TA (cat. no. T0801) was purchased from Shanghai Topscience Co., Ltd. N-Acetylcysteine (NAC; cat. no. HY-B0215) was purchased from MedChemExpress. LPS (from Escherichia coli O55:B5) was purchased from Sigma-Aldrich; Merck KGaA, and the rat heart myoblast cell line H9C2(2-1) (cat. no. GNR 5) was obtained from The Cell Bank of Type Culture Collection of The Chinese Academy of Sciences. High-glucose DMEM (cat. no. 12800017) and fetal bovine serum (FBS; cat. no. 10270106) were purchased from Gibco; Thermo Fisher Scientific, Inc. Phosphate-buffered saline (PBS; cat. no. SH30256.01) and trypsin (cat. no. SH30042.01) were purchased from HyClone; Cytiva. Pierce™ BCA Protein Assay kit (cat. no. 23227) was obtained from Thermo Fisher Scientific, Inc. Cellular ROS Detection Assay kit was purchased from Nanjing Jiancheng Bioengineering Institute. Cell Counting Kit-8 (CCK-8) assay was purchased from Seven Seas Futai. Culture dishes (cat. nos. 430166 and 430167) were purchased from Corning, Inc.

The primary antibody directed against β-actin (cat. no. ab8227) was purchased from Abcam. Primary antibodies against cleaved caspase-3 (cat. no. BF0711), cleaved caspase-9 (cat. no. AF5240), Bax (cat. no. AF0120), Bcl-2 (cat. no. AF6139), ATF6 (cat. no. DF6009), CHOP (cat. no. F6025), phosphorylated (p)-PERK (cat. no. DF7576), PERK (cat. no. AF4799), p-IRE1 (cat. no. AF7150), IRE1 (cat. no. DF7709), p-JNK (cat. no. AF3318) and JNK (cat. no. AF6318) were obtained from Affinity Biosciences. HRP-conjugated secondary antibodies (cat nos. ab6721 and ab6885) were purchased from Abcam.

The H9C2 cells were cultured in high-glucose DMEM supplemented with 10% (v/v) FBS, 1.5 g/l NaHCO3 and 1% penicillin-streptomycin (Gibco; Thermo Fisher Scientific, Inc.) at 37°C in a humidified atmosphere containing 5% CO2 and were maintained in a logarithmic growth phase for all experiments. According to a previous study (31), H9C2 cells were treated with LPS (15 µg/ml) for 12 h, and with different concentrations of TA (0, 5, 10, 20, 30 and 40 µM) for 12 or 24 h. In addition, H9C2 cells were pretreated with 10 µM TA 2 h prior to treatment with LPS (15 µg/ml) for 12 h. For ROS inhibition experiments, the inhibitor NAC (2 mM) was added for 1 h prior to exposure to 10 µM TA. H9C2 cells were treated with TA (10 µM) and NAC (2 mM) alone for 12 h. All treatments were performed at 37°C in a humidified atmosphere containing 5% CO2.

The H9C2 cells were seeded in a 96-well plate at a density of 5,000 cells/well, and treated as follows: Fresh culture medium alone (control group); fresh culture medium with different concentrations (0–50 µM) of TA and/or fresh culture medium with 15 µg/ml LPS for the indicated times. All treatments were performed at 37°C in a humidified atmosphere containing 5% CO2. Cell viability was assessed using the CCK-8 assay, according to the manufacturer's instructions. Briefly, after treatment, CCK-8 solution (10 µl) was added to each well and the cultures were incubated at 37°C in humidified atmosphere containing 95% air and 5% CO2 for 1 h. Absorbance was measured at 450 nm using a microplate reader (Bio-Rad Laboratories, Inc.). Values were expressed as the percentage of viable cells as follows: Relative viability (%)=[OD450 (treated) - OD450 (blank)]/[OD450 (control) - OD450 (blank)] × 100. To exclude cell toxicity, one concentration of TA (10 µM) was used in subsequent experiments.

