The rat H9c2 cardiomyocyte cell line was obtained from the Cell Bank of Chinese Academy of Sciences (Shanghai, China). To establish a model of MI/R in vitro, the cardiomyocytes were first cultured in serum/glucose-free Dulbecco's modified Eagle's medium (DMEM; Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) and an anoxic environment (95% N₂ and 5% CO₂) at 37°C for 2 h. The gas mixture of 95% N₂ and 5% CO₂ was replaced with air at a flow velocity of 2 l/min. Subsequent to the hypoxia process, the cardiomyocytes were transferred into fresh medium (DMEM) and maintained in an oxygen-rich incubator (37°C; 95% O₂ and 5% CO₂). Following incubation for 1 h, the MI/R cell model was harvested.

Five treatment groups were prepared, which were the control group (cardiomyocytes with no treatment), MI/R group (cardiomyocytes treated with OGD/R injury), 20 µM RP+MI/R group (cardiomyocytes treated with OGD/R injury, and subsequently treated with 20 µM RP at 37°C for 1 h), 40 µM RP+MI/R group (cardiomyocytes treated with OGD/R injury, and subsequently treated with 40 µM RP at 37°C for 1 h) and 80 µM RP+MI/R group (cardiomyocytes treated with OGD/R injury, and subsequently treated with 80 µM RP at 37°C for 1 h). RP was purchased from Beijing Solarbio Science & Technology Co., Ltd. (Purity >98%; Beijing, China).

A Cell Counting Kit-8 (CCK-8; Beyotime Institute of Biotechnology, Haimen, China) was performed to assess the cell viability of the cardiomyocytes. Cardiomyocytes in the logarithmic phase were seeded into 96-well plates at a density of ~6×10⁴ cells/ml/group, and maintained in a 5% CO₂ atmosphere at 37°C for 12 h. Subsequently, the cardiomyocytes were maintained for 12, 24 and 48 h. A total of 10 µl CCK reagent was supplemented into the wells of the 96-well plates. The cardiomyocytes were maintained for 3 h. A microplate reader (Bio-Rad Laboratories, Inc., Hercules, CA, USA) was used to record the absorbance at 450 nm. Cell viability was evaluated as the percentage of cell survival compared with the control.

The lactate dehydrogenase (LDH; cat. no. C0016) detection kit, malondialdehyde (MDA; cat. no. S0131) detection kit, superoxide dismutase (SOD; cat. no. S0101) assay kit and glutathione peroxidase (GPx; cat. no. S0058) assay kit were purchased from Beyotime Institute of Biotechnology. Detection of MDA content, LDH activity, SOD activity and Gpx activity were performed according to the manufacturer's protocols.

Reactive oxygen species (ROS) levels were detected using 2′,7′-dichlorodihydrofluorescein diacetate (DCFH-DA; Sigma-Aldrich; Merck KGaA, Darmstadt, Germany). A total of 2×10⁶ cells/ml were seeded in a 6-well plate and 10 mM DCFH-DA in PBS was added to the cells for 20 min at 37°C. DCF fluorescence was measured using a BD LSR II flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA). The flow cytometry data were analyzed using FlowJo software version 7.5.5 (FlowJo LLC, Ashland, OR, USA).

Flow cytometry (FCM) was conducted to assess the apoptosis of cardiomyocytes. Following washing with PBS, cultured cardiomyocytes were trypsinized with 0.25% trypsin (Beyotime Institute of Biotechnology). The cells were harvested and centrifuged at 1,000 × g for 5 min at 4°C. The supernatant was removed and the cardiomyocytes were suspended in the incubation buffer at a density of 1×10⁶ cells/ml for assessment. Apoptosis of cardiomyocytes was detected with the Annexin V-fluorescein isothiocyanate and propidium iodide detection kit (cat. no. V13242, Invitrogen; Thermo Fisher Scientific, Inc.), according to the manufacturer's instructions. A flow cytometer (FACSCalibur) running BD CellQuest™ software version 1.2 (both from BD Biosciences) was used to assess cell apoptosis.

