HUVECs (cat. no. iCell-h110) were purchased from iCell Bioscience, Inc. and were incubated in Endothelial Cell Culture Medium (Cellverse Bioscience Technology Co., Ltd.) containing 100 U/ml penicillin and 100 µg/ml streptomycin in the presence of 5% CO₂ at 37˚C. To mimic AGE-induced DVT, HUVECs were treated with modified glycated human serum albumin (M-HSA) for 24 h (26) and M-HSA was prepared by co-incubation of HSA (15 mg/ml; Sigma-Aldrich; Merck KGaA) and 3-deoxyglucosone (3-DG, 1 mM; Sigma-Aldrich; Merck KGaA) for 2 weeks as described in a previous study conducted by the authors (16). Tunicamycin (TM; Sigma-Aldrich; Merck KGaA), the agonist of ERS, was used for pre-treatment (5 µg/ml) for 6 h prior to M-HSA induction in HUVECs.

The sequences of PPARγ were cloned into the pcDNA 3.1 vector (Invitrogen; Thermo Fisher Scientific, Inc.) to establish a PPARγ overexpression vector (oe-PPARγ). The pcDNA 3.1 vector was used as a negative control (oe-NC). HUVECs were transfected with oe-PPARγ (15 nM) or oe-NC (15 nM) using Lipofectamine™ 3000 reagent (Thermo Fisher Scientific, Inc.) at 37˚C strictly in line with the manufacturer's guidelines upon reaching 80% confluence. At 48 h post-transfection, the transfected HUVECs were harvested for subsequent experiments.

PPARγ activity was evaluated as previously reported (27). In brief, nuclear extracts were initially obtained from cultured HUVECs using a Nuclear Extraction Kit (cat. no. ab113474; Abcam). Subsequently, PPARγ activity was determined using a PPARγ Transcription Factor Assay Kit (cat. no. ab133101; Abcam) by measuring the absorbance at 450 nm.

Cell viability was determined using the Cell Counting Kit-8 (CCK-8) assay (16). In brief, HUVECs were cultured in 96-well plates (3.0x10³ cells/well) and incubated for 24, 48 and 72 h, respectively. At different time points, 10 µl CCK-8 solution (cat. no. KGA9305-500; Nanjing KeyGen Biotech Co., Ltd.) was added into each well and HUVECs were cultured in the incubator for an additional 2 h at 37˚C. Finally, the absorbance at 450 nm of each well was detected using a microplate reader.

Cell apoptosis was assessed using the TUNEL assay (28). HUVECs (5x10⁴ cells/well) were cultured in 6-well plates with cell culture silicon slides. Following treatment, the cells were fixed with 4% paraformaldehyde for 30 min, permeabilized with 0.2% Triton X-100 for 5 min and blocked with 3% BSA (Wuhan Servicebio Technology Co., Ltd.) for 30 min at room temperature. Subsequently, the cells were incubated with a TUNEL reaction mixture (Roche Diagnostics) at 37˚C for 1 h and the cell nuclei were stained with 1 mg/ml 4',6-diamidino-2-phenylindole (Invitrogen, Thermo Fisher Scientific, Inc.) for 10 min at 37˚C in the dark. Anti-fluorescence quenching liquid was used for sealing. The apoptotic cells were observed in five random fields using an inverted fluorescence microscope (Olympus IX71; Olympus Corporation).

The culture medium was harvested and centrifugated at 12,000 x g, 4˚C for 10 min and the supernatant was then collected. ELISA kits for 6-keto prostaglandin-F1 α (6-K-PGF1α; E-EL-0054, Elabscience), ET-1 (ml025101), TNF-α (ml077385), IL-1β (ml058059) and IL-6 (ml028583; all from Shanghai Enzyme-linked Biotechnology Co., Ltd.) were applied to evaluate the corresponding protein levels in the culture supernatant in accordance with the manufacturer's instructions (16).

The levels of ROS, malondialdehyde (MDA) and superoxide dismutase (SOD) were detected to evaluate the degree of oxidative stress (16). For ROS measurement, HUVECs were stained with 20 µM 2',7'-dichlorodihydrofluorescein diacetate strictly in line with the instructions of the Fluorometric Intracellular ROS Kit (cat. no. MAK143; Sigma-Aldrich; Merck KGaA). The fluorescence intensity was detected by a microplate fluorometer (Molecular Devices, LLC). For MDA and SOD measurements, the cell supernatant was measured with Lipid Peroxidation MDA Assay Kit (cat. no. S0131S) and Total Superoxide Dismutase Assay Kit (cat. no. S0101S) (both from Beyotime Institute of Biotechnology) in accordance with the manufacturer's guidelines, respectively.