The Cellular ROS Detection Assay kit contains the fluorogenic dye, 2,7-dichlorofluorescin-diacetate (DCFDA), which measures ROS activity within the cell. After diffusion into the cell, DCFDA is deacetylated by cellular esterases to a non-fluorescent compound, which is later oxidized by ROS into 2′,7′-dichlorofluorescein (DCF). DCF is a highly fluorescent compound, which can be detected by fluorescence spectroscopy with excitation/emission at 485/535 nm. In the present study, the H9C2 cells (70–80% confluent) were incubated with DCFDA (10 µM) for 30 min at 37°C in the dark. The cells were subsequently washed twice with PBS and observed under a confocal laser scanning microscope (Leica Microsystems, Inc.), and the images were captured. The mean fluorescence intensity of DCF was analyzed using ImageJ (version 1.46) software (National Institutes of Health).

The H9C2 cells were also collected to assess malondialdehyde (MDA) and nicotinamide adenine dinucleotide phosphate (NADPH) levels; 0.1 ml lysate or homogenate extracted from 10⁶ cells was used. After homogenization or pyrolysis, the supernatant was centrifuged at 6,000 × g for 10 min at 4°C and the supernatant was then stored at −80°C. Subsequently, MDA (cat. no. S0131S) and NADPH (cat. no. ST360) levels were detected according to the instructions of commercial kits (Beyotime Institute of Biotechnology).

Annexin V-FITC was used to measure the apoptosis of H9C2 cells as described previously (32). The necrotic cells were counterstained with propidium iodide (PI). Briefly, cells were plated in 60-mm dishes and exposed to different treatments once they reached 80–85% confluence. Detached and adherent cells were collected, washed twice with cold PBS, and then resuspended in 1X binding buffer [BD Pharmingen™ FITC Annexin V Apoptosis Detection kit I (cat. no.556547)] at a concentration of 1×10⁶ cells/ml. The resuspended solution containing 1×10⁵ cells (100 µl) was transferred to a 5-ml culture tube, to which 5 µl Annexin V-FITC and 5 µl PI were added, and gently vortexed prior to 15-min incubation at 25°C in the dark. Subsequently, 1X binding buffer (400 µl) was added to each tube at room temperature and the percentages of early + late apoptotic cells were analyzed by flow cytometry (Beckman Coulter; COULTER-XL.MCL) within 1 h and EXPO32 (Applied Cytometry Systems) was used for analysis.

RT-qPCR analysis was performed as described previously (33). The primer sequences are described in Table I. Briefly, total RNA was extracted using TRIzol^® (cat. no. 15596026; Invitrogen; Thermo Fisher Scientific, Inc.) and DNase was used to remove internal DNA contamination. Random primers (cat. no. K1622; Thermo Fisher Scientific, Inc.) were used in RT reactions to obtain first-strand cDNA according to the manufacturer's instructions, and SYBR-Green qPCR Master Mix (cat. no. FP202; Tiangen Biotech Co., Ltd.) was used to amplify the target fragments. Gapdh cDNA amplification was used as an internal control. The general qPCR procedure included an initial denaturing step at 95°C for 15 min, followed by 37 cycles of denaturing at 95°C for 10 sec, annealing at 57°C for 30 sec and extension at 72°C for 30 sec, with a plate read at 95°C for 5 sec. GAPDH was used as a reference gene with the DCT method to quantify the results (33). Each experimental group was assessed in triplicate. IBM SPSS (version 19) software was used to analyze differences in expression.