PBS was added to the cultured cardiomyocytes until the cell concentration reached 1×10⁶ cells/ml. Rho 123 (5 µM; cat. no. R8004; Sigma-Aldrich; Merck KGaA) or Fluo-3AM [3 µM; cat. no. 70-F1243; Multisciences (Lianke) Biotech Co., Ltd., Hangzhou, China] were added to the cardiomyocytes. The cardiomyocytes were incubated at room temperature and kept away from light for 10 min. FCM was performed to assess the MMP and Ca²⁺ level in cardiomyocytes, and the excitation wavelength was 488 nm. In total, 10,000 cells were collected from each sample. The data was analyzed using BD CellQuest™ software version 1.2 (BD Biosciences).

Mitochondria were isolated from cells using a Mitochondrial extraction kit (cat. no. SM0020; Beijing Solarbio Science & Technology Co., Ltd.), according to the manufacturer's protocols. Tumescent fluid (containing 120 mmol/l KCl, 20 mmol/l MOPS, 10 mmol/l Tris-HCl and 5 mmol/l KH₂PO₄; pH 7.4; Beyotime Institute of Biotechnology) was added to the mitochondria and the mixture was diluted to the point where the protein concentration reached 0.25 g/l at 25°C. Subsequently, 200 µmol/l CaCl₂ was added to the mitochondria and the absorbance at 520 nm was observed for 15 min. The alterations in absorbance suggested the degree of mitochondrial swelling, which indicated the opening of mPTP.

Proteins were isolated using NP-40 lysis buffer (Beyotime Institute of Biotechnology, China). The BCA Protein quantification kit (cat. no. ab207002; Abcam, Cambridge, UK) was used to determine protein concentration. A total of 25 µg/lane protein was separated by 12% SDS-PAGE. The separated products were transferred to a polyvinylidene difluoride membrane (EMD Millipore, Billerica, MA, USA). The membranes were blocked with 5% skimmed milk at room temperature for 2 h. Western blotting was performed with specific primary antibodies at 4°C overnight: anti-inactive-caspase (1:2,000; cat. no. ab184787; rabbit anti-rat); anti-active-caspase-3 (1:500; cat. no. ab49822; rabbit anti-rat); anti-inactive-caspase-9 (1:500; cat. no. ab138412; rabbit anti-rat); anti-active-caspase-9 (1:1,000; cat. no. ab25758; rabbit anti-rat); anti-mitochondrial cyto c (1:5,000; cat. no. ab133504; rabbit anti-rat); anti-mitochondrial AIF (1:1,000; cat. no. ab32516; rabbit anti-rat); anti-cyto c (1:1,000; cat. no. ab110325; rabbit anti-rat); anti-AIF (1:1,000; cat. no. ab1998; rabbit anti-rat); anti-cyclooxygenase (COX) IV (1:2,000; cat. no. ab16056; rabbit anti-rat); and anti-actin (1:5,000; cat. no. ab179467; rabbit anti-rat) (all from Abcam). Horseradish peroxidase-conjugated secondary antibodies (1:5,000; Abcam; cat. no. ab205718; goat anti-rabbit) were supplemented and incubated at room temperature for 1 h. Enhanced chemiluminescent (ECL) reagents (EMD Millipore) in combination with an ECL system (GE Healthcare, Chicago, IL, USA) were used to assess the results. The density of the protein bands was quantified using Quantity One^® software version 4.2.1 (Bio-Rad Laboratories, Inc.).

