The present study was performed in accordance with the Declaration of Helsinki and the guidelines of the Ethics Committee of No. 1 People's Hospital (Suzhou, China). All experimental protocols were approved by the Ethics Committee of No.1 People's Hospital. The experiments on animals were performed in accordance with the Guide to the Care and Use of Experimental Animals (Vol. 1, 2nd ed., 1993; and Vol. 2, 1984; Canadian Council on Animal Care, Ottawa, ON, Canada; www.ccac.ca), or the Guide for the Care and Use of Laboratory Animals (1996; National Academy Press, Washington, DC, USA).

Male Sprague-Dawley rats (n=150, 8–10 weeks, weight, 250±10 g) were purchased from the Experimental Animal Center, Academy of Military Medical Science and were housed under standard laboratory conditions (27°C, 40–60% humidity, a 12-h light and dark cycle). A commercial pellet diet and fresh drinking water were given ad libitum. Rats were randomly divided into the following groups, with n=6 in each: i) Sham group; ii) PBS; iii) low dose of liraglutide; and iv) high dose of liraglutide. Rats were intraperitoneally anaesthetized with sodium pentobarbital (30 mg/kg). The animals were subsequently incubated and ventilated with a volume-regulated respirator during surgery. Subsequent to a left lateral thoracotomy and pericardectomy, the left coronary artery was identified and ligated with a 6.0 prolene suture. Freshly prepared liraglutide (Sigma-Aldrich; Merck KGaA, Darmstadt, Germany) was administered via the caudal vein at doses of 0.09 mg/kg (low dose group) or 0.18 mg/kg (high dose group) for 28 successive days post-infarction. These concentrations of drugs were selected as they were previously reported to be effective in reducing cardiac damage (10).

The hearts were excised and rapidly frozen in Optimal Cutting Temperature medium at −20°C (Agar Scientific, Ltd., Stansted, UK) for the preparation of frozen sections (4-µm thickness). A total of 10 sections were prepared at 10 different transversal levels at the site of tissue necrosis, equally distributed from base to apex. Sirius red staining was performed at room temperature in sections to quantify the cardiac fibrosis and was observed using an inverted microscope (magnification, ×40; BX51; Olympus Corp., Tokyo, Japan). The degrees of collagen fiber accumulation in the infarcted area were evaluated by measuring the fluorescence density of fibrotic region in the left ventricular area, which was calculated using RS Image Pro, version 4.5 (Media Cybernetics, Inc., Rockville, MD, USA). Terminal deoxynucleotidyl-transferase-mediated dUTP nick end labeling (TUNEL) staining was performed to assess cellular apoptosis according to the protocol described below. The degree of apoptosis was calculated as the number of TUNEL-positive cells per 500 cellular nuclei. The nuclei were stained with the chromatin dye DAPI. Briefly, cells were fixed for 1 h in 4% (w/v) paraformaldehyde at room temperature. Following labeling with TUNEL for ~30 min at room temperature, the cells were exposed to DAPI in the dark for 5 min. Then, the stained cells were observed using an inverted microscope (magnification, ×100; BX51; Olympus Corp.). In order to determine the inflammatory response, heart sections were stained with anti-tumor necrosis factor (TNF) ^α (1:1,000; Cell Signaling Technology, Inc., Danvers, MA, USA; cat. no. 6945). In brief, samples were fixed with 4% paraformaldehyde for 10 min at room temperature, permeabilized with 0.3% Triton X-100 for 5 min, and blocked with 10% goat serum albumin (Invitrogen; Thermo Fisher Scientific, Inc., Waltham, MA, USA) for 1 h at room temperature. Specimens were subsequently incubated with transforming growth factor (TGF) ^β (1:1,000; Cell Signaling Technology, Inc.; cat. no. 3711) overnight at 4°C, then washed with PBS three times, and incubated with horseradish peroxidase AffiniPure Goat Anti-Rabbit secondary antibody (1:2,000; Beyotime Institute of Biotechnology, Haimen, China; cat. no. A0277) for 45 min at room temperature. Images were captured using an inverted microscope (magnification, ×40; BX51; Olympus Corp.).

