Sitagliptin-mediated preservation of endothelial progenitor cell function via augmenting autophagy enhances ischaemic angiogenesis in diabetes

Animals

db/db (FVB background) male mice at 2–3 months of age were used in this study. All animal procedures were approved by the Animal Policy and Welfare Committee of Wenzhou Medical University and/or the Institutional Animal Care and Use Committee of the University of Louisville, which conform to the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (publication No. 85-23, revised 1996).

HLI and sitagliptin treatment

The db/+ founder mice [FVB.BKS(D)-Lepr^db/+/ChuaJ, FVB background] were purchased from the Jackson Laboratory (Bar Harbor, ME, USA) and maintained under specific pathogen-free conditions at the University of Louisville Animal Facility (Louisville, KY, USA) or the Wenzhou Medical University. The db/db mice were generated by breeding male db/+ to female db/+ mice following Jackson Laboratory's instructions. Twelve-week-old male db/db mice were equally divided into two groups (n = 16/group) to develop the HLI model as described in our previous report 12. One group was pretreated with sitagliptin (25 mg/kg, daily) for 1 week before HLI surgery and continually treated with sitagliptin for an additional 7 or 35 days after surgery via gavage administration. Another group was treated with H₂O as the vehicle control. At 7 or 35 days after surgery, eight mice from each group were sacrificed to collect blood and gastrocnemius muscle samples. The HLI procedure is briefly described below: under sufficient anaesthesia with isoflurane (1–3% isoflurane in 100% oxygen at a flow rate of 1 l/min.), the hind limbs were shaved and the entire right superficial femoral artery and vein (from just below the deep femoral arteries to the popliteal artery and vein) were ligated with 6-0 silk sutures, cut and excised with an electrical coagulator (Fine Science Tools Inc., Foster City, CA, USA). The overlying skin was closed with 4-0 silk sutures.

Measurement of blood flow perfusion with a Pericam perfusion speckle imager (PSI)

To evaluate the limb perfusion ratio [ischaemic limb (right)/normal limb (left)], real-time microcirculation imaging analysis was performed using a Pericam PSI based on laser speckle contrast analysis technology (Perimed Inc., Kings Park, NY, USA) before surgery and at day 0, 3, 7 14, 21, 28 and 35 post-surgery.

Quantification of circulating EPCs by flow cytometry

At days 3 and 7 after surgery, blood samples were collected in 0.1 mol/l EDTA-2Na-coated tubes from the tail vein. Whole blood was incubated with CD34-PE and VEGFR-2-APC antibodies 13, 14 (BD Pharmingen, San Jose, CA, USA) for 1 hr, then lysed and fixed in FACS™ lysing solution (BD Biosciences, San Jose, CA, USA) for 5 min. Flow cytometry analysis was performed with a BD FACS (BD Biosciences) to count VEGFR-2 and CD34 double-positive EPCs. Isotype control IgG (BD Pharmingen) was used to exclude false-positive cells.

Determination of SDF-1 and glucagon-like peptide-1 (GLP-1) in plasma

At day 14 after sitagliptin administration (7 days after surgery), the plasma was collected to detect the levels of SDF-1 and GLP-1. The levels of SDF-1α and GLP-1 were measured with commercially available enzyme-linked immunosorbent assay (ELISA) kits CXCL12/SDF-1α immunoassay (R&D systems, Minneapolis, MN, USA) and Glucagon Like Peptide-1 (Active) ELISA kit (Millipore, Billerica, MA, USA), respectively.

Histological assessment

The extent of angiogenesis at day 35 post-ischaemic surgery was assessed by measuring capillary density using isolectin B4 staining. Ischaemic gastrocnemius muscle tissues were fixed with 4% paraformaldehyde and embedded with paraffin. Paraffin sections (5 μm) were stained with Alexa Fluor^® 594 conjugated isolectin GS-IB4 antibody (Thermo Scientific, Waltham, MA, USA) to evaluate the capillary density. The capillaries were counted in randomly selected fields for a total of 20 different fields (×40 magnification) per section and three sections per animal. The capillary density is presented as capillary number per muscle fibre.