Total RNA was extracted from the cultured cardiomyocytes using TRIzol^® reagent (Thermo Fisher Scientific, Inc.). RNA was reverse transcribed to cDNA using BeyoRT™ II cDNA Synthesis kit (Beyotime Institute of Biotechnology), according to the manufacturer's protocol. qPCR was performed using a SYBR-Green PCR Master Mix kit (Applied Biosystems; Thermo Fisher Scientific, Inc.) on a ABI 7500 Thermocycler (Applied Biosystems). PCR cycling conditions were as follows: 10 min pretreatment at 94°C, at 95°C for 15 sec, at 68°C for 45 sec (45 cycles), at 95°C for 15 sec, at 68°C for 1 min, at 94°C for 15 sec, a final extension at 75°C for 10 min and held at 4°C. The primers were purchased from Invitrogen (Thermo Fisher Scientific, Inc.): Caspase-3, forward, 5′-TGTCGATGCAGCTAACCTCA-3′ and reverse, 5′-GCAGTAGTCGCCTCTGAAGA-3′ (product: 241 bp); caspase-9, forward, 5′-CATTGGTTCTGGCAGAGCTC-3′ and reverse, 5′-AGCAGTCAGGTCGTTCTTCA-3′ (product: 238 bp); cyto c, forward, 5′-CAACTGCACAAGACAACCCA-3′ and reverse, 5′-ATAGCACAATCCCCACCACA-3′ (product: 204 bp); AIF, forward, 5′-ACATGCGACCTCCTCTTTCA-3′ and reverse, 5′-TGCCTCTTACATCCAGGTGG-3′ (product: 213 bp); COX IV, forward, 5′-TGAAGGAGAAGGAGAAGGCC-3′ and reverse, 5′-ACCCAGTCACGATCAAAGGT-3′ (product: 229 bp); actin, forward, 5′-CAACATGGATGAGCGGAAGG-3′ and reverse, 5′-GCAGTGTAGCAGCATCGAAA-3′ (product: 233 bp). Actin was used as the control of the input RNA level. The 2^−ΔΔCq method was used to analyze the relative gene expression (28).

Results in the present study are presented as the mean ± standard error of the mean with at least three independent experiments repeated. All experimental data were analyzed one-way analysis of variance followed by the Dunnett's test. P<0.05 was considered to indicate a statistically significant difference. Data analysis was performed using GraphPad Prism version 6.0 (GraphPad Software, Inc., La Jolla, CA, USA).

The cell viabilities of cardiomyocytes treated with different concentrations of RP were measured in the present study. It was observed that compared with the control group, the cell activities of cardiomyocytes treated with RP at concentrations of 10, 20, 40 and 80 µM exhibited no significant difference (Fig. 1A). However, it was identified that with the treatments of 160 and 320 µM RP, the cell viabilities of cardiomyocytes were significantly decreased, particularly at 48 h treatment (P<0.05; Fig. 1A). These results suggested that low concentrations of RP exhibited a limited effect on the cell viability of cardiomyocytes. Therefore, the cell viabilities of cardiomyocytes subjected to MI/R injury and MI/R-induced cardiomyocytes treated with 20, 40 and 80 µM RP were further evaluated. According to the results, it was observed that MI/R injury decreased the cell viability of cardiomyocytes; whereas, RP increased the cell viability of MI/R-induced cardiomyocytes in a dose-dependent manner (Fig. 1B). The results indicated that RP was able to enhance the cell viability of cardiomyocytes, which were subjected to MI/R injury. Therefore, it was confirmed that MI/R resulted in low cell viability of cardiomyocytes; whereas, RP increased the cell viability of MI/R-induced cardiomyocytes within a certain concentration range.

Oxidative stress has been proposed to contribute to the development and progression of MI/R (5). Therefore, the levels of oxidative stress markers, including ROS, MDA, LDH, SOD and GPx in cardiomyocytes treated with MI/R and different concentrations of RP were assessed. The ROS, MDA and LDH content in cardiomyocytes treated with MI/R in advance were markedly higher compared with the control, while treatment with ≥40 µM RP significantly decreased the ROS, MDA and LDH content in MI/R-induced cardiomyocytes (P<0.05; Fig. 2A-C). However, MI/R was observed to be able to reduce the activities of SOD and GPx in cardiomyocytes. The SOD and GPx activity in MI/R-induced cardiomyocytes was significantly increased (P<0.05; Fig. 2D and E) following treatment with ≥40 µM RP. It was suggested that RP reduced oxidative stress in MI/R-induced cardiomyocytes. The opening degree of mPTP in cardiomyocytes was assessed by spectrophotometry. The results revealed that MI/R increased the mPTP opening degree in cardiomyocytes (P<0.05; Fig. 2F); whereas, RP decreased the opening degree of mPTP in MI/R-induced cardiomyocytes.

