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Introduction

Oxidative stress is important in various disease processes, including in cancer, inflammation, cardiovascular diseases, atherosclerosis, central nervous system disorders, neurode-generative diseases, diabetes and respiratory diseases. Almost all human organs can be damaged by oxidative stress (1–5). In the cardiovascular system, reactive oxygen species (ROS) induce the oxidation of low density lipoprotein, cholesterol, cholesterol-derived species and protein modifications, which can lead to foam cell formation, atherosclerotic plaques and vascular thrombosis (6). Various studies have previously demonstrated that cardiomyocyte damage induced by heart ischemia/reperfusion is predominantly due to the generation of ROS (7–9). Other studies also indicated that ROS can damage the sarcoplasmic reticulum of cardiac cells, inducing contractile dysfunction and Ca2+ release by modifying the structure and function of cardiac proteins, which may be important in the formation of myocardial ischemia/reperfusion injury (9,10). Several investigations have demonstrated that ROS induce cardiomyocyte apoptosis by activating various signaling pathways, including mitogen-activated protein kinase 14 (p38MAPK), MAPK 1 (also known as ERK1/2), MAPK 8 (also known as JNK) and v-akt murine thymoma viral oncogene homolog 1 (Akt1) signaling, which may contribute to the development and progression of cardiac dysfunction and heart failure (11–13). Additionally, angiotensin II stimulates ROS-mediated activation of the transcription factor nuclear factor-κB, which is understood to be involved in the induction of cardiac hypertrophy. ROS also regulates the transcription of jun proto-oncogene, which influences the expression of other genes in cardiac hypertrophy (14). In summary, oxidative stress participates in a variety of pathological mechanisms associated with cardiomyocyte diseases.

Numerous in vitro studies of oxidative stress in cardiomyocytes have been performed using H9c2 cardiomyocytes. H9c2 cells are a clonal cardiomyocyte cell line derived from embryonic rat ventricles (15), with a similar profile of signaling mechanisms to adult cardiomyocytes. Under oxidative stress, H9c2 cardiomyocytes respond in a similar manner to myocytes in primary cultures or isolated heart experiments (16). H9c2 cells have been demonstrated to be a useful tool for the study of the cellular mechanisms and signal transduction pathways of cardiomyocytes (17–20).

H2O2-treated H9c2 cells have been commonly used as an in vitro model for studying oxidative stress in cardio-myocytes, and to evaluate the cardioprotective effects of drugs against oxidative damage (21–24). However, to the best of our knowledge, H9c2 cells have not been previously used for high-throughput drug screening. The current study used this model to establish a cell-based screening assay in a high throughput format. From a library of traditional Chinese medicine (TCM) extracts, 17 primary hits were identified, 2 of which were further validated as cardiopro-tective agents against oxidative damage. The present study demonstrated the used of the H2O2-induced cell damage model in a high-throughput screening (HTS) assay, which may be established as an efficient and low-cost HTS assay for the identification of candidate drugs that reduce oxidative damage from large TCM extract/chemical libraries.

Materials and methods

Cell culture

H9c2 cells (Cell Resource Centre of the Shanghai Institutes for Biological Sciences, Chinese Academy of Science, Shanghai, China) were maintained in Dulbecco's modified Eagle's medium (Gibco; Thermo Fisher Scientific, Inc., Waltham, MA, USA) containing 10% FBS (Gibco; Thermo Fisher Scientific, Inc.) and incubated at 37°C in a humid atmosphere of 5% CO2. Following expansion, cells at passage 3 were used for all experiments.

Cell counting kit (CCK)-8 assays

H9c2 cells were used to establish the cell model of oxidative damage. H9c2 cells (100 µl/well) were seeded into 96-well plate at a density of 3.0×104 cells/ml and incubated at 37°C, 5% CO2 overnight. The cells were then treated with 100 µl 50 µmol/l H2O2 (Shandong Siqiang Chemical Group Co., Ltd., Liaocheng, China) for 3 h. Following H2O2 treatment, a CCK-8 assay kit (BestBio, Shanghai, China) was used to detect cell viabilities according to the manufacturer's instructions. In brief, following treatment with H2O2, 10 µl CCK-8 solution was added to each well. After 1–4 h incubation, cell viability was determined by measuring the absorbance at 450 nm using a Flex Station 3 microplate spectrophotometer (Molecular Devices, LLC, Sunnyvale, CA, USA).

