Human renal mesangial cells (HRMCs) were purchased from the Cell Bank of Type Culture Collection, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences (Shanghai, China). The human podocyte cells (HPCs) were obtained from Celprogen Inc., (Torrance, CA, USA; cat. no. 36036-08). HPCs and HRMCs were treated with standard glucose medium (5.5 mmol/l, Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) and high glucose medium (25 mmol/l, Gibco; Thermo Fisher Scientific, Inc.) to generate the control and hyperglycemic groups, respectively, for 12 h to induce hyperglycemic damage, as described previously (23). In the present study, SCH772984 (Selleck Chemicals, Houston, TX, USA; cat. no. S7101) was added into HRMC to inhibit the activity of ERK for 2 h. Liraglutide (0–20 nM, Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) was used to treat HRMC and HPC for 12 h at 37°C in an atmosphere containing 5% CO₂.

The cellular ROS production was measured via immunofluorescence using Dihydrorhodamine 123 (Molecular Probes, USA, Cat. no. D23806). Briefly, cells were incubated with Dihydrorhodamine 123 for 30 min in the dark at 37°C in an atmosphere containing 5% CO₂. Subsequently, PBS was used to wash the cells, to remove the free probe. Finally, cells were observed using a fluorescent microscope (BX51; Olympus Corporation, Tokyo, Japan; magnification, ×200). The fluorescence intensity was recorded as the ROS production and analyzed using Image-Pro Plus version 6.0 (Media Cybernetics, Rockville, MD, USA), as described previously (24).

The mitochondrial potential was measured via an immunofluorescence assay using JC-1 staining. First, cells were washed with PBS three times, and then the culture medium was replaced with PBS. Subsequently, the JC-1 probe Mitochondrial membrane potential assay kit with JC-1 (Beyotime Institute of Biotechnology, Haimen, China; cat. no., C2006) was added into the medium for 30 min in the dark. Then, samples were centrifuged at 10,000 × g for 20 min at 4°C to acquire pellets and the cell supernatant was discarded, and PBS was used to wash the cells at least three times at room temperature. Finally, the cell nuclei were stained using DAPI for 3 min in the dark. Then, the samples were observed using a fluorescence microscope (BX51; Olympus Corporation, magnification, ×200). The fluorescence intensity was recorded and analyzed using Image-Pro Plus 6.0 (Media Cybernetics, Rockville, MD, USA). To examine ATP production, an ATP Production Assay Kit Enhanced ATP Assay kit (Beyotime Institute of Biotechnology; cat. no., S0027) was used according to the manufacturer's protocol, as described previously (25).

Mitochondrial function was measured via mPTP opening analysis using a calcein-AM/cobalt method, as previously described (26). Cells were cultured in 96-well plates (1×10⁶) and 5 µM Calcein-AM was added to the medium for 30 min at 37°C in the dark. Following washing 3 times with PBS, the samples were observed using a Varioskan^® Flash microplate reader (Thermo Fisher Scientific, Inc., Waltham, MA, USA) with an excitation wavelength of 488 nm and an emission wavelength of 525 nm. In the EdU assay, cells (1×10⁴) were cultured in 24-well plates. Then, the EdU Proliferation kit (Abcam, Cambridge, MA, USA; cat. no., ab219801) was used to observe the cellular proliferation. Cell nuclei were labelled by DAPI (Guangzhou RiboBio Co., Ltd., Guangzhou, China), and the EdU-positive cells were observed by laser scanning microscope (magnification, ×200). The fluorescence intensity was recorded and analyzed using Image-Pro Plus version 6.0 (Media Cybernetics, Rockville, MD, USA) (27).

LDH release is a hallmark of cell death. Accordingly, the LDH concentration, as an indicator of the level of cell death, was measured in the medium. The LDH content in the cell medium was measured via an LDH Release Commercial kit (Beyotime Institute of Biotechnology). The concentration of antioxidants was measured via ELISA. The ELISA kits for glutathione (GSH; Thermo Fisher Scientific, Inc.; cat no., T10095), GSH peroxidase (GPx; Beyotime Institute of Biotechnology; cat no., S0056) and superoxide dismutase (SOD; Thermo Fisher Scientific, Inc.; cat. no., BMS222TEN) were used according to the manufacturer's instructions as described previously (28).