Furthermore, the Ca²⁺ and MMP levels in cardiomyocytes treated with MI/R and different concentrations of RP were measured. Based on the FCM data, it was identified that MI/R markedly increased the Ca²⁺ level in cardiomyocytes. Following treatment with 40 and 80 µM RP, the Ca²⁺ level in MI/R-induced cardiomyocytes was significantly decreased compared with the MI/R group (P<0.05; Fig. 3A), suggesting that RP was able to decrease the Ca²⁺ level in MI/R-induced cardiomyocytes. Additionally, the FCM results on MMP revealed that MI/R significantly decreased the MMP level in cardiomyocytes; whereas, treatment with 40 and 80 µM RP was observed to significantly increase the MMP levels in MI/R-induced cardiomyocytes (P<0.01; Fig. 3B). These results suggested that RP was able to modulate the Ca²⁺ and MMP levels in MI/R-induced cardiomyocytes.

The mitochondrial mechanisms of RP in protecting cardiomyocytes against MI/R induced injury were examined. In this analysis, the mitochondrial cyto c, mitochondrial AIF, cytosolic cyto c and cytosolic AIF expression in cardiomyocytes were measured. In the cardiomyocytes from the MI/R groups, the mitochondrial cyto c expression was decreased compared with the control, while the expression level of mitochondrial AIF was markedly increased compared with the control. Increases were observed in the mitochondrial cyto c expression in MI/R-induced cardiomyocytes treated with 40 and 80 µM RP. It was additionally identified that RP was able to downregulate the expression level of mitochondrial AIF in MI/R-induced cardiomyocytes, with significant decreases observed in the 40 and 80 µM RP+MI/R groups (P<0.01; Fig. 4A and B). Additionally, the expression level of cytosolic cyto c in cardiomyocytes was upregulated upon MI/R injury, while cytosol AIF expression was reduced by MI/R injury (P<0.05). Following treatment with RP, the expression level of cytosolic cyto c was decreased; whereas, the cytosolic AIF expression was significantly increased in MI/R-induced cardiomyocytes (P<0.05; Fig. 4C and D). Therefore, it was demonstrated that RP upregulated the mitochondrial cyto c and cytosolic AIF expression, and downregulated the expression levels of mitochondrial AIF and cytosolic cyto c in MI/R-induced cardiomyocytes.

From the aforementioned results, it was determined that RP was able to enhance the cell viability of MI/R-induced cardiomyocytes; therefore, the apoptotic capacity of MI/R-induced cardiomyocytes treated with RP was evaluated. It was demonstrated that the proportion of apoptotic cardiomyocytes in MI/R group was 26.46%, which was significantly increased compared with the control (6.32%). Furthermore, following treatment with 20, 40 and 80 µM RP, the percentage of apoptotic MI/R-induced cardiomyocytes decreased from 26.46 to 23, 18.52 and 14.1%, respectively (Fig. 5). In particular, significant decreases in the percentage of apoptotic cells were observed in the 40 and 80 µM RP+MI/R groups (P<0.05). These data suggested that RP markedly decreased the apoptotic capacity of MI/R-induced cardiomyocytes in a dose-dependent manner.

According to the apoptosis results above, it was demonstrated that RP inhibited the apoptosis of cardiomyocytes induced by MI/R. Therefore, the apoptosis-associated proteins in cardiomyocytes were further investigated. According to the RT-qPCR data, it was observed that the caspase-3 and caspase-9 expression in cardiomyocytes from MI/R group was significantly increased compared with the control; whereas, the caspase-3 and caspase-9 expressions in MI/R-induced cardiomyocytes were decreased by RP in a dose-dependent manner, with significant differences observed with 40 and 80 µM RP (P<0.05; Fig. 6A). Furthermore, the western blot analysis revealed that the expression levels of inactive caspase-3 and −9 in cardiomyocytes were markedly downregulated with MI/R injury; however, were upregulated upon treatment with different concentrations of RP. Additionally, it was identified that MI/R upregulated the active caspase-3 and −9 expression in cardiomyocytes. However, RP was able to decrease the expression levels of caspase-3 and −9 in MI/R-induced cardiomyocytes compared with the control (Fig. 6B). Therefore, it was demonstrated that RP modulated the apoptosis-associated protein expression in MI/R-induced cardiomyocytes.