For drug activity assays, 100 µl H9c2 cells/well were seeded into 96-well plates at a density of 3.0×104 cells/ml, and incubated overnight. Each plate contained 8 negative and 8 positive control wells, and all cells, excluding the positive controls, were treated with 100 µl/well H2O2 (50 µmol/l) for 3 h. Following H2O2 treatment, 0.1 µl/well dimethyl sulfoxide (DMSO; Sigma-Aldrich, St. Louis, MO, USA) was added to the positive control wells, and 0.1 µl/well TCM extract samples were added to the all other wells, excluding the negative controls. Cells were then incubated for an additional 3 h. Cell viabilities were tested using the CCK-8 assay kit to assess drug activities.

Lactate dehydrogenase (LDH) activity, malondialdehyde (MDA) content and superoxide dismutase (SOD) activity assays

LDH, MDA and SOD were measured using the respective assay kits (Beyotime Institute of Biotechnology, Haimen, China) according to the manufacturer's instructions. Briefly, 1,000 µl/well H9c2 cells were seeded into 24-well plates at a density of 3.0×104 cells/ml. Following a 3-h treatment with 50 µmol/l H2O2, and a 3-h incubation with 50 µmol/l quercetin and 25 µg/ml TCM extracts, the cells were centrifuged at 400 × g for 5 min, and 120 µl of the supernatant was then transferred to a 96-well plate for LDH activity determination. Subsequently, all cells were lysed and centrifuged at 1,600 × g for 10 min, the supernatants were then collected and stored at -80°C prior to MDA and SOD detection.

For LDH assays, 60 µl work ing solution, containing 20 µl lactic acid solution, 20 µl 1X INT solution and 20 µl enzyme solution, was added to each sample (total, 180 µl) for an additional 30 min incubation by gently agitating at room temperature. The maximum LDH release of target cells was determined by lysing target cells for 45 min and subsequently measuring the LDH from the culture medium. Absorbance values after the colorimetric reaction were measured at 490 nm with a reference wavelength of 655 nm, using a Flex Station 3 microplate spectrophotometer (Molecular Devices, LLC, Sunnyvale, CA, USA).

For MDA assays, 200 µl working solution was added to 100 µl samples for an additional 15 min incubation at 100°C, and were subsequently centrifuged at 1,000 × g for 10 min after the samples cooled to room temperature. Subsequently, 200 µl supernatant was transferred to a 96-well plate for MDA detection by Flex Station 3 microplate spectrophotometer at 532 nm absorbance with a reference wavelength of 450 nm.

For SOD assays, 180 µl working solution was added to 20 µl sample for an additional 30 min incubation at 37 °C. SOD activities were detected at 490 nm with a reference wavelength of 600 nm, using the same microplate reader as before.

