Primary HUVECs were isolated from different six neonatal umbilical cords as previously described (26). Briefly, HUVECs at passages 2–4 were maintained in M199 medium (Invitrogen, Thermo Fisher Scientific, Inc.) supplemented with 20% fetal bovine serum (Hyclone, GE Healthcare Life Sciences) and 60 µg/ml endothelial cell growth supplement (BD Biosciences) at 37°C in a 5% CO₂ incubator and then exposed to the desired treatment in triplicate. The isolation procedure for HUVECs was approved by the Research Committee at the Third Affiliated Hospital of Sun Yat-sen University (approval nos. 2010-2–48 and 2018-02-057-01). The donors were negative for human immunodeficiency virus and hepatitis B virus and provided written informed consent to donate the umbilical cords.

To induce senescence, isolated HUVECs were treated with 60 µM H₂O₂ (Sigma-Aldrich, Merck KGaA) for 1 h and then cultured for another 24 h at 37°C. Rb1 (16071307, Chengdu Pufei De Biotech Co., Ltd.) used in the present study was extracted from Panax ginseng by HPLC according to the manufacturer's instructions and the purity of Rb1 used in the present study was 98.85%. To evaluate the effect of Rb1 on senescence, the cells were pretreated with 10 or 20 µM Rb1 for 30 min prior to H₂O₂ treatment. To measure the effect of the SIRT1 inhibitor nicotinamide (NAM; Sigma-Aldrich, Merck KGaA) and AMPK inhibitor compound C (Sigma-Aldrich, Merck KGaA), the cells were incubated with 20 mM NAM (27) or 8 µM compound C for 30 min as reported previously (18,28,29) and then treated with or without Rb1 at concentrations of 10 or 20 µM for 30 min before H₂O₂ treatment. At the end of each experiment, the cultured supernatants and monolayered cells were harvested for analyses.

NO production was evaluated by measuring the accumulation of nitrites. The Griess method (30) was used to detect NO using a NO assay kit (Beyotime Institute of Biotechnology), following the manufacturer's instructions. Briefly, after the cells were cultured and treated as described above, 50 µl culture supernatant was incubated with 50 µl Greiss reagent I and 50 µl Greiss reagent II in a 96-well microplate at room temperature for 30 min. The optical density was measured with a Victor microplate reader (PerkinElmer, Inc.) at 540 nm. Nitrite concentrations in the medium were calculated according to a standard curve.

Senescence was detected using a senescence-associated β-galactosidase (SA-β-gal)-positive approach according to a published protocol (31). After HUVECs were washed twice with prechilled PBS, the cells were fixed with 2% formaldehyde plus 1% glutaraldehyde for 5–10 min at room temperature. The cells were then washed twice with prechilled PBS for 3 min and stained with a staining solution [40 mmol/l citric acid/sodium phosphate buffer, 5 mmol/l potassium ferrocyanide (K₄[Fe(CN)₆]3H₂O), 5 mmol/l potassium ferricyanide (K₃[Fe(CN)₆]), 150 mmol/l sodium chloride, 2 mmol/l magnesium chloride and 1 mg/ml X-gal] overnight at 37°C without CO₂. Senescent cells were identified as blue-stained cells under a TS100 inverted microscope (Nikon Corporation) at ×100 magnification. At least 400 cells were examined to determine the percentage of SA-β-gal-positive cells in each group.

The cellular NADP⁺/NADPH ratio was determined using a NAD⁺/NADH Quantification Kit (Beyotime Institute of Biotechnology), according to the manufacturer's instructions. In brief, HUVECs (1×10⁶/well) were seeded in six-well plates and exposed to the experimental conditions. To measure the NADP⁺/NADPH ratio, the cells were harvested, lysed with 200 µl NAD⁺/NADH buffer and gently pipetted to promote cell lysis. Then, 50–100 µl of the samples were collected and incubated for 30 min at 60°C. Then, 20 µl of the reacted samples were added to a 96-well plate and analyzed at 450 nm as the reference wavelength in the Victor microplate reader. The NAD⁺/NADH ratio was calculated according to a standard curve and normalized to the cell number, following the manufacturer's protocol.

