Centre of Anhui Medical University (Hefei, China). The rats were randomly divided into the following four groups: i) control group (n=7); ii) DM group (n=6); iii) DM + low-dose sitagliptin (10 mg/kg) treatment group (n=6); and iv) DM + high-dose sitagliptin (30 mg/kg) treatment group (n=6). All groups were subjected to a 12:12 h light-dark cycle (lights on at 06:00) under controlled conditions of temperature (22±1°C) and humidity (50–60%). With the exception of the control group, all of the rats received a high-fat (HF) diet (2% cholesterol, 10% lard, and 88% normal diet) and sufficient water for 8 weeks, after which time, they were subjected to an intraperitoneal glucose tolerance test (IPGTT) and an intraperitoneal insulin tolerance test (IPITT). The rats in the DM and the sitagliptin treatment groups were injected once intraperitoneally with a dose of streptozotocin (25 mg/kg; Sigma-Aldrich, St. Louis, MO, USA) to induce diabetes and blood glucose levels were tested 1 week after the streptozotocin injection. The animals with glucose levels ≥11.1 mmol/l were considered diabetic. The rats in the control and DM groups were given normal saline. The rats in the low-dose (10 mg/kg) and high-dose (30 mg/kg) sitagliptin (Merck Serono Co., Ltd., Guangzhou, China) groups received sitagliptin once daily by oral gavage for 12 weeks. The rats were weighed every 3 days and the dosage was adjusted accordingly. All the rats were euthanized at the end of week 21. All experiments were approved by the Ethics Committee of Anhui Medical University (Hefei, China).

Non-invasive, transcutaneous ultrasound evaluation of the blood flow velocity was performed two days prior to the end of the experiment as previously described (16,17). Briefly, the rats were anaesthetized using 10% chloral hydrate solution and placed in the dorsal decubitus position for the shaving of their abdomens. After 10 min of rest in the supine position, ultrasonic examination of the abdominal aorta was performed using a 13-MHz ultrasound probe (GES6 two-dimensional Color Doppler Ultrasound Diagnostic Apparatus; GE Healthcare, Piscataway, NJ, USA). The transducer was lubricated with ultrasound gel and placed at the level of the abdominal aorta with minimal pressure. The probe was placed 1.0 cm precisely below the renal artery to obtain a longitudinal axis view of the abdominal aorta. Image settings were optimized to allow the clearest definition of the abdominal aorta. Images of the abdominal aorta were recorded continuously throughout the entire procedure. The ultrasound probe remained in the same position for the duration of the measurement period. The end-diastolic velocity (EDV) and the peak systolic velocity (PSV) were recorded.

Following the withdrawal of food overnight, the rats were deeply anesthetized using 10% chloral hydrate solution (0.3 ml/100 g body weight). Blood samples were obtained from the abdominal aorta. The aortic arch and the thoracic aorta were removed. The aortic arch was longitudinally cut and fixed in 4% paraformaldehyde for subsequent pathological and immunohistochemical assays. Part of the thoracic aortas were placed in a dish containing ice-cold Krebs solution (composition in mmol/l: NaCl, 120; KCl,4.7; KH₂PO₄, 1.18; CaCl₂, 2.25; NaHCO₃, 24.5; MgSO₄•7H₂O, 1.2; glucose, 11.1; EDTA, 0.03) and continuously aerated with 95% O₂ and 5% CO₂. The samples were then cut into rings (3 mm in length) for the measurement of isometric contractile tension. The remaining thoracic aortas were frozen in liquid nitrogen and stored at −80°C for reverse transcription-quantitative polymerase chain reaction (RT-qPCR) and western blot analysis.

Blood samples were taken by cardiac puncture, and the levels of serum fasting blood glucose (FBG), total cholesterol (Tch), low-density lipoprotein cholesterol (LDL-c) and triglyceride (TG) were measured using commercially available spectrophotometric assay kits (Beijing BHKT Clinical Reagent Co., Ltd., Beijing, China). Fasting and postprandial serum GLP-1 levels were measured using an enzyme-linked immunosorbent assay kit (Cusabio, Wuhan, China). Serum levels of ET-1 and insulin were measured using a radioimmunoassay kit (Beifang Biology, Beijing, China). Serum NO levels were measured using a commercial kit (Jiancheng, Nanjing, China). The insulin sensitivity index was calculated using the [1/(FBG × fasting insulin)] formula (18). All analyses were performed in duplicate.