Western blotting

The protein expression levels of the apop-totic proteins, caspase-3, B-cell CLL/lymphoma 2 (Bcl-2), Bcl-2-associated X protein (Bax), and the MAPK subfamily proteins, p38, JNK and ERK1/2, were detected by western blotting. A total of 1.5×106 cells/ml/well were seeded into 6-well plates. Following 3 h treatment with 12.5 µmol/l (for MAPK proteins) or 50 µmol/l (for apoptotic proteins) H2O2, cells were incubated with 25 µg/ml active extracts for 3 h. The cells were then rinsed twice with ice-cold 1X phosphate-buffered saline (PBS) and harvested under non-denaturing conditions by incubation at 4°C with lysis buffer (Qiagen, Hilden, Germany) containing protease and phosphorylase inhibitors for 10 min. The cell lysates were centrifuged at 14,000 × g for 10 min at 4°C to remove insoluble precipitates. The protein content in each sample was determined by Bradford assay (Applygen Technologies, Inc., Beijing, China). Total protein (50 µg) from cell culture samples was denatured and separated by SDS-PAGE on 12% acrylamide gels, and subsequently electrophoretically transferred to polyvinylidene fluoride membranes. Nonspecific binding sites were blocked by incubation of membranes in Tris-buffered saline with Tween (TBS-T) containing 5% bovine serum albumin (BSA; Amresco, LLC, Solon, OH, USA) for 2 h. The membranes were then incubated with primary antibodies against rabbit anti-caspase-3 (cat. no. sc-7148; Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), rabbit anti-Bcl-2 (cat. no. 2870S Cell Signaling Technology, Inc., Danvers, MA, USA), rabbit anti-Bax (cat. no. sc-526; Santa Cruz Biotechnology, Inc.), rabbit anti-p38 (cat. no. 9212; Cell Signaling Technology, Inc.), rabbit anti-phospho (p)-p38 (cat. no. 4631; Cell Signaling Technology, Inc.), rabbit-anti-JNK (cat. no. 9258; Cell Signaling Technology, Inc.), rabbit anti-p-JNK (cat. no. 4671; Cell Signaling Technology, Inc.), rabbit anti-ERK1/2 (cat. no. 9102S; Cell Signaling Technology, Inc.), rabbit anti-p-ERK1/2 (cat. no. 9101S; Cell Signaling Technology, Inc.) all at 1:1,000 dilution, overnight at 4°C. The membranes were washed with TBS-T and were subsequently incubated with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (cat. no. 7074; 1:2,000; Cell Signaling Technology, Inc.) for 2 h and visualized using a Chemi Doc XRS+ detection system (Bio-Rad Laboratories, Inc., Hercules, CA, USA). β-tubulin was used as a loading control.

Immunofluorescence assays for early growth response-1 (Egr-1)

H9c2 cells (100 µl/well) at a density of 5.0×104 cells/ml were seeded into 96-well plates. The cells were untreated (control) or incubated with 12.5 or 200 µmol/l H2O2 alone, or with 12.5 µmol/l H2O2 and 25 µg/ml active extracts for 2 h. Subsequently, the cells were fixed with 4% (v/v) formaldehyde (Amresco, LLC) in 1X PBS at room temperature for 15 min, then washed with 1X PBS 3 times, and blocked with 1% (w/v) BSA (Amresco, LLC) in 1X PBS containing 0.3% (v/v) Triton X-100 (Amresco, LLC) at room temperature for 30 min. The primary antibody against Egr-1 (cat. no. sc-110; 1:100; Santa Cruz Biotechnology, Inc.) was incubated with the cells at 37°C for 2 h. Subsequently, cells were washed with PBS and incubated with goat anti-rabbit IgG-CruzFluor 488 (cat. no. sc-362262; 1:250; Santa Cruz Biotechnology, Inc.) at 37°C for 1 h. Following 3 washes with 1X PBS, cell nuclei were stained using Hoechst 33258 (Sigma-Aldrich) at a final concentration of 2 µg/ml for 15 min. Fluorescent images were captured using a DMI 4000B fluorescence microscope (Leica Microsystems GmbH, Wetzlar, Germany).

The library of TCM extracts

Each TCM herb (500 g) was soaked in water for 1 h prior to extraction, and extraction was performed twice by boiling in 10- and 8-fold volumes of water (v/v) for 2 h. Extracts were filtered through gauze, and all crude extractions were combined and concentrated to 500 ml. The concentrates were then separated on macroporous resins (specification, Φ 5×60 cm; 1:1 weight ratio of the concentrates; HaiGuang Chemical Co., Ltd., China) by successive elution with water and different concentration gradients of ethanol (20–95%), with 3 bed volumes (BV) of eluent volume at a flow rate of 1 BV/h. Eight samples from each TCM herb were collected, and concentrated at 70°C. Following freeze drying, 5 mg of each sample was dissolved into 200 µl DMSO, then dispensed into 96-well plates.