Isolation and culture of bone marrow-derived EPCs

EPCs were isolated from the bone marrow of wild-type (WT) FVB mice and cultured according to our established methods with minor modifications 15. Briefly, bone marrow mononuclear cells (MNCs) were isolated from the femurs and tibias of the mice by density gradient centrifugation with histopaque-1083 (Sigma-Aldrich, St. Louis, MO, USA). After two washes, MNCs were plated on vitronectin-coated culture dishes (Sigma-Aldrich) and maintained in endothelial growth factor-supplemented media (EGM-2 bullet kit; Lonza, Basel, Switzerland) with 10% foetal bovine serum. Cells were cultured at 37°C with 5% CO₂ in a humidified atmosphere. EGM-2 medium was replaced after the first 24 hrs and every 3 days thereafter. Cell colonies appeared after 7 days of culture and were expanded to the fourth or fifth passage for further analysis.

Characterization of bone marrow-derived MNCs

After 14 days of culture in endothelial-specific media and the removal of non-adherent MNCs, the remaining cells were characterized by uptake of Dil-Ac-LDL and ulex europaeus lectin-1 binding. For the uptake of Dil-Ac-LDL and ulex europaeus lectin-1 binding assay, cells were seeded on a vitronectin-coated 8-well μ-slide (ibidi, Martinsried, Germany) one day before the procedure, and then, the cells were incubated with 5 μg/ml acetylated DiI lipoprotein from human plasma (Dil-Ac-LDL, Thermo Fisher Scientific, Waltham, MA, USA) at 37°C for 4 hrs, followed by three washes with Dulbecco's phosphate-buffered saline (DPBS) and fixed with 4% paraformaldehyde. Then, the cells were incubated with 10 μg/ml fluorescein isothiocyanate-labelled ulex europaeus lectin-1 (FITC-UEA-1; Sigma-Aldrich) for 1 hr at room temperature. After incubation, the cells were rinsed with DPBS three times and were visualized via confocal microscopy.

To further confirm the identity of these cells, immunofluorescence staining was performed to analyse the expression of specific cell markers VEGFR2 and Sca-1. For the immunofluorescence staining, cells were seeded on a vitronectin-coated 8-well μ-slide one day before the procedure and then fixed with 4% paraformaldehyde after three washes with DPBS. The fixed cells were incubated with PE-VEGR2 and FITC-Sca-1 16, 17 antibodies (Abcam, Cambridge, MA, USA) overnight at 4°C. DAPI (4′,6-diamidino-2-phenylindole, dihydrochloride) was used to label nuclei.

DPP-4 enzymatic activity assay

DPP-4 enzymatic activity was assayed in the culture medium of EPCs under different treatment conditions using a DPP-4 Activity Assay Kit (Sigma-Aldrich). EPCs were seeded on 12-well plates and maintained under basal culture conditions with or without HG (25 mmol/l) for 24 hrs 18 in the presence or absence of different concentrations of sitagliptin. The culture medium was collected for the DPP-4 enzymatic activity assay. Briefly, a 50 μl volume of medium was diluted with 48 μl of DPP-4 assay buffer, mixed with 2 μl substrate gly-pro-7-amino-4-methylcoumarin (AMC) and then incubated at 37°C for 30 min. The release of AMC from the substrate was measured with a fluorescence spectrophotometer at 360 nm excitation and 460 nm emission.

Apoptosis assay

EPCs were seeded on 12-well plates (1 × 10⁵ cells/well) and maintained under basal culture conditions or with HG (25 mmol/l) for 24 hrs in the presence or absence of sitagliptin. Non-adherent cells were removed by washing with PBS. In some experiments, cells were pretreated with the autophagy inhibitor 3-methyladenine (3-MA, 5 mmol/l; Sigma-Aldrich) or the autophagy activator rapamycin (10 μmol/l; Sigma-Aldrich) for 30 min. and then continually exposed to HG for the indicated durations in the presence or absence of sitagliptin. Subsequently, adherent cells were released with 0.25% trypsin without EDTA. EPCs were collected by centrifugation and stained with an APC-conjugated Annexin V Apoptosis Detection Kit according to the manufacturer's instructions (Biolegend, San Diego, CA, USA). Apoptotic EPCs were detected by flow cytometry. Early apoptotic cells were defined as AnnexinV⁺/PI^–.