Fixed aortic specimens were dehydrated, embedded in paraffin, sectioned (4 µm thickness) and stained with H&E as previously described (19). The artery wall was observed using a microscope (Leica, Wetzlar, Germany) and the mean artery wall thickness was calculated.

Briefly, the sections of the thoracic aorta were deparaffinised, rehydrated, and fixed with methanol-0.3% H₂O₂ solution for 20 min at room temperature. The antigens were unmasked by cooling for 30 min at room temperature after high compression heating. Primary antibodies against ET-1 (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA) were added and incubated with the tissue sections overnight followed by sequential incubation with a biotinylated antibody (anti-rat IgG) and horseradish streptavidin for 30 min at 37°C. Finally, the samples were incubated with diaminobenzidine for coloration and counterstained with haematoxylin for 2 min to stain the nuclei. Positively-stained cells appeared brown in colour. Images were captured using a microscope (Leica), and the integral optical density values were measured using the JD-801 pathological imaging analysis system (Jiangsu JEDA Science-Technology Development Co., Ltd., Jiangsu, China).

Individual aortic rings were vertically suspended between two stainless steel wire hooks in a jacketed organ bath containing 25 ml Krebs solution. The Krebs solution was replaced at 15-min intervals. The bathing solution was bubbled continuously with a mixture of 95% O₂ and 5% CO₂ at 37°C. The isometric contractile tension was recorded continuously using a BL-420F biological function experimental system (Chengdu Taimeng Science and Technology Co., Ltd., Chengdu, China). The resting tension applied to the aortic rings was increased incrementally to a final tension of 2 g. The rings were then equilibrated for 45 min. Following equilibration, the rings were precontracted with 1 µmol/l phenylephrine (Phe); once a stable contraction plateau was obtained, either 10⁻⁸ to 10⁻⁴ mol/l acetylcholine (Ach) or 10⁻⁸ to 10⁻⁴ mol/l sodium nitroprusside (SNP) was added to the organ bath cumulatively until a maximal vasodilator response was achieved. Cumulative vasodilator response data are expressed as the percentage of relaxation relative to the Phe-induced precontraction.

SYBR-Green RT-qPCR was performed in order to detect the mRNA levels of ET-1. Total RNA was extracted from the aortic tissue of rats using TRIzol reagent (Invitrogen, Grand Island, NY, USA). RNA was reverse transcribed to cDNA using the PrimeScript RT reagent kit (Takara Bio, Otsu, Japan). qPCR was performed with PikoReal (Thermo Fisher Scientific, Waltham, MA, USA) using SYBR Premix Ex Taq (Takara Bio). The relative quantification values for the expression of these genes were calculated using the ΔΔCT method and corrected using a housekeeping gene. The following primer sequences for the analysis of ET-1 and GAPDH mRNA were used: ET-1 forward, 5′-CAACCAGACACCGTCCTCTT-3′ and reverse, 5′-CTTGGAAAGCCACAAACAGC-3′; and GAPDH forward, 5′-TCATTGACCTCAACTACA-3′ and reverse, 5′-CAAAGTTGTCATGGATGACC-3′.

The aortas were washed three times in TBS, and then lysed in RIPA buffer. The lysates were centrifuged at 14,000 × g for 30 min at 4°C. The total protein concentration of each sample was measured using a Micro BCA Protein Assay reagent kit (Pierce, Rockford, IL, USA). The same amount of lysate from each line in SDS sample buffer was resolved by 10% SDS-polyacrylamide gel electrophoresis and electroblotted onto a PVDF membrane, which was then blocked with 5% fat-free milk in TBST (TBS, 0.1% Tween-20) for 1 h at room temperature. The ET-1 Ab (1:250 dilution; sc-21625-R; Santa Cruz Biotechnology, Inc.); phosphorylated (p-)AMP-activated protein kinase (AMPK) (Thr¹⁷²) Ab (1:500 dilution; no. 2535) and p-nuclear factor (NF)-κB p65 Ab (1:500 dilution; no. 3033) (both from Cell Signaling Technology, Inc., Danvers, MA, USA); and p-IκBα Ab (1:250 dilution; sc-8404; Santa Cruz Biotechnology, Inc.) were incubated overnight at 4°C, followed by incubation with secondary antibodies conjugated with horseradish peroxidase (Beijing Zhongshan Jinqiao Biotechnology Co., Ltd., Beijing, China). Specific bands of the target proteins were stained with chemiluminescence reagent (Pierce). Densitometric scanning of the exposed X-ray film was used for semi-quantitative measurement of the protein bands. β-actin levels were used to normalise the relative protein expression levels.

