Sitagliptin (phosphate) was provided by Cayman Chemical Company (Ann Arbor, MI, USA) and 5-aminoimidazole-4-carboxamide riboside (AICAR) was purchased from Beyotime Institute of Biotechnology (Haimen, China). Compound C was obtained from Sigma-Aldrich; Merck KGaA (Darmstadt, Germany). Monoclonal rabbit anti-phospho-AMPKα antibody (catalog no. 2535p) and anti-AMPK antibody (catalog no. 2603p) were obtained from Cell Signaling Technology, Inc. (Danvers, MA, USA). The following antibodies were also used: Monoclonal mouse anti-β-actin antibody (catalog no. sc-47778) and horseradish peroxidase-conjugated goat anti-rabbit/mouse secondary antibody (catalog no. sc-2004/sc-2005) from Santa Cruz Biotechnology, Inc. (Dallas, TX, USA).

HUVECs were isolated by collagenase digestion from fresh cord umbilical veins, as previously described (14). The flesh cord umbilical veins were obtained from normal cesarean section surgery. This was approved by Air Force General Hospital ethics committee with informed written consent. HUVECs between passages 3 and 6 were cultured in endothelial cell medium (ScienCell Research Laboratories, Inc., Carlsbad, CA, USA) containing basal medium, supplemented with 5% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) and 1% endothelial cell growth supplement with antibiotics (100 U/ml penicillin G and 100 µg/ml streptomycin sulfate). This was conducted in a humidified atmosphere containing 5% CO2, at 37°C.

To determine the effect of sitagliptin on AMPK activation, the HUVECs were treated with 1 µM sitagliptin for 0.5, 1, 2 and 4 h or 100 µM AMPK activator AICAR, for 0.5 h. To detect the inhibitory effect of the AMPK inhibitor compound C on sitagliptin-induced AMPKα phosphorylation, HUVECs were incubated with 1 µM sitagliptin, 10 µM compound C or 1 µM sitagliptin plus 10 µM compound C for 2 h. The cytoplasmic protein of cells was extracted with ice-cold hypotonic lysis buffer [50 mM Tris-HCl, pH 7.5, 15 mM EGTA, 0.1% (vol/vol) Triton X-100, 100 mM NaCl and complete protease inhibitor cocktail] as previously described (15). Cell lysates were first snap frozen in liquid nitrogen and then centrifuged at 12,000 × g at 4°C for 20 min, for collection of the supernatant. Protein concentration was measured using the BCA method. Equal amounts of protein (10 µg per sample) were separated on 10% sodium dodecyl sulfate-polyacrylamide gels electrophoresis and blotted onto polyvinylidene difluoride membranes. Following incubation with no fat milk at 25°C for 20 min, the membranes were reacted with anti-phospho-AMPKα antibody (1:1,000) and anti-AMPKα antibody (1:1,000) at 4°C overnight, then reacted with appropriate horseradish peroxidase-conjugated secondary antibodies (1:3,000) for 2 h at 25°C. Proteins were visualized with an enhanced chemiluminescence kit, as previously described (16). Densitometry analysis was performed for three independent experiments using the Image J Gel Analysis tool (National Institutes of Health, Bethesda, MD, USA).

HUVECs (1×10⁵) were incubated with HG (33 mM) in the presence of 1 µM sitagliptin, 100 µM AICAR or 1 µM sitagliptin plus 10 µM AMPK inhibitor compound C for 48 h. Induction of apoptosis in the treated groups was assessed by Annexin V-fluorescein isothiocyanate (FITC)/propidium iodide (PI) double staining detection kit (Nanjing KeyGen Biotech Co., Ltd., Nanjing, China), according to the manufacturer's protocol. Briefly, cells were incubated with 33 mM D-glucose in the presence of the aforementioned agents for 48 h and gently digested with 1 ml 0.25% trypsin (Thermo Fisher Scientific, Inc.) for 2 min. The trypsinized cells were washed once with endothelial cell medium containing 5% fetal bovine serum prior to collection by centrifugation at 1,000 × g and room temperature for 20 min. Cells were resuspended in 500 µl of 1X binding buffer, followed by incubation with 5 µl of Annexin V-FITC and 5 µl of PI (50 µg/ml) for 10 min in the dark. Binding buffer, Annexin V-FITC and PI are components of the detection kit. All procedures subsequent to cell incubation were performed at room temperature. Stained cells were monitored by flow cytometry (BD FACSCalibur; BD Biosciences, San Jose, CA, USA) and analyzed with BD FACSDiva^™ software (version 6.0; BD Biosciences).