Statistical analysis

All data are presented as the mean ± standard deviation (SD). Statistical analysis was performed using one-way analysis of variance and Tukey's post-hoc tests using SPSS software version 17.0 (SPSS, Inc., Chicago, IL, USA). P<0.05 was considered to indicate a statistically significant difference.

Results

Creation and optimization of oxidative damage cell model for HTS

To establish a stable HTS assay that generates reliable outcomes, the present study optimized several factors that may affect the assay results.

Sodium pyruvate-supplemented medium vs. sodium pyruvate-free medium

Sodium pyruvate, a supplement in cell culture medium, may affect the screening assays. As demonstrated in Fig. 1, when the H9c2 cells were maintained in DMEM containing 110 mg/l sodium pyruvate, 12.5–200 µmol/l H2O2 induced a decrease in cell viabilities (<7%; Fig. 1A and B). However, H2O2 treatment of the cells in sodium pyruvate-free DMEM resulted in a more marked decrease (~40%) in cell viability (Fig. 1C and D). Therefore, sodium pyruvate-free DMEM was used during the oxidative damage model.

Figure 1 Effect of different concentrations and incubation times of H2O2 on the viability of H9c2 cells. H9c2 cells (100 µl/well) at a density of 3.0×10 cells/µl were treated with different concentrations (0–400 µmol/l) of H2O2 at 37°C and 5% CO2 for 0–4 h. Cell viabilities were then analyzed using cell counting kit-8 assay with absorbance measured at 450 nm. H9c2 cells incubated in medium supplemented with 110 mg/ml sodium pyruvate were treated with (A) different concentrations of H2O2 for 4 h and (B) 50 µmol/l H2O2 for different time periods. H9c2 cells incubated in sodium pyruvate-free medium were treated with (C) different concentrations of H2O2 for 4 h and (D) 50 µmol/l H2O2 for different time periods. OD, optical density.

Concentration and incubation time

Optimization experiments were also performed to determine the optimal working concentration and incubation time of H2O2. H9c2 cells were exposed to varying degrees of oxidative stress by treatment with H2O2 for 0–4 h. The results demonstrated that H2O2 reduced cell viability in a dose- and time-dependent manner in the pyruvate-free groups, and exhibited an almost 40% injury at 50 µmol/l for 3 h. Higher concentrations or longer incubation time did not result in a more significant change to the OD450 values (Fig. 1C and D).

Cell density

A low number of cells per well may cause low response values, however, a large cell number is not conducive to cell growth, due to the contact inhibition. Thus, determining the appropriate number of cells is essential for drug screening. As demonstrated in Fig. 2, 1.5–4.0×104 cells/well treated with 50 µmol/l H2O2 for 3 h exhibited consistent results, whereas higher seeding densities exhibited reduced cell viability. Thus, the current study used the cell density of 3.0×104 cells/well for the HTS assays.

Figure 2 Effects of cell density on the viability of H2O2-treated H9c2 cells. H9c2 cells (100 µl/well) at the densities of 1.5, 3.0, 4.0, 5.0, 6.0, 8.0 and 10×104 cells/ml were seeded into 96-well plates. The cells were treated with 50 µmol/l H2O2 at 37°C and 5% CO2 for 3 h, and cell viabilities were then assessed using cell counting kit-8 assay, measuring absorbance at 450 nm. OD, optical density.