Cellular apoptosis was measured via TUNEL staining and MTT assays. For TUNEL staining, cells were fixed with 3.7% paraformaldehyde at 4°C for 30 min. Then, a TUNEL staining kit (Roche Applied Science, Indianapolis, IN, USA) was used to tag apoptotic cells. Following staining of the cells with DAPI (5 mg/ml) for 30 min in the dark at 37°C in an atmosphere containing 5% CO₂, cells were observed using a fluorescent microscope (BX51; Olympus Corporation; magnification, ×200). In addition, cellular viability was also measured via MTT assays. Cells (1×10⁴) were seeded onto a 96-well plate. Then, MTT solution (2 mg/ml; Beyotime Institute of Biotechnology) was added to the medium for 4 h in the dark. Subsequently, dimethyl sulfoxide was used to dissolve the MTT solution, and the OD of the cells was recorded at an absorbance of 490 nm using a spectrophotometer (Epoch 2; BioTek Instruments, Inc., Winooski, VT, USA). Caspase-3 and caspase-9 activity levels were measured using the Caspase 3 Activity Assay kit (Beyotime Institute of Biotechnology; cat. no., C1115) and Caspase 9 Activity Assay kit (Beyotime Institute of Biotechnology; cat. no, C1158) were measured according to the manufacturer's instructions (29).

For the immunofluorescence assay, cells were fixed with 3.7% paraformaldehyde at 4°C for 30 min, followed by permeabilization using 0.1% Triton X-100 for 30 min at room temperature. Subsequently, samples were washed with PBS and then blocked with 15% non-fat dried milk in TBS containing 0.05% Tween-20 for 45 min at room temperature. Subsequently, samples were treated with primary antibodies overnight at 4°C. Following extensive washing with PBS, samples were treated with secondary antibodies (Alexa Fluor^® 488 donkey anti-rabbit antibody; 1:1,000; cat. no. A-21206 and Alexa Fluor^® Plus 647 goat anti-rabbit antibody; 1:1,000; cat. no. A-32733; Invitrogen; Thermo Fisher Scientific, Inc.) for 45 min at room temperature. Following loading with DAPI (5 mg/ml) for 30 min in the dark at 37°C in an atmosphere containing 5% CO₂, cells were observed using an inverted fluorescence microscope (BX51; Olympus Corporation; magnification, ×200). The primary antibodies used in the present study were as follows: Phosphorylated (p)-ERK (1:1,000; Abcam; cat. no., ab176660), Sirt3 (1:1,000; Abcam; cat. no., ab86671) and cytochrome-c (Cyt-c; 1:1,000; Abcam; cat. no., ab90529). The fluorescence intensity was recorded and analyzed using Image-Pro Plus 6.0 (Media Cybernetics, Rockville, MD, USA).

In the present study, cells were lysed in radioimmunoprecipitation assay lysis buffer (Beyotime Institute of Biotechnology). Total protein was analyzed using the bicinchoninic acid assay (Beyotime Institute of Biotechnology) and 60 µg of protein was loaded onto a 10% SDS-PAGE gel and then transferred onto a polyvinylidene fluoride membrane (Roche Applied Science, Penzberg, Germany). The membranes were subsequently blocked with 5% non-fat milk for 1 h at room temperature prior to incubation with the primary antibodies at 4°C overnight. Bands were detected via enhanced chemiluminescence reagents (Applygen Technologies, Inc., Beijing, China). The primary antibodies used in the present study were as follows: p-ERK (1:1;000; Abcam; cat. no., ab65142), total-ERK (t-ERK, 1:1,000; Abcam; cat. no., ab54230), B-cell lymphoma 2 (Bcl-2; 1:1,000; Cell Signaling Technology, Inc., Danvers, MA, USA; cat. no., 3498), Bcl-2-assocaited X protein (Bax; 1:1,000; Abcam; cat. no., ab32503), Cyclin D1 (1:1,000; Abcam; cat. no., ab134175), Cyclin E (1:1,000; Abcam; cat. no., ab171535), cellular inhibitor of apoptosis protein 1 (c-IAP; 1:1,000; Cell Signaling Technology, Inc.; cat. no., 4952), Bcl-2-associated death promotor (Bad; 1:1,000; Abcam; cat. no., ab90435), caspase-9 (1:1,000; Cell Signaling Technology, Inc.; cat. no., 9504), tumor necrosis factor α (TNF-α; 1:1,000; Abcam; cat. no., ab6671), matrix metalloproteinase 9 (MMP-9; 1:1,000; Abcam; cat. no., ab38898), interleukin-8 (IL-8; 1:1,000; Abcam; cat. no., ab7747), Yap (1:1,000; Cell Signaling Technology, Inc.; cat. no., 14074), Sirt3 (1:1,000; Abcam; cat. no., ab86671), pro-caspase-3 (1:1,000; Cell Signaling Technology, Inc.; cat. no., 9662) and cleaved caspase-3 (1:1,000; Cell Signaling Technology, Inc.; cat. no., 9664). The secondary antibodies used in the present study were: Horseradish peroxidase (HRP)-coupled secondary antibodies (1:2,000; cat. nos. 7074 and 7076; Cell Signaling Technology, Inc.). Bands were detected using an enhanced chemiluminescence substrate (Applygen Technologies, Inc., Beijing, China). GAPDH (1:1,000; cat. no. 5174; Cell Signaling Technology, Inc.) and β-actin (1:1,000; cat. no. 4970; Cell Signaling Technology, Inc.) were used as internal controls. The blots were analyzed using Quantity One 4.6 software (Bio-Rad Laboratories, Inc., Hercules, CA, USA).