Tube formation assay

The in vitro angiogenic capability of EPCs was determined by a Matrigel tube formation assay. Briefly, 48-well plates were coated with growth factor-reduced matrix gel (150 μl/well, BD Biosciences). EPCs (5 × 10⁴ cells/well) in 200-μl basal culture medium or medium containing HG in the presence or absence of different concentrations of sitagliptin were incubated at 37°C with 5% CO₂ for 12 hrs to form tubes. In some experiments, cells were pretreated with 3-MA (5 mmol/l) or rapamycin (10 μmol/l) for 30 min. and then continually exposed to HG for the indicated durations in the presence or absence of sitagliptin. Images of tubes in each well were acquired using an inverted microscope (Nikon Eclipse E600, Nikon, Kanagawa, Japan). The tube lengths were calculated by ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Quantitative determination of oxidative stress

To detect the reactive oxygen species (ROS) levels of EPCs with HG treatment in the presence or absence of different concentrations of sitagliptin, a dihydroethidium (DHE; Molecular Probes, Eugene, OR, USA) probe was used to stain EPCs. DHE is cell permeable and able to react with superoxide to form ethidium, which in turn intercalates with DNA and produces nuclear fluorescence. EPCs were seeded on 24-well plates, treated with HG in the presence or absence of sitagliptin for 6 hrs and then incubated with 5 μmol/l DHE in PBS for 30 min. at 37°C. Nuclear DHE-positive staining indicates superoxide generation in cells. Fluorescence intensity was detected by a microplate reader (SpectraMax M3; Molecular Devices, Sunnyvale, CA, USA) under specific wave length conditions (excitation = 518 nm; fluorescence = 605 nm). Meanwhile, fluorescence images of random EPCs were captured at 20× magnification (XI 71 Olympus, Tokyo, Japan).

Autophagic signalling assay

Autophagic signalling was assayed by detecting microtubule-associated protein 1A/1B-light chain 3 (LC3) and p62 protein expression levels and LC3-bound puncta formation. For the LC3 and p62 protein expression assay, EPCs were seeded on 6-well plates (1 × 10⁵ cells/well) and maintained under basal culture conditions with or without HG for 24 hrs in the presence or absence of sitagliptin. Non-adherent cells were removed by washing with PBS. In some experiments, cells were pretreated with AMP-activated protein kinase (AMPK) inhibitor compound C (10 μmol/l; Sigma-Aldrich) for 30 min. and then continually exposed to HG for the indicated durations in the presence or absence of sitagliptin. Then, EPCs were harvested for a Western blot assay of LC3 and p62 protein expression levels. For the LC3-bound puncta assay, EPCs were seeded on an 8-well microculture chamber (2 × 10⁴ cells/well) and maintained under basal culture conditions with or without HG for 24 hrs in the presence or absence of sitagliptin. Non-adherent cells were removed by washing with PBS. The remaining cells were stained with a LC3B primary antibody (1:100; Novus Biologicals, Littleton, CO, USA) and an appropriate FITC-conjugated secondary antibody. LC3 puncta were observed with immunofluorescence microscopy.

Western blot assay

Western blotting was performed as described in a previous study 15. Gastrocnemius muscle tissues and harvested cells were homogenized or lysed in ice-cold RIPA lysis buffer (Santa Cruz Biotechnology, Dallas, TX, USA). The protein concentration was determined using a Bradford protein assay kit (Bio-Rad, Hercules, CA, USA). The total proteins were separated by 10% sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to nitrocellulose membranes (Bio-Rad). The membranes were blocked in tris-buffered saline with 5% non-fat milk and 0.5% bovine serum albumin for 1 hr and then incubated with primary antibody overnight at 4°C, followed by an incubation with secondary antibodies for 1 hr at room temperature after standard washing procedures. The primary antibodies against VEGF (1:10,000), glyceraldehyde 3-phosphate dehydrogenase (GAPDH, 1:5000) and β-actin (1:3000) were purchased from Santa Cruz Biotechnology; SDF-1 (1:2000), AMPK and p-AMPK (1:2000), mTOR and p-mTOR (1:2000), unc-51-like autophagy activating kinase 1 (ULK1) and p-ULK1 and P62 (1:2000) were purchased from Cell Signaling Technology (Danvers, MA, USA); and LCB (1:2000) was purchased from Novus Biologicals (Littleton, CO, USA). All horseradish peroxidase (HRP)-conjugated secondary antibodies were purchased from Santa Cruz Biotechnology. Blots were visualized with SuperSignal West Femto Maximum Sensitivity Substrate (Thermo Scientific) and quantified with Quantity 5.2 software (Bio-Rad).