In vitro, chronic hypoxia of cardiomyocytes was used to mimic the infarcted heart following AMI. Hypoxic conditions were produced using fresh Hank's solution (Beyotime Institute of Biotechnology) with 95% N₂ and 5% CO₂. The dishes (1×10⁶ cells) were placed into a hypoxic incubator at 37°C, that was equilibrated with 95% N₂ and 5% CO₂ and the actual oxygen concentration was 0. Ambient O₂ levels in the hypoxic incubator were monitored with an O2 analyzer (series-2000; Alpha Omega Instruments, Lincoln, RI, USA). Chronic hypoxia of cardiomyocytes cell line H9C2 (purchased from the American Type Culture Collection, Manassas, VA, USA) was performed for ~48 h. For liraglutide treatment, H9C2 were treated with liraglutide (0–50 nM) for 48 h in the presence of hypoxia. To activate SIRT1, pretreatment with SRT1720 (SRT; 10 µM; Sigma-Aldrich; Merck KGaA) was performed for ~4 h. To inhibit the SIRT1, Selisistat (10 µM; Sigma-Aldrich; Merck KGaA) was used for 6 h. To suppress the role of liraglutide, Exendin 9–39 (Ex9-39; 10 nM; Sigma-Aldrich; Merck KGaA) was used to block the glucagon-like peptide 1 receptor (GLP1R) under hypoxic conditions.

The 1×10⁶ cells were seeded into the dishes then were washed in PBS and permeabilized for 10 min at 4°C in a solution of 0.1% Triton X-100 and 0.1% sodium citrate in PBS. Then, samples were blocked with 10% goat serum albumin for 1 h at room temperature and subsequently were incubated with primary antibodies against overnight at 4°C (30). Following three rinses with PBS, the secondary antibody, Alexa Fluor 488 donkey anti-rabbit antibody (1:1,000; cat. no. A-21,206; Invitrogen; Thermo Fisher Scientific, Inc.) was added to the samples for 1 h at room temperature. The primary antibodies used in the present study were as follows: Mitochondrial import receptor subunit TOM20 homolog (1:1,000; Abcam, Cambridge, UK; cat. no. ab78547), lysosome-associated membrane glycoprotein 1 (1:1,000; Abcam; cat. no. ab24170), cytochrome-c (1:1,000; cyt-c; Abcam; cat. no. ab133504), Parkin (1:1,000; Cell Signaling Technology, Inc.; cat. no. 2132) and Sirt1 (1:1,000; Cell Signaling Technology, Inc.; cat. no. 9475). Images were taken using an inverted microscope (magnification, ×40; BX51; Olympus Corp.).