Western blotting was performed as described previously (34). Briefly, cells were lysed in lysis buffer containing 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 0.25% sodium deoxycholate and 1% NP-40, with a protease inhibitor cocktail and phosphatase inhibitor cocktail; the insoluble components were removed by centrifugation at 12,000 × g for 15 min at 4°C. Subsequently, protein concentrations were measured using the BCA method. After denaturation, the total proteins (25 µg) were separated by SDS-PAGE on 10% gels and transferred onto 0.2- or 0.45-µm polyvinylidene fluoride membranes. After blocking with 5% (w/v) non-fat milk for 2 h at room temperature, the membranes were incubated with primary antibodies diluted in blocking buffer (1:1,000) overnight at 4°C. The membranes were subsequently washed three times with 1X TBS-0.1% Tween-20 (TBST), and further incubated with HRP-conjugated secondary antibodies (1:5,000) for 2 h at room temperature. After subsequent washes of the membrane, three times with 1X TBST, antibody detection was performed using the Immobilon Western Chemiluminescent HRP Substrate (cat. no. WBKLS0500; Merck KGaA) and a ChemiDoc™ XRS+ system (Bio-Rad Laboratories, Inc.). ImageJ (version 1.46; National Institutes of Health) was used to compare the gray values of the target bands; GAPDH served as an internal control.

Data are presented as the mean ± standard error of the mean from at least three independent experiments. GraphPad Prism 8.0.2 (GraphPad Software Inc.) was used for statistical analysis. Statistical significance was determined by one-way ANOVA with Bonferroni correction for multiple comparisons. P<0.05 was considered to indicate a statistically significant difference.

Prior to investigating whether TA protected H9C2 cells from LPS-induced cell injury, the cytotoxicity of the extract was determined. Cell viability was assessed using the CCK-8 assay and the results revealed that cell viability was slightly increased in response to TA concentrations ≤10 µM; however, these increases were not statistically significant when compared with the control group (Fig. 1A). Following treatment of H9C2 cells with TA concentrations ≥30 µM, cell viability was significantly decreased; therefore, to exclude cell toxicity, 5, 10 and 20 µM concentrations were selected for the subsequent experiment. To examine the effects of TA on LPS-induced cell injury, the viability of H9C2 cells pretreated with 0, 5, 10, 15 and 20 µM TA and then treated with 15 µg/ml LPS was determined. As shown in Fig. 1B, LPS treatment significantly diminished cell viability to 50.65±0.5% compared with the control group. Notably, pretreatment with TA (5, 10 and 15 µM) prior to LPS treatment significantly improved cell viability compared with the LPS-treated group to 68.3±0.8, 84.6±2.1 and 90.6±3.3%, respectively. Cell viability was slightly decreased in response to 20 µM TA compared with 15 µM TA. These findings demonstrated that pretreatment with TA dose-dependently increased cell viability, which was diminished by LPS treatment, thus suggesting that TA may be capable of protecting H9C2 cells from LPS-induced cell injury.

To further evaluate the effect of TA on LPS-induced apoptosis, H9C2 cells were treated with LPS or a combination of TA and LPS. Cell apoptosis was measured by flow cytometry, and the expression levels of apoptosis-associated proteins in H9C2 cells were measured by western blotting. As presented in Fig. 2A and B, the percentage of apoptotic cells was significantly increased after LPS treatment compared with that in the control group. Conversely, pretreatment with TA significantly decreased the percentage of apoptotic cells induced by LPS treatment. Consistent with the results of flow cytometry, LPS increased the protein expression levels of apoptotic markers, Bax/Bcl2, cleaved caspase-3 and cleaved caspase-9, compared with those in the control group. However, pretreatment with TA decreased the expression levels of these markers compared with treatment with LPS alone (Fig. 2C and D). Notably, the detected levels of all markers in the TA-only group were comparable to those detected in the control (medium) group. These results suggested that TA alleviated LPS-induced H9C2 cell apoptosis.

The level of ROS is a crucial index of oxidative stress. To further evaluate the effects of TA on LPS-induced oxidative stress, the levels of intracellular ROS were monitored by detecting changes in DCF fluorescence by laser confocal microscopy. As shown in Fig. 3A and B, LPS treatment significantly elevated ROS generation compared with that in the control group; however, the LPS-mediated increase in DCF fluorescence was significantly decreased with TA pretreatment. Notably, the ROS levels detected in the TA-only group were comparable to those in the control group treated with medium only, demonstrating the minimum cell toxicity of the chosen TA dose.