Statistical analysis

All data are presented as the mean ± S.D. Statistical analysis was performed using Origin 7.5 (OriginLab data analysis and graphing software, Northampton, MA, USA) with one-way or two-way anova, followed by post hoc multiple comparisons with Scheffe's test. Statistical significance was considered P < 0.05.

Sitagliptin enhances blood perfusion and angiogenesis in HLI in type 2 diabetic mice

To evaluate the angiogenic effect of DPP-4 inhibition in diabetes, db/db mice were pretreated with sitagliptin (Januvia) for 1 week; then, the mice were subjected to HLI surgery and continually treated with sitagliptin for an additional 5 weeks until to the end of the experiments. Blood flow was evaluated by PSI at day 3, 7, 14, 21, 28 and 35 after surgery. The results showed that sitagliptin treatment time dependently improved blood perfusion in db/db mice underwent HLI (Fig. 1A), which was accompanied by promoted angiogenesis mirrored by increased capillary density (Fig. 1B).

Sitagliptin increases proangiogenic factor expression levels and enhances EPCs mobilization in type 2 diabetic mice

To evaluate the effect of DPP-4 inhibition on the expression levels of proangiogenic factors in plasma and ischaemic tissue, peripheral blood and gastrocnemius muscles in ischaemic limbs were collected at day 7 after surgery. The ELISA results showed that mice that received sitagliptin had elevated levels of GLP-1 and SDF-1 in plasma (Fig. 2A and B). Proangiogenic factors in gastrocnemius muscle were detected by Western blotting, and the results showed that VEGF and SDF-1 increased significantly in sitagliptin-treated mice (Fig. 2C).

At days 3 and 7 after surgery, peripheral blood was collected to evaluate the number of EPCs in circulation. A flow cytometry assay showed that the number of circulating EPCs (CD34+/VEGFR2+) significantly increased in sitagliptin-treated mice (Fig. 2D), indicating that sitagliptin promoted EPCs mobilization from bone marrow to peripheral blood.

Characterization of bone marrow-derived EPCs

MNCs isolated from bone marrow were cultured in EGM-2 medium on vitronectin-coated culture dishes. By 14–21 days of culture, endothelium-like cells with the cobblestone-like morphology of late EPCs were nearly confluent. Late EPCs were further characterized by Dil-ac-LDL uptake and FITC-UEA-1 lectin binding. As shown in Figure 3A, most late EPCs were positive for Dil-ac-LDL and FITC-UEA-1 lectin staining. To further confirm the identity of these cells, immunofluorescence staining was performed to analyse the expression of specific cell markers (Fig. 3B, upper panel). The results showed that the majority of cells are both VEGFR2 and Sca-1 positive (Fig. 3B, lower panel).

Sitagliptin improves the survival and angiogenic function of EPCs treated with HG

To validate the potency of sitagliptin, DPP-4 activity was measured in the medium of EPCs using a DPP-4 Activity Assay Kit. As expected, HG increased the DPP-4 activity of EPCs, which was inhibited by sitagliptin treatment in a dose-dependent manner (Figure S1). To determine whether DPP-4 inhibition could restore the survival and angiogenic function of EPCs under diabetic conditions, EPCs were exposed to HG to mimic the dysmetabolic stress of hyperglycaemia. The results demonstrated that HG significantly increased EPC apoptosis, as measured by Annexin V/PI staining (Fig. 4A). Sitagliptin treatment dose dependently prevented HG-induced apoptosis of EPCs within the lower dose range, although this protection was blunted at higher doses (Fig. 4A). Tube formation assays were used as one measure of angiogenic function in vitro. Similarly, HG significantly impaired the tube formation abilities of EPCs, and sitagliptin treatment significantly prevented the detrimental effects of HG on EPCs function at the same dose levels that prevented apoptosis (Fig. 4B).