Cells were washed with PBS and lysed in Laemmli Sample Buffer (Bio-Rad Laboratories, Inc., Hercules, CA, USA), and further homogenized with a rotor-stator homogenizer. Proteins were isolated and concentrations were determined using the Bicinchoninic Acid Protein Assay kit (Thermo Fisher Scientific, Inc.) (31). A mass of 20–80 µg proteins was loaded on a 12–15% SDS-PAGE gel. Following electrophoresis, proteins were transferred to a polyvinylidene fluoride western blotting membrane (Roche Applied Science, Penzberg, Germany). Membranes were blocked with 5% nonfat dried milk [in TBS-Tween 20 (TBST)] for 2 h at room temperature and incubated overnight at 4°C with primary antibodies. The membrane was subsequently washed with TBST (5 min; three times) and incubated with horseradish peroxidase-conjugated secondary antibodies (Cell Signaling Technology, Inc.) for 1 h at room temperature. Following washing with TBST (5 min; three times), bands were detected using an enhanced chemiluminescence substrate (Applygen Technologies, Inc., Beijing, China). Band intensities were normalized to the respective internal standard signal intensity (^β-actin antibody; 1:2,000; Abcam; cat. no. ab8224). The experiment was repeated three times. The primaries antibodies used in the present study were against the following proteins: Pro-caspase3 (1:1,000; Cell Signaling Technology, Inc.; cat. no. 9662), cleaved caspase3 (1:1,000; Cell Signaling Technology, Inc.; cat. no. 9664), baculoviral IAP repeat-containing protein 2 (c-IAP1; 1:2,000; Cell Signaling Technology, Inc. cat. no. 7065), caspase9 (1:1,000; Abcam; cat. no. ab32539), Parkin (1:1,000; Cell Signaling Technology, Inc.; cat. no. 2132), microtubule-associated protein light chain (LC) 3II (1:1,000; Cell Signaling Technology, Inc.; cat. no. 3868), sequestome-1 (p62; 1:1,000; Abcam; cat. no. ab56416), Beclin1 (1:1,000; Cell Signaling Technology, Inc.; cat. no. 3495), autophagy protein 5 (Atg5; 1:1,000; Cell Signaling Technology, Inc.; cat. no. 12,994), Sirt1 (1:1,000; Cell Signaling Technology, Inc., cat. no. 9475), TGF^β (1:1,000; Cell Signaling Technology, Inc.; cat. no. 3711), matrix metalloproteinase (MMP) 9 (1:1,000; Cell Signaling Technology, Inc.; cat. no. 13667) and poly (ADP ribose) polymerase 1 (PARP; 1:1,000; Cell Signaling Technology, Inc.; cat. no. 9532) The mean densities of the bands were represented as the optical density in units/mm² and normalized to that of ^β-actin (Quantity One, version 4.6.2; Bio-Rad Laboratories, Inc.).

Cells were incubated in serum-free Dulbecco's modified Eagle's medium (DMEM; Gibco; Thermo Fisher Scientific, Inc.) containing dichloro-dihydro-fluorescein diacetate (10 µM) for 30 min (32). Cells were subsequently washed with PBS. Flow cytometric analyses were performed using a BD FACSCalibur™ flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA) using BD CellQuest Pro software (version 5.1; BD Biosciences).

The mitochondrial potential was assessed using the probe JC-1, a sensitive fluorescent dye used to detect alterations in mitochondrial potential. Following treatment, cells were incubated with 10 mg/ml JC-1 for 10 min at 37°C in the dark and monitored with a fluorescence microscope (magnification, ×100; BX51; Olympus Corp.) (33). Red-orange fluorescence was attributable to a potential-dependent aggregation in the mitochondria. Green fluorescence, reflecting the monomeric form of JC-1, appeared in the cytosol following mitochondrial membrane depolarization.

The level of ATP in cells was determined using an ATP Bioluminescence Assay kit (Beyotime Institute of Biotechnology) Harvested cultured cells (1×10⁶) were lysed with a PBS (Sigma-Aldrich; Merck KGaA), followed by centrifugation at 10,000 × g for 2 min at 4°C. The level of ATP was determined by mixing 50 µl supernatant with 50 µl luciferase reagent, which catalyzed the light production from ATP and luciferin. The emitted light was linearly compared with the ATP concentration and measured using a microplate luminometer.

MTT experiments were performed in 96-well plates. The cells were applied to the scaffold at a density of 10⁴ cells/well. Following 2–3 days of culture, the samples were washed 3 times with PBS, and 50 µl MTT was added to each well. The samples were subsequently incubated for 4 h at 37°C in a humid atmosphere containing 5% CO₂. The MTT solution was removed and 200 µl dimethyl sulfoxide was added to each sample and incubated for 10 min. Following the addition of Sorensen's buffer, the absorbance was determined at a wavelength of 570 nm (34).

A TUNEL assay to detect DNA fragmentation in the cell nuclei was performed using an In Situ Cell Death Detection kit (Roche Diagnostics GmbH, Mannheim, Germany) (35), according to the manufacturer's protocol. DAPI was used to label the nuclei (at room temperature for ~30 min). The results are expressed as apoptotic cells per tubule cross-section.