Determination of intracellular ATP in the indicated groups of HUVECs was performed by a bioluminescence assay (ATP Assay kit; Beyotime Institute of Biotechnology), according to the manufacturer's instructions. Briefly, the cells were washed twice with prechilled PBS and then lysed in lysis buffer on ice. Then, the samples were harvested and centrifuged at 12,000 × g for 5 min at 4°C and the supernatants were collected for subsequent analysis. After the reaction solutions containing luciferase and luciferin were added and background luminescence was measured, the ATP standard solution and samples were added and luminescence was measured. Then, the background luminescence was subtracted and the standard curve was constructed. The ATP concentrations were calculated from the standard curve and normalized to total protein content.

HUVECs were grown in six-well plates. After the indicated treatments, the cells were washed twice with prechilled PBS and then lysed in radioimmunoprecipitation buffer with a protease inhibitor cocktail, phenylmethylsulfonyl fluoride and sodium orthovanadate (Santa Cruz Biotechnology, Inc.). The protein concentration was measured by the Bradford method. Proteins (30 µg) in 30 µl reducing sample buffer were boiled for 5 min at 100°C and then resolved by SDS-PAGE (8 or 12% gels) for 2 h at 100 V. The proteins were transferred onto a polyvinylidene difluoride membrane for 90 min at 100 V. After transfer, the membrane was incubated in 25 ml blocking buffer [1X Tris-buffered saline (TBS) and 0.1% Tween-20 with 5% non-fat dry milk] for 1 h at room temperature. The primary antibodies were rabbit polyclonal anti-eNOS (1:500; Cell Signaling Technology, Inc.), Rabbit polyclonal anti-PAI-1 (1:500; Cell Signaling Technology, Inc.), Rabbit polyclonal anti-AMPK (1:2,000; Cell Signaling Technology, Inc.), rabbit polyclonal anti-phosphorylated AMPK (Thr172; 1:1,000; Cell Signaling Technology, Inc.), rabbit polyclonal anti-SIRT1 (1:1,000; Cell Signaling Technology, Inc.), and rabbit polyclonal anti-GAPDH (control, 1:5,000; Cell Signaling Technology, Inc.). The membrane was incubated with the primary antibody in 10 ml primary antibody dilution buffer with gentle agitation overnight at 4°C. After washing three times for 10 min each with 15 ml of 10X TBS/0.1% TBST, the membrane was incubated with a horseradish peroxidase-conjugated secondary antibody (1:3,000; cat. no. 7074; Cell Signaling Technology, Inc.) in 10 ml blocking buffer with gentle agitation for 1 h at room temperature, followed by 3 washes for 10 min each. Membranes were developed using the enhanced chemiluminescence detection method (EMD Millipore). The signals were quantified using Quantity One software (version 4.6.9; Bio-Rad Laboratories, Inc.).

Results were obtained from at least three independent experiments (n≥3) if not otherwise stated. Data are expressed as the mean ± standard deviation (n≥3). Statistical significance was calculated by one-way or two-way analysis of variance with Bonferroni's post-hoc test using GraphPad Prism software (version 5.0; GraphPad Software, Inc.). P<0.05 was considered to indicate a statistically significant difference.

A number of studies have reported that H₂O₂ is a major inducer of cell senescence and oxidative stress in HUVECs and other different cell lines (32–35), which plays a central role in cell dysfunction. H₂O₂ was used at a concentration of 60 µM in the present study based on our previous studies (16,36,37) in which 60 µM H₂O₂ was reported to induce cell senescence and oxidative stress sufficiently. Senescence-associated β-galactosidase (SA-β-gal) staining was used to determine the degree of cell senescence. Aging cells stained blue indicated the senescent phenotype of HUVECs. In line with our previous studies (16,36,37), it was demonstrated that 60 µM H₂O₂ increased the SA-β-gal⁺ cell number (blue staining). In addition, the H₂O₂ treated group had more cells with a clear outline, enlarged cell body and difficulty in adhering to the bottom of the culture plate, all characteristics of stressed senescent cells (38,39). By contrast, pretreatment with Rb1 decreased the SA-β-gal⁺ cell number and attenuated cell senescence induced by H₂O₂ in a dose-dependent manner (Fig. 1A and B). PAI-1 and eNOS expression were then measured after the same treatment. The results demonstrated that 60 µM H₂O₂ increased PAI-1 expression (Fig. 1C) and decreased eNOS expression (Fig. 1D) in HUVECs, which were restored by Rb1 in a dose-dependent manner.