To inhibit Sirt3 expression, siRNA was used to knock down its expression as described previously (30). The siRNA against Sirt3 was obtained from Yangzhou Ruibo Biotech Co., Ltd. (Yangzhou, China). The siRNA (70 nM/well) was transfected into cells using Lipofectamine^® 2000 transfection reagent (Invitrogen; Thermo Fisher Scientific, Inc.). After 72 h of transfection, cells were collected, and the expression of Sirt3 was determined via western blot analysis. The Sirt3 siRNA sequence was: 5′-ACUCCCAUUCUUCUUUCAC-3′.

In the present study, at least 3 independent experiments were performed and all data are presented as the mean ± standard error. The data were analyzed by one-way analysis of variance followed by a Tukey's Honestly Significant Difference post-hoc test. P<0.05 was considered to indicate a statistically significant difference.

To determine the protective role of liraglutide in hyperglycemia-mediated HRMC damage, different doses of liraglutide were added into the medium of HRMCs. Then, the cellular viability was measured via MTT assay. As demonstrated in Fig. 1A, compared with the control group, hyperglycemia significantly decreased HRMC viability, and this effect was reversed by liraglutide in a dose-dependent manner. Similarly, TUNEL assays also indicated that hyperglycemia increased the number of TUNEL-positive cells compared with the control group (Fig. 1B and C). However, liraglutide treatment dose-dependently decreased the ratio of TUNEL-positive cells (Fig. 1B and C). This information suggested that liraglutide protected the renal mesangial cells against hyperglycemia-mediated damage. Similar results were also observed using HPCs (Fig. 1D and E). These results indicated that liraglutide protected HRMC and HPC viability in the presence of hyperglycemia stress. Notably, as the minimal effective concentration of liraglutide on HRMC and HPC viability in a hyperglycemic state was 5 nM (Fig. 1A-E), liraglutide was used at 5 nM in subsequent experiments.

Subsequently, EdU assays were used to observe the proliferative capacity of the renal mesangial cells with liraglutide treatment. The number of EdU-positive cells was decreased during hyperglycemic stress and was reversed to almost normal levels with liraglutide treatment (Fig. 1F and G). Similar results were also observed using HPCs (Fig. 1H and I). Cellular proliferation is primarily regulated by Cyclin proteins, in particular Cyclin D1 and Cyclin E. Notably, hyperglycemia-inhibited Cyclin D1 and Cyclin E expression levels were markedly upregulated by liraglutide (Fig. 1J and L). Altogether, these data demonstrated that liraglutide decreased renal mesangial cell damage and promoted cell proliferation in the context of the hyperglycemic challenge.

Oxidative stress and the inflammatory response have been acknowledged as the primary pathophysiological events involved in diabetic renal damage. Therefore, cellular ROS were measured via immunofluorescence in HRMCs. As demonstrated in Fig. 2A and B, hyperglycemia treatment elevated ROS production, and this effect was reversed by liraglutide. As a consequence of ROS overproduction, the contents of antioxidants, including SOD, GPX and GSH, were correspondingly decreased in the hyperglycemia-treated cells (Fig. 2C-E). However, liraglutide treatment reversed the concentrations of antioxidants. These data indicated that hyperglycemia-mediated oxidative stress was repressed by liraglutide. In addition, the effect on the inflammatory response with liraglutide treatment was also observed. Compared with the control group, hyperglycemia increased the expression of MMP-9, IL-8 and TNF-α (Fig. 2F-I); these effects were abrogated by liraglutide treatment. Altogether, this information suggested that hyperglycemia promoted oxidative stress and inflammatory damage in HRMC and that this effect was alleviated by liraglutide.

To explain the beneficial action of liraglutide on diabetic renal damage, its anti-apoptotic effect was examined, in particular mitochondrial apoptosis. The early molecular characterization of mitochondrial apoptosis involves decreased mitochondrial potential (31). The present study identified that hyperglycemia decreased the mitochondrial potential and that this effect was reversed by liraglutide (Fig. 3A and B). As a consequence of decreases in mitochondrial potential, the permeability of the mitochondria is increased, leading to mPTP opening (32). By analyzing the rate of mPTP opening, it was demonstrated that hyperglycemia-mediated mPTP opening was inhibited by liraglutide (Fig. 3C). Excessive mPTP opening may irreversibly facilitate mitochondrial pro-apoptotic factor release into the nucleus (33). As indicated in Fig. 3D, hyperglycemia promoted more Cyt-c leakage from mitochondria into the nucleus; this effect was inhibited by liraglutide treatment. Upon release from mitochondria into the cytoplasm/nucleus, Cyt-c interacts with caspase-9 to activate the apoptosome, which then additionally activates the caspase-3-mediated cellular apoptotic system (34). In the present study, it was identified that hyperglycemia increased the expression of caspase-3, caspase-9, Bax, and Bad (Fig. 3E-K). However, the anti-apoptotic proteins Bcl-2 and c-IAP were downregulated by hyperglycemia. These data indicated that hyperglycemia triggered mitochondrial apoptosis in HRMCs. Notably, liraglutide administration markedly inhibited the activation of pro-apoptotic proteins and reversed the expression of anti-apoptotic factors (Fig. 3E-K), effectively correcting hyperglycemia-mediated mitochondrial apoptosis.