Cells (1×10⁶) in the treatment groups were lysed, centrifuged for 10 min at 1,600 × g at 4°C, and the supernatant (10 µl) was mixed with the pre-mixed reagent (according to the manufacturer's protocol) using a vortex mixer. This mixture was placed in a thermostatic water bath for 20 min at 37°C. The mixture was incubated with the color-developing agent for 10 min at room temperature. The absorbance value was determined at 560 nm using a microplate reader. The superoxide dismutase (SOD), glutathione (GSH) and malondialdehyde (MDA) levels were calculated using the standard formula, according to the protocols of the commercially-available ELISA kits (Beyotime Institute of Biotechnology, Haimen, China; cat. nos. S0086, S0131 and S0058, respectively).

Total RNA was isolated from cells using TRIzol™ reagent (Invitrogen; Thermo Fisher Scientific, Inc.). A total of 1 µg RNA from each sample was reverse-transcribed into cDNA using an RT kit (Eurogentec, Liege, Belgium) (36). The qPCR was performed with primers and matched probes from the Universal Fluorescence-labeled Probe Library (Roche Diagnostics GmbH). Quantification of gene expression was performed using an ABI PRISM 7500 Sequence Detection system (Applied Biosystems; Thermo Fisher Scientific, Inc.) with SYBR^® Green (Beiijing Transgen Biotech Co., Ltd., Beijing, China). The relative mRNA expression levels were normalized to that of ^β-actin using the 2^−ΔΔCq method (12). The thermocycling conditions were as follows: 95°C for 10 min, followed by 40 cycles of 95°C for 15 sec and 72°C for 35 sec, for PCR. The experiments were repeated three times with triplicates. The primers used in the present study were as follows: Interleukin (IL) 6, forward primer 5′-TCACCGATGTCTACCTGCTG-3′, reverse primer 5′-CACAGGGTTGAGCCAAAAGT-3′, C-C motif chemokine 2 (MCP1), forward primer 5′-CTCAACATCATGAAGGTCTC-3′, reverse primer 5′-GGCATTCAGTTCCAGGTCAG-3′, and GAPDH, forward primer 5′-ACGACATAGACGGCATCCA-3′, reverse primer 5′-GCTGTGGTTCAGTTGTGGTG-3′.

Caspase-3/9 activity kits (Beyotime Institute of Biotechnology) were used, according to the manufacturer's protocols (34). The relative caspase-3/9 activity was calculated from the ratio of treated cells to untreated cells (1×10⁶). The assays were repeated three times.

LDH is a fairly stable enzyme that is released from the cytosol into the culture medium as a consequence of cellular integrity damage. Thus, we used an LDH assay (Beyotime Institute of Biotechnology) to evaluate the presence of cell injury or damage. The level of LDH released was expressed as a percentage of the control group.

In order to evaluate the functional role of Parkin, small interfering (si)RNA was used to reduce its expression. The selective siRNA duplex (5′-GAGGAUGACAACGACAUAATT-3′, antisense, 5′-UUAUGUCGUUGUCAUCCUCTT-3′) and a nonspecific control duplex (5′-UUCUCCGAACGUGUCACGUTT-3′; antisense, 5′-ACGUGACACGUUCGGAGAATT-3′). These were obtained from YangZhou Ruibo Biotech Co., Ltd. (Yangzhou, China). Transfection of siRNA into cells was performed using Lipofectamine^® 2000 transfection reagent (Thermo Fisher Scientific, Inc.), according to the manufacturer's protocol (37). Cultured cells were washed with Opti-Minimal Essential Medium (Invitrogen; Thermo Fisher Scientific, Inc.) without serum or antibiotics and seeded in 6-well plates to 30–40% confluence. The transfection reagent and siRNA were diluted separately in serum-free medium, mixed and incubated for 10 min at room temperature to form the siRNA/lipid complex. This complex was added to each well at a final concentration of 70 nM/well siRNA. At 48 h post-transfection, cells were harvested to determine Parkin protein expression levels by western blot analysis according to the protocol described above.