A number of studies have shown that SIRT1 exhibits anti-inflammatory (40–42) and antioxidant effects (43–45) in the endothelium. The effect of Rb1 on SIRT1 and phosphorylation of the catalytic subunit of AMPK (Thr172) in the presence or absence of H₂O₂ was examined. The results demonstrated that 60 µM H₂O₂ inhibited SIRT1 expression (Fig. 2A and B) as well as phosphorylation of AMPK (Fig. 2A and C) and that treatment with Rb1 at 10 and 20 µM restored Sirt-1 expression (Fig. 2A and B) and AMPK phosphorylation (Fig. 2A and C) in a dose-dependent manner.

The intracellular NAD⁺/NADH ratio in the presence or absence of Rb1 was further examined and it was identified that treatment with 60 µM H₂O₂ reduced the NAD⁺/NADH ratio, which was restored with Rb1 pretreatment (Fig. 2D). The data confirmed that H₂O₂-induced decrease in the NAD⁺/NADH ratio were associated with reduced SIRT1 expression and it was demonstrated for the first time, to the best of the authors' knowledge, that treatment with Rb1 at 10 and 20 µM Rb1 prevented the reductions in SIRT1 and the NAD⁺/NADH ratio, and inhibited phosphorylated AMPK, in HUVECs exposed to H₂O₂.

Next, it was determined whether AMPK was involved in the inhibitory effects of Rb1 on the H₂O₂-induced oxidative response. HUVECs were treated with 10 or 20 µM Rb1 for 24 h in the presence or absence of compound. C, a specific inhibitor of AMPK. As shown in Fig. 3A, compound C clearly downregulated the phosphorylation of AMPK at 6, 8 and 10 µM in a dose-dependent manner. According to the results, 8 µM compound C was chosen to inhibit the phosphorylation of AMPK in the following experiment. Pretreatment of HUVECs with compound C significantly abolished the inhibitory effects of Rb1 on H₂O₂-induced PAI-1 expression (Fig. 3C). Rb1 also significantly restored eNOS expression (Fig. 3D) and NO production (Fig. 3E) and this beneficial effect was markedly reversed by pretreatment of HUVECs with compound C (Fig. 3). The results suggest that AMPK activation is essential for the inhibitory effect of Rb1 on H₂O₂-induced senescence and the oxidative response in HUVECs.

To further understand the role of SIRT1 in the AMPK pathway, it was investigated whether inactivation of SIRT1 by nicotinamide affected H₂O₂-induced cell injury. As Guo et al (27) reported that 20 mM NAM decreases SIRT1 gene expression significantly, HUVECs were treated with H₂O₂ in the presence or absence of 20 mM NAM, followed by observation of the effects on H₂O₂-induced senescence and the expression of antioxidant genes. As shown in Fig. 4, NAM reduced the phosphorylation of AMPK (Fig. 4A and B), indicating that AMPK is the downstream protein modulated by SIRT1.

As AMPK is one of the most important proteins in modulating the cellular energy metabolism and ATP is a major downstream product of mitochondrial energy coordinators, the effects of NAM with or without Rb1 on the ATP level were next investigated. Rb1 protected HUVECs from H₂O₂-induced PAI-1 expression (Fig. 4C) and rescued the downregulation of eNOS expression (Fig. 4D) and NO production (Fig. 4E). As shown in Fig. 4F, treatment with Rb1 increased the ATP level under H₂O₂-induced oxidative stress. However, the promotional effect on the ATP level by Rb1 was not observed in the NAM-treated group (Fig. 4F). The beneficial effects of Rb1 on H₂O₂-induced injuries were reversed by treatment with NAM (Fig. 4). These in vitro data confirmed that inhibition of SIRT1 by NAM blocks Rb1 to increase phosphorylation of AMPK, ATP production, eNOS expression and NO production, which collectively indicate that Rb1 plays a protective role against H₂O₂-induced injury of HUVEC via the SIRT1/AMPK pathway.