Sitagliptin attenuates ROS levels in EPCs treated with HG

Oxidative stress is a major factor responsible for the dysfunction of diabetic EPCs 19, 20. We used DHE staining to show superoxide production in EPCs. The results demonstrated that HG significantly increased superoxide production levels, whereas sitagliptin treatment attenuated HG-induced superoxide production in EPCs at the same optimal dose level that prevented apoptosis and protected angiogenic function (Fig. 4C). Therefore, the optimal dose of sitagliptin at 3 μmol/l in EPCs protection was used in the following mechanistic studies.

Sitagliptin improves the survival and function of EPCs treated with HG via augmenting autophagy

Under diabetic conditions, impaired autophagy contributes to endothelial dysfunction, and restoration of autophagy can improve endothelial cell survival and function 21. In this study, we found that sitagliptin improved the survival and function of EPCs treated with HG. Therefore, we investigated whether the protective effects of sitagliptin on EPCs were mediated by autophagy activation in the following studies. EPCs were treated with HG (25 mmol/l) in the presence or absence of sitagliptin (3 μmol/l), and cells treated with 5.5 mmol/l glucose plus 20.5 mmol/l mannitol were used as a control. Immunofluorescence staining analysis demonstrated that elevated glucose levels reduced the number and distribution of LC3 puncta staining, suggesting a reduction in autophagosome formation, which was significantly prevented by sitagliptin treatment (Fig. 5A). In addition, we also detected the expression of p62 and the ratio of LC3II to LC3I by Western blotting. The results showed that the ratio of LC3II to LC3I decreased in EPCs with HG treatment, which was reversed by sitagliptin, whereas p62 expression exhibited the opposite pattern (Fig. 5B). These results indicate that sitagliptin activates autophagy in EPCs under HG treatment conditions.

To confirm whether the protective effects of sitagliptin on EPCs were dependent on augmented autophagy, the autophagy inhibitor 3-MA and the activator rapamycin were tested against the protection of sitagliptin. The result showed that inhibition of autophagy by 3-MA completely abolished the sitagliptin-stimulated increase in LC3II and decrease in p62 expression in EPCs (Fig. 6A), resulting in complete abolishment of sitagliptin-induced cell survival (Fig. 6B) and tube formation (Fig. 6C) in EPCs under HG treatment conditions, whereas activation of autophagy by rapamycin almost completely prevented HG-induced pathological changes in EPCs, which are comparable to that of sitagliptin treatment (Fig. 6A–C). These results suggest that sitagliptin preserves EPC function via restoring autophagy in EPCs.

Sitagliptin restores autophagy in EPCs treated with HG via activating the AMPK/ULK1 signalling pathway

AMPK/ULK1 signalling is critical for autophagy regulation under diabetic conditions 22, 23. A previous study has shown that sitagliptin can activate AMPK 24. Therefore, we first detected AMPK activation in EPCs. The results showed that HG inhibited AMPK activity, as estimated by the decreased phosphorylation of AMPK, whereas sitagliptin treatment prevented HG-induced decreases in the phosphorylation of AMPK (Fig. 7A). Meanwhile, we also found that sitagliptin prevented HG-induced phosphorylation of mTOR, which was accompanied by an increase in the phosphorylation of ULK1 at Ser555 (Fig. 7A). To further confirm the critical role of AMPK activation in sitagliptin-induced augmentation of autophagy in EPCs, the AMPK inhibitor compound C was tested against sitagliptin-induced activation of autophagy. The result showed that compound C completely abolished the sitagliptin-stimulated increase in LC3II and decrease in p62 expression in EPCs (Fig. 7B). These results suggest that AMPK is a critical mediator of the protective effects of sitagliptin on EPC function via augmenting autophagy.