The following experiments were performed to determine the mechanism by which liraglutide protected mitochondrial function within a high glucose stress environment. A previous study suggested that Sirt3 was vital for mitochondrial homeostasis (35). Accordingly, the present study assessed whether liraglutide modulated diabetic renal damage via Sirt3. Western blot analysis data in Fig. 4A and B demonstrated that Sirt3 expression was downregulated in response to hyperglycemia and was upregulated with liraglutide treatment. Subsequently, to verify whether Sirt3 was necessary for the anti-apoptotic properties of liraglutide on mitochondrial homeostasis, siRNA was used to knock down Sirt3 expression. The knockdown efficiency was confirmed via western blot analysis (Fig. 4C and D) and immunofluorescence (Fig. 4E and F). Subsequently, mitochondrial function was examined again in the Sirt3-silenced cells. Mitochondrial membrane potential was decreased with hyperglycemia stress and was reversed to almost normal levels following liraglutide treatment (Fig. 4G and H). However, deletion of Sirt3 abrogated the protective action of liraglutide on mitochondrial membrane potential (Fig. 4G and H). In addition, hyperglycemia-mediated ROS overproduction (Fig. 4I-J) and antioxidant downregulation (Fig. 4K and M) were also reversed by liraglutide in a Sirt3-dependent manner. With regard to mitochondrial apoptosis, caspase-9 (Fig. 4N) and caspase-3 activities (Fig. 4O) were increased in response to hyperglycemia stress and was decreased in liraglutide-treated cells. However, Sirt3 deficiency abrogated the inhibitory effect of liraglutide on caspase-9 and caspase-3 activation. Altogether, these data demonstrated that liraglutide maintained mitochondrial homeostasis in HRMC via Sirt3.

Experiments were then performed to determine the upstream regulatory factors for Sirt3 upregulation in response to liraglutide treatment. Previous studies have identified the ERK-Yap axis as a novel signaling pathway that modifies mitochondrial function (36–38). Therefore, the present study investigated whether the ERK-Yap signaling pathway was involved in liraglutide-mediated Sirt3 activation. Western blot analysis data in Fig. 5A-D suggested that p-ERK was downregulated in the hyperglycemia-treated cells, which was also accompanied with a decreased Yap expression. Notably, liraglutide treatment reversed the expression of p-ERK and Yap. Subsequently, to confirm whether the ERK-Yap signaling pathway was the upstream mediator of Sirt3, a pathway blocker was used. Following inhibition of ERK activity using SCH77298, p-ERK and Yap expression levels were downregulated in liraglutide-treated cells (Fig. 5A-D). In addition, the positive effect of liraglutide on Sirt3 activation was also nullified by SCH772984 (Fig. 5A-D). These data suggested that liraglutide modulated Sirt3 expression via the ERK-Yap signaling pathway. This conclusion was additionally verified with immunofluorescence via p-ERK and Sirt3 co-staining (Fig. 5E-G). Activated p-ERK expression was positively associated with increased Sirt3 expression in liraglutide-treated cells compared with the control group (Fig. 5E-G). In comparison, ERK inhibition promoted Sirt3 downregulation despite treatment with liraglutide (Fig. 5E-G).

Finally, TUNEL assays were performed to investigate whether the ERK-Yap signaling pathway was involved in hyperglycemia-mediated HRMC apoptosis. Similar to the aforementioned data, hyperglycemia-triggered apoptosis was markedly inhibited by liraglutide (Fig. 5H-I). However, inhibition of ERK negated the anti-apoptotic effect of liraglutide on HRMCs (Fig. 5H-I). Furthermore, caspase-3 activity was increased in response to hyperglycemic stress and was decreased to almost normal levels with liraglutide treatment (Fig. 5J). However, inhibition of ERK abrogated the beneficial effects of liraglutide on HRMCs survival (Fig. 5J). Taken together, these results confirmed that ERK-Yap axis was activated by liraglutide and protected HRMCs against hyperglycemia-mediated damage by sustaining Sirt3 (Fig. 6).