All data are expressed as the mean ± standard deviation. Statistical analyses were performed with SPSS software (version 17.0; SPSS, Inc., Chicago, IL, USA). Statistical significance between two groups was determined by Student's t-test. Results for more than two groups were evaluated by one-way analysis of variance with the least significant difference test. P<0.05 was considered to indicate a statistically significant difference.

To examine the protective role of liraglutide in repairing the infarcted heart, fibrosis and cardiomyocytes death were assessed. Compared with the control group, the post-infarcted heart (28 days following myocardial infarction) exhibited extensive fibrosis, as evidenced by Sirius red staining (Fig. 1A and B). However, liraglutide had the ability to reduce cardiac fibrosis in a dose-dependent manner. To provide further evidence for the role of liraglutide in cardiac fibrosis in the post-infarcted heart, the expression of TGFβ and MMP9 was examined via western blotting. TGFβ and MMP9 are key feature of fibrosis. Compared with the control group, the post-infarcted heart exhibited more TGFβ and MMP9 expression (Fig. 1C-E). However, these phenotypic alterations were rescued by liraglutide in a concentration-dependent manner.

Since cardiomyocyte death is the primary reason for the development of cardiac fibrosis and heart failure following myocardial infarction, a TUNEL assay was used to observe cardiomyocyte apoptosis. As presented in Fig. 1F and G, the post-infarcted heart had an increased number of TUNEL-positive cells. However, treatment with liraglutide was able to reduce the ratio of TUNEL-positive cells. In order to provide further evidence for cellular death during cardiac remodeling, apoptotic protein expression was detected. Compared with the control group, caspase3 and its primary cleavage target PARP were increased in the post-infarcted heart (Fig. 1H-J), indicative of myocardial death in the post-infarcted heart. However, these alterations were reversed by treatment with liraglutide.

Apart from fibrosis and cellular death, the present study additionally examined the inflammatory response which is associated with cardiomyocyte death and fibrosis accumulation. Compared with the control group, increased expression of TNFα was observed in the myocardial tissue in the post-infarcted heart (Fig. 1K). Additionally, the transcriptional levels of IL6 and MCP1 were increased in the post-infarcted heart (Fig. 1L-M). However, treatment with liraglutide was able to repress the expression of the inflammatory markers. These data suggested that liraglutide had the ability to repair the damaged heart following myocardial infarction.

To gain an insight into the mechanism through which liraglutide maintained cardiomyocyte viability, a chronic hypoxia model with H9C2 cells was used to mimic the infarcted heart, and the molecular signaling responsible for the anti-apoptotic properties of liraglutide was examined (Fig. 2). Following 48 h of hypoxia stimulation, H9C2 apoptosis was detected via MTT and LDH release assays. Chronic hypoxia reduced cell viability, as evidenced by a reduced MTT value (Fig. 2A) and increased LDH release (Fig. 2B). However, treatment with liraglutide was able to inhibit the apoptotic signaling in a dose-dependent manner. Considering that the minimum protective concentration of liraglutide was 5 nM, this concentration was used in the following experiments. To further quantify H9C2 apoptosis under chronic hypoxia, a TUNEL assay was used. It was observed that hypoxia increased the number of TUNEL-positive cells (Fig. 2C and D), which was inhibited by liraglutide. Since liraglutide, a type of GLP1R agonist, exerts protective action on cells via GLP1R, Ex9-39 was used to block the GLP1R. Following the blockade of GLP1R, the protective effect of GLP1 on H9C2 apoptosis disappeared (Fig. 2C and D). In order to investigate whether mitochondrial apoptosis was involved in H9C2 apoptosis, cyt-c was assayed. In response to mitochondrial apoptosis, cyt-c was released from the mitochondria into the cytoplasm, where it interacts with caspase9 and activates caspase3. Therefore, immunofluorescence was used to observe the cyt-c cellular location. In the normal cells, cyt-c was observed in the cellular cytoplasm (Fig. 2G). However, hypoxia induced the diffusion of cyt-c into the cytoplasm and nucleus (Fig. 2G), indicating the activation of mitochondrial apoptosis. By contrast, liraglutide had the ability to reverse such alterations. Notably, the GLP1R blocker was able to reverse the inhibitory action of liraglutide on cyt-c diffusion.