Total protein was extracted from cells using a radioimmunoprecipitation assay lysis buffer (Beyotime Institute of Biotechnology), followed by the determination of protein concentrations using an Enhanced BCA protein assay kit (Beyotime Institute of Biotechnology). The same amount (30 µg/lane) of protein was separated by 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis and transferred onto polyvinylidene fluoride membranes. Following blocking with 5% non-fat milk at room temperature for 2 h, the membranes were incubated with primary antibodies against PPARγ (1:1,000; cat. no. ab178860; Abcam), C/EBP homologous protein (CHOP; 1:1,000; cat. no. 2895; Cell Signaling Technology, Inc.), glucose-regulated protein 78 (GRP78; 1:1,000; cat. no. ab21685; Abcam), phosphorylated (p)-protein kinase (PKR)-like ER kinase (p-PERK; 1:200; cat. no. orb504147; Biorbyt), PERK (1:500; cat. no. orb1294328; Biorbyt), p-inositol requiring enzyme 1α (p-IRE1α; 1:1,000; cat. no. ab243665; Abcam), IRE1α (1:1,000; cat. no. ab37073; Abcam) and GAPDH (1:2,500; cat. no. ab9485; Abcam) at 4˚C overnight. On the following day, the membranes were washed with Tris-buffered saline containing 0.1% Tween-20, and subsequently incubated with horseradish peroxidase-conjugated goat anti-rabbit (1:5,000; cat. no. ab6721; Abcam) or goat anti-mouse (1:5,000; cat. no. ab6789; Abcam) secondary antibodies at room temperature for 2 h. The signals were visualized using Amersham ECL Prime Western Blotting Detection Reagent (Amersham; Cytiva) and were semi-quantified by ImageJ software (Version 1.52; National Institutes of Health).

All data were expressed as mean ± standard deviation from at least three independent experiments. Data analysis was conducted using GraphPad Prism 8 (GraphPad Software; Dotmatics). One-way ANOVA analysis followed by the Tukey's post-hoc test was used to compare the differences among groups. P<0.05 was considered to indicate a statistically significant difference.

To investigate the role of PPARγ in AGE-induced VT, HUVECs were stimulated by M-HSA to mimic AGE-induced DVT and the expression levels of PPARγ were detected. As demonstrated in Fig. 1A, the protein expression levels of PPARγ were significantly reduced following M-HSA stimulation in HUVECs. Therefore, a gain-of function experiment was conducted to upregulate PPARγ (Fig. S1). The expression levels of PPARγ in the M-HSA + oe-PPARγ group were significantly higher than those in the M-HSA + oe-NC group (Fig. 1A). In addition, PPARγ activity was also weakened by M-HSA stimulation while it was increased following PPARγ overexpression (Fig. 1B). Subsequently, the data from the CCK-8 and TUNEL assays indicated that M-HSA led to a significant reduction in cell viability and an apparent elevation in TUNEL-positive cells, whereas these changes were inhibited when PPARγ was overexpressed (Fig. 1C-E), suggesting that PPARγ had the ability to alleviate M-HSA-induced cell viability loss and apoptosis in HUVECs. In addition, the downregulated 6-K-PGF1α levels and upregulated ET-1 levels in HUVECs, which were caused following M-HSA induction, were also partly abolished by PPARγ overexpression (Fig. 1F and G). These data suggested that PPARγ overexpression attenuated M-HSA-induced endothelial injury in HUVECs by improving cell viability, inhibiting cell apoptosis, upregulating 6-K-PGF1α levels and downregulating ET-1 levels.

Since oxidative stress and inflammation are common pathological processes responsible for vascular endothelial cell damage, the regulatory role of PPARγ was also investigated on inflammation and oxidative stress in M-HSA-stimulated HUVECs. As expected, M-HSA resulted in excessive production of TNF-α, IL-1β and IL-6 in HUVECs, while PPARγ overexpression was capable to suppress the overproduction of these markers (Fig. 2A-C). Furthermore, elevated levels of ROS and MDA and a reduced level of SOD were observed in HUVECs following M-HSA stimulation; these effects were partly reversed by PPARγ overexpression (Fig. 2D-F). The aforementioned data indicated a protective role of PPARγ against M-HSA-stimulated inflammation and oxidative stress in HUVECs.

Subsequent studies investigated the potential regulatory mechanism by which ERS is induced by a variety of physiological and pathological factors including oxidative stress. The protein expression levels of CHOP, GRP78, p-PERK and p-IRE1α were significantly increased following M-HSA stimulation, suggesting that M-HSA triggered the activation of ERS in HUVECs (Fig. 3). However, this activation was weakened by PPARγ overexpression, as demonstrated by the restoration of the protein expression changes following PPARγ overexpression in M-HSA-stimulated HUVECs.

To verify the critical role of ERS during the regulation of PPARγ in M-HSA-induced HUVECs, the agonist of ERS, TM, was used and the aforementioned cellular experiments were re-conducted. It was observed that the inhibitory effects of PPARγ on M-HSA-induced cell viability loss and cell apoptosis in HUVECs were weakened by TM (Fig. 4A-C). Moreover, additional treatment of TM caused a decrease in 6-K-PGF1α levels and an increase in ET-1 levels compared with the corresponding levels noted in the M-HSA + oe-PPARγ group (Fig. 4D and E). In addition, the protective effects of PPARγ against M-HSA-induced inflammation and oxidative stress were also weakened by TM in HUVECs (Fig. 5A-F). Therefore, these data suggested that the protective role of PPARγ against M-HSA-induced HUVEC injury could be diminished by TM, implying that PPARγ may exert its functions by inhibiting the activation of ERS.