A ROS-specific fluorescent probe, 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA; Molecular Probes; Thermo Fisher Scientific, Inc.) was used for the measurement of cytosolic ROS production. HUVECs were incubated with 33 mM D-glucose in the presence of 1 µM sitagliptin, 100 µM AICAR or 1 µM sitagliptin plus 10 µM compound C for 48 h, then cells were stained with 10 µM H2DCFDA fluorescent probe in serum-free endothelial cell medium at 37°C for 30 min. The labeled cells were then washed twice with serum-free endothelial cell medium and the levels of ROS were immediately analyzed by flow cytometry (BD FACSCalibur; BD Biosciences).

To measure ΔΨm, HUVECs were treated with 1 µM sitagliptin, 100 µM AICAR or 1 µM sitagliptin plus 10 µM compound C prior to exposure to 33 mM D-glucose for 48 h. Following incubation, cells were collected and stained with 2 µM ΔΨm-specific fluorescent dye JC-1 (Molecular Probes; Thermo Fisher Scientific, Inc.) at 37°C in an atmosphere containing 5% CO2, for 20 min. Flow cytometry (BD FACSCalibur; BD Biosciences) was used to detect ΔΨm for each treatment group. JC-1 accumulates in mitochondria in a potential-dependent manner. In normal mitochondria with high ΔΨm, JC-1 aggregates with red fluorescence. In apoptotic cells with injured mitochondria membrane, JC-1 alters to monomers, and emits green fluorescence. ΔΨm is determined by red/green fluorescence intensity ratio.

Data are expressed as the mean ± standard error of the mean. One-way analysis of variance was used to determine differences among the mean values of treatments. SPSS software, version 20.0 (IBM SPSS, Armonk, NY, USA) was used for the statistical data analysis. P<0.05 was considered to indicate a statistically significant difference.

The present study examined the effect of sitagliptin on HUVECs incubated with HG. Cell apoptosis of the pretreated groups was measured by Annexin V-FITC/PI double staining and monitored by flow cytometry (Fig. 1A). It was observed that HG significantly increased cell apoptosis, and this HG-induced endothelial cell apoptosis was prevented by sitagliptin or the AMPK activator, AICAR. However, compound C, an AMPK inhibitor, reversed the inhibition of apoptosis by sitagliptin (Fig. 1B). This therefore indicated that AMPK is important in the regulatory action of sitagliptin.

As AMPK was observed to be involved in sitagliptin-mediated prevention of endothelial cell apoptosis, the present study aimed to determine the effect of sitagliptin on AMPK activity. HUVECs were incubated with 1 µM sitagliptin at different times ranging from 0.5–4 h. Phosphorylation of AMPKα (p-AMPKα) was determined by western blotting (Fig. 2A). Sitagliptin stimulated AMPKα (Thr¹⁷²) phosphorylation from 2 h, and this phosphorylation activity prevailed until 4 h. AICAR enhanced AMPK phosphorylation in endothelial cells in a similar manner to sitagliptin, following incubation with the cells for 0.5 h (Fig. 2B). The effect of compound C on sitagliptin-induced AMPKα phosphorylation was additionally examined (Fig. 2C). As presented in (Fig. 2D), sitagliptin-stimulated AMPKα activation was significantly inhibited by compound C. These findings suggested that sitagliptin induces AMPKα phosphorylation.

In vascular endothelial cells, the hyperglycemia load increases generation of ROS (17), which subsequently contributes to cell apoptosis. To observe the effect of sitagliptin pretreatment on HG-induced cytosolic ROS generation, cytosolic ROS levels were detected via flow cytometry (Fig. 3A). It was observed that high glucose significantly increased ROS production, however this was suppressed with pretreatment with 1 µM sitagliptin. In addition, AICAR effectively inhibited generation of ROS, whereas compound C diminished the inhibitory effect of sitagliptin (Fig. 3B). These data suggested that sitagliptin inhibits cytosolic ROS via AMPK activation. The effect of sitagliptin on ROS-mediated mitochondrial dysfunction, under conditions of hyperglycemia were then examined.

ROS-mediated ΔΨm collapse was previously demonstrated to initiate mitochondrial-dependent apoptosis in DM (18). The present study proceeded to characterize HG-induced ΔΨm alterations and examine if sitagliptin protects against ΔΨm collapse. JC-1 staining detection by flow cytometry was performed. Mitochondrial depolarization is determined by a decrease in aggregate/monomer fluorescence ratio (Fig. 4A). In a similar manner to that exhibited by AICAR, 1 µM sitagliptin restored HG-induced ΔΨm collapse, and this effect was blocked by compound C (Fig. 4B). These results suggested that AMPK is important in the regulatory actions of sitagliptin in HG-induced endothelial apoptosis.