To provide further evidence for mitochondrial apoptosis, alterations in the expression of proteins associated with cyt-c leakage were assessed. As presented in Fig. 2F and H-J, hypoxia elevated the expression of caspase3, caspase9 and Bax, and reduced the expression of c-IAP1. However, liraglutide was able to promote anti-apoptotic protein expression and limit pro-apoptotic protein expression (Fig. 2F and H-J). Notably, the anti-apoptotic effect of liraglutide was mediated by GLP1R. These data indicated that liraglutide sustained H9C2 survival in the context of chronic hypoxia damage by abating mitochondrial apoptosis.

As mitophagy is important for the maintenance of mitochondrial homeostasis, it was hypothesized that liraglutide may preserve mitochondrial balance via mitophagy (38). To validate this hypothesis, the expression of mitophagy markers was examined. Compared with the control group, hypoxia induced a downregulation of mitophagy, as evidenced by decreased LC3II, p62, Beclin1 and Atg5 expression (Fig. 3). However, liraglutide was able to reverse the expression of mitophagy parameters, and this effect was nullified by Ex9-39 (Fig. 3A, B and D-G). Notably, as Parkin is an important receptor in mitophagy, alterations in Parkin expression were examined. It was observed that hypoxia progressively reduced the expression of Parkin (Fig. 3A and B), which was rescued by treatment with liraglutide. However, with the blockade of GLP1R, Parkin expression was decreased (Fig. 3A and B). These data indicated that liraglutide triggered Parkin-associated mitophagy under hypoxia.

Additionally, to provide direct evidence for the role of liraglutide in the activation of mitophagy, immunofluorescence analysis was used to observe the overlap of mitochondria and lysosomes. As presented in Fig. 3C, in the control group, a number of mitochondria were tagged with lysosomes. However, following chronic hypoxic stimulation, the majority of mitochondria were separated from lysosomes (Fig. 3C), indicative of mitophagy inhibition. However, treatment with liraglutide was able to reverse the interaction between mitochondria and lysosomes, and contributed to the overlap of mitochondria and lysosomes. However, with blockade of GLP1R, mitophagy was inhibited despite treatment with liraglutide (Fig. 3C). These data demonstrated that liraglutide had the ability to reverse mitophagy activity under chronic hypoxic conditions.

To examine the consequences of mitophagy activation under treatment with liraglutide, mitochondrial function was evaluated. Since the generation of energy is the principal role of mitochondria, the cellular ATP content was detected (39). As presented in Fig. 4A, hypoxia repressed ATP production when compared with the control group. However, treatment with liragutide reversed the effect on ATP content. To examine whether Parkin-associated mitophagy was responsible for the protective role of liraglutide on ATP production, siRNA was used to knock down the expression of Parkin. Following silencing of Parkin in liraglutide-treated H9C2 cells, the ATP content was decreased (Fig. 4A). These data indicated that Parkin-associated mitophagy was involved in liraglutide-mediated ATP production.