All statistical analyses were performed using SPSS software, version 18.0. All data are presented as the means ± standard deviation (SD). Comparisons between the groups were conducted using one-way analysis of variance (ANOVA) and the Student-Newman-Keuls (SNK) method. A P-value <0.05 was considered to indicate a statistically significant difference.

Following 8 weeks of HF feeding, IPGTT and IPITT results revealed that the blood glucose levels in the rats fed an HF diet were significantly higher than those in the control rats at baseline (IPITT only) and at 15, 30, 60 and 120-min time-points (P<0.05). The area under the curve (AUC) for the glucose level increased significantly in the rats fed an HF diet for 8 weeks (P<0.05) (Fig. 1). These results suggested that the rats fed an HF diet were insulin resistant.

Statistical analysis showed that the body weight and the blood glucose levels were significantly changed in the DM group, and that sitagliptin reduced the extent of weight loss as well as the high FBG levels during the experimental period (Fig. 2). Following 12 weeks of treatment, FBG, Tch, TG, LDL-c and insulin resistance were significantly higher in the diabetic rats compared with the control group (P<0.05). The administration of high-dose sitagliptin significantly reduced FBG levels and insulin resistance (P<0.05). However, the levels of Tch, TG and LDL-c were not significantly affected by low dose of sitagliptin (P>0.05) (Table I).

The induction of diabetes significantly increased the serum levels of ET-1 and reduced the levels of fasting and postprandial GLP-1 as well as NO (P<0.05). The adminstration of high- and low-dose sitagliptin significantly reduced the serum levels of ET-1 and elevated the serum levels of GLP-1 as well as NO to varying extents (P<0.05) (Fig. 3).

We then examined the effects of sitagliptin on vascular endothelial function in the diabetic rats. Isometric contractile tension experiments on isolated aortic rings were performed. As illustrated in Fig. 4, in the DM group, we observed inhibition of the cumulative contraction-relaxation response to Ach in the thoracic aortic rings compared with the controls (P<0.05), whereas there were no significant changes in the cumulative contraction-relaxation response to SNP (P>0.05). The administration of different doses of sitagliptin partly reversed the diabetes-induced impairment of the cumulative concentration-vasodilator responses to Ach in the thoracic aortic rings precontracted with Phe (P<0.05) and had no effect on the SNP-induced cumulative contraction-relaxation responses (Fig. 4C and D). The contractile response to Phe in the untreated diabetic rats was lower than that in the control rats and the rats treated with high-dose sitagliptin (Fig. 4B). In addition, performing non-invasive transcutaneous ultrasound measurement of blood flow velocity in the abdominal aorta showed that PSV was significantly reduced in the DM group, and was not significantly increased by the administration of sitagliptin (P<0.05) (Fig. 4A).

We examined the morphological characteristics of the arterial lesions in the H&E-stained tissue sections. The endothelial cells adhering to inner elastic plates showed structural changes in the DM group compared with those in the control group. Indeed, the vascular smooth muscle cells (VSMCs) of the tunica media were arranged irregularly, as shown by irregular nuclei and uneven staining of the cytoplasm, and the media layer thickness of the aorta was also markedly increased. However, the administration of sitagliptin attenuated the remodeling of the aortic intima, corrected the disarrangement of the VSMCs and reduced the media thickness of the aorta (Fig. 5A and C).

As shown in Fig. 5B and D, a high ET-1 expression level was found in the intima of the DM rats, in comparison with that in the control rats (P<0.05). High- and low-dose administration of sitagliptin led to a significant downregulation of ET-1 expression in the intima of the arteries (P<0.05).

As shown in Fig. 6A and B, hyperglycaemia and dyslipidaemia stimulated ET-1 expression at the transcriptional and translational levels, whereas the administration of sitagliptin significantly inhibited ET-1 expression (P<0.05). Western blot analysis also revealed that the rats in the DM group exhibited markedly enhanced expression of p-NF-κB, NF-κB and p-IκBα (P<0.05), and reduced expression of p-AMPK as well as IκBα (P<0.05) (Fig. 6C–E). Notably, the administration of sitagliptin reversed these effects on protein expression (P<0.05), thus, reversing the detrimental effects of hyperglycaemia and dyslipidaemia in the rat aorta.