The results of the present study demonstrated that LPS significantly increased the content of MDA in cells compared with that in the control group. The levels of MDA in the TA pretreatment group were significantly lower than those in the LPS treatment group (Fig. 3C). In addition, LPS treatment decreased the content of NADPH compared with that in the control group, whereas TA treatment significantly improved the levels of NADPH compared with those in the LPS treatment group (Fig. 3D). These results suggested that TA was capable of reducing intracellular ROS accumulation following LPS stimulation. The ability of TA to inhibit intracellular ROS production may contribute to its efficacy at protecting H9C2 cells against LPS-induced apoptosis.

To determine the effect of TA on ERS in H9C2 cells induced by LPS, the expression levels of ERS associated proteins were examined. As presented in Fig. 4A-C, the expression levels of ERS associated proapoptotic proteins, CHOP, ATF6, p-PERK, PERK and p-IRE1/IRE1, were significantly increased following exposure to LPS compared with those in the control group. Moreover, both phosphorylated and total PERK levels increased following treatment, indicating that changes in phosphorylation of PERK may serve an important role in the PERK signaling pathway. By contrast, the expression levels of ERS associated proteins were markedly downregulated in the TA + LPS group compared with those in the group treated with LPS only. This result suggested that pretreatment with TA significantly downregulated the expression of ERS associated proteins, which alleviated LPS-induced ERS. Based on this finding, it may be hypothesized that the ERS associated proapoptotic proteins CHOP, ATF6, PERK and IRE1 could serve underlying roles in alleviating LPS-induced H9C2 cell apoptosis by TA.

The present study further examined the protective effects of TA on H9C2 cells and explored the relationship between ROS and ERS-associated apoptosis in LPS-induced H9C2 cells. To determine whether LPS induced ROS-modulated ERS in H9C2 cells, the ROS scavenger, NAC, was used to evaluate the relationship between ROS, ERS and apoptosis. H9C2 cells were pretreated with 1 mM NAC or 10 µM TA, and then treated with 15 µg/ml LPS. The protein expression levels of ERS associated proteins (CHOP, ATF6, p-PERK, PERK, p-IRE1 and IRE1) and apoptosis-associated proteins (p-JNK, JNK, cleaved caspase-3, Bax/Bcl-2 and cleaved caspase-9) were detected by western blotting (Figs. 5A-C and 6A-C). In addition, the mRNA expression levels of ER stress associated genes [Chop, Atf6, Perk, Ire1 and spliced X box-binding protein 1) and apoptosis-associated genes (Bax, caspase-3, caspase-12 and cytochrome c (Cyt c)] were detected (Figs. 5D-H and 6D-G). LPS caused a significant increase in the expression of ERS associated proteins and apoptosis-associated proteins; however, these protein expression levels were markedly decreased following TA or NAC pretreatment (Figs. 5 and 6). Moreover, TA in the presence of ROS inhibitor NAC had no further effect on expression of ERS associated proteins and apoptosis-associated proteins compared with TA alone. The levels of all markers detected in the NAC-only group were similar to those observed in the control group. The present results indicated that ERS activation and apoptosis activation were inhibited by TA in a ROS-dependent manner, but not in the presence of NAC. ERS is considered one of the mechanisms contributing to ROS-mediated cell apoptosis (18). The present results also demonstrated that LPS-induced ROS modulated ERS in H9C2 cells (Fig. 5), suggesting that the ROS-mediated ERS pathway may be one of the important mechanistic pathways modulating apoptosis in LPS-induced H9C2 cells. Taken together, TA may prevent LPS-induced ROS-mediated ERS, which may be responsible for the TA-induced decrease in H9C2 cell apoptosis.