ATP generation results from mitochondrial membrane potential (38). Mitochondria convert the proton gradient of mitochondrial membrane potential to chemical energy. Accordingly, the alteration in membrane potential was investigated. Through JC1 staining, it was observed that hypoxia reduced the mitochondrial membrane potential, as revealed by increased green fluorescence (Fig. 4B and C). However, liraglutide had the ability to maintain the membrane potential and this effect was reversed by Parkin knockdown. Aside from the collapse of mitochondrial membrane potential, it was observed that hypoxia induced excessive cellular oxidative stress, as evidenced by elevated ROS (Fig. 4D and E). Liraglutide had the ability to repress ROS release, whereas this effect was neutralized by Parkin silencing (Fig. 4D and E). As a consequence of ROS overproduction, a marked decline in the expression of antioxidant factors was observed, including GSH and SOD (Fig. 4F-H). By contrast, the expression of MDA, the lipid peroxide, was increased. Liraglutide had the ability to reverse the effect on GSH and SOD, although it repressed MDA in a Parkin-dependent manner (Fig. 4F-H). In order to examine whether mitophagy was associated with cellular death under chronic hypoxia, caspase3 activity was detected. As presented in Fig. 4I, the caspase3 activity was elevated under treatment with hypoxia and was reduced in response to liraglutide application. Notably, loss of Parkin increased caspase3 activity despite treatment with liraglutide. These data indicated that Parkin-associated mitophagy was responsible for the beneficial effects of liraglutide on mitochondria involving energy metabolism, mitochondrial membrane potential, cellular oxidative stress, redox biology and cellular apoptosis.

The molecular mechanism through which liraglutide regulated Parkin-dependent mitophagy was investigated in the present study (Fig. 5). SIRT1 has been identified to be a novel factor for mitochondrial homeostasis by sustaining mitochondrial membrane potential, reducing oxidative injury and promoting mitochondrial biogenesis (40). Based on this, it was hypothesized that SIRT1 may be the upstream signaling molecule for Parkin-dependent mitophagy. Western blotting was performed in the infarcted heart to examine the alterations in SIRT1 expression (Fig. 5A and B). It was observed that SIRT1 expression was downregulated in the infarcted heart. However, treatment with liraglutide was able to could increase SIRT1 expression in a dose dependent manner (Fig. 5A and B). Similarly, the downregulation of SIRT1 was also observed in H9C2 cells under chronic hypoxia (Fig. 5C-E), and this tendency was reversed by liraglutide treatment.

To demonstrate whether SIRT1 was involved in Parkin-dependent mitophagy, SRT (10 µM for 4 h), an activator of SIRT1, was used to reverse the decrease in SIRT1 expression under treatment with hypoxia. The SIRT1 inhibitor Selisistat (10 µM for 6 h) was applied to reduce SIRT1 expression in liraglutide-treated cells. As presented in Fig. 5C-E, activation of SIRT1 under hypoxic conditions was able to reverse the effect on Parkin expression, consistent with the results from the liraglutide group. However, in the liraglutide-treated cells, Selisistat not alleviated the SIRT1 expression and repressed Parkin expression (Fig. 5C-E). In order to provide further evidence for the regulatory role of SIRT1 in Parkin, co-staining for SIRT1 and Parkin was performed. It was observed that hypoxia reduced the level of SIRT1 and Parkin expression (Fig. 5H). However, these alterations were rescued by treatment with liraglutide or SRT (Fig. 5H). By contrast, following inhibition of SIRT1 with Selisistat, the beneficial effect of liraglutide on SIRT1 and Parkin disappeared.

Caspase9 activity and LDH release assays were performed to investigate whether SIRT1 was involved in mitochondrial protection and cellular survival. It was observed that caspase9 activity and LDH content were increased in the hypoxia group (Fig. 5F and G). However, treatment with liraglutide or SRT application was able to abate the caspase9 activity and LDH release (Fig. 5F and G). Notably, following the blockade of SIRT1 with Selisistat, caspase9 activity and LDH content were increased despite treatment with liraglutide. These data illustrated that SIRT1 was the upstream molecule for Parkin-dependent mitophagy, and that SIRT1 contributed to mitochondrial protection and cellular survival. The beneficial role of liraglutide in the post-infarcted heart through regulation of SIRT1-mediated mitophagy was summarized in Fig. 6.