Variability and robustness of model

To assess whether the model of oxidative damage can be applied to an HTS format, the present study applied the optimized conditions to establish the H2O2-induced cell damage model. The data of the cell viabili-ties from 30 wells of positive control (H2O2-free) and 30 wells of negative control (H2O2-treated) were obtained to analyze variability between wells and the robustness of the cell model of oxidative stress using the Z′ factor, which is calculated from the following formula: Z′=1-[3×(δc+ -δc−)⁄(µc+ - µc−)]; δc+ =SD of positive control; δc− =SD of negative control; µc+= mean of positive control; and µc− =mean of negative control. Z′≥0.5 indicates the assay method can be performed effectively in HTS (25). As presented in Fig. 3, treatment of H9c2 cells with 50 µmol/l H2O2 for 3 h produced a Z′ value of ~0.85 and small critical values (CVs) (0.7% for positive control and 1.5% for negative control). These results demonstrated that there was an appropriate separation between the SDs of the H2O2-induced cell damage model and the untreated controls.

Figure 3 Testing the reproducibility of the oxidative damage model in an automated 96-well assay format. H9c2 cells (100 µl/well) at 3.0×104 cells/µl were seeded into 96-well plates. H2O2-free wells (30 wells) and H2O2-treated wells (30 wells) were used as positive and negative controls, respectively. Following treatment with 50 µmol/l H2O2 at 37°C and 5% CO2 for 3 h, cell viabilities were assessed using CCK-8 assay by measuring absorbance at 450 nm. The data are presented as (A) CCK-8 readings from individual wells and (B) as the mean ± standard deviation of 30 wells. *P<0.001 vs. H2O2-free wells. OD, optical density; CCK-8, cell counting kit-8.

The incubation time of the TCM extracts may be an important factor that affects the result of the assay. A positive control drug, quercetin, was used to optimize the incubation time of the cells with the drug following oxidative damage. Following 3 h treatment with 50 µmol/l H2O2, 25 µmol/l quercetin was added to H9c2 cells and incubated for an additional 1–6 h. The results demonstrated that the optimal drug incubation time following oxidative damage was 3 h (Fig. 4A).

Figure 4 Optimization of the incubation time of H2O2-treated cells with samples and evaluation of the robustness of the screening method in a high-throughput format. H9c2 cells (100 µl/well) at a density of 3.0×104 cells/µl were plated into 96-well plates. H9c2 cells were treated with 50 µmol/l H2O2 for 3 h (model group) or incubated with 25 µmol/l quercetin for a further 1–6 h. Control group was treated with dimethyl sulfoxide only. The cell viabilities were then assessed using cell counting kit-8 assay, measuring absorbance at 450 nm. (A) The data are presented as the mean ± standard deviation of 30 wells at the different time points, and (B) as individual wells following the 3-h treatment with 50 µmol/l H2O2 and 25 µmol/l quercetin for an additional 3 h. Data from 30 wells of positive controls (H2O2 + quercetin) and 30 wells of negative controls (H2O2 + vehicle) were used for the calculation of Z' factor and critical value. OD, optical density.

The optimized incubation time was used to establish the screening assays, and Z′ factor calculation validated whether the assays were suitable for an HTS format. Quercetin-treated cells (pretreated with 50 µmol/l H2O2 for 3 h) were used as the positive control and the cells treated with H2O2 only as the negative control to further investigate the robustness of the screening assays in an HTS format. As demonstrated in Fig. 4B, quercetin increased the cell viabilities compared with 50 µmol/l H2O2 treatment. The Z′ factor was 0.71, and the CVs of the positive and negative controls were 0.9 and 1.5%, respectively. This indicated that the H2O2-induced cell damage model was suitable for HTS assays.

Identification of active extracts from TCM

The optimized 96-well plate HTS system was then used to identify extracts with antioxidative activity from a TCM library. After a 3-h treatment with 50 µmol/l H2O2, and a 3-h incubation with TCM extracts at 37°C and 5% CO2, the cell viabilities were determined using the CCK-8 kit and absorbance measured at 450 nm. In addition to the extracts, 0.1 µl/well 25 mg/ml DMSO solution was added to the cells. The final concentration of the samples in each well was ~25 µg/ml. This concentration exhibited low cytotoxic effects on the cells (data not shown). The extracts that exhibited an OD450 value >the mean ± 3SD of the negative control (H2O2-treated) were considered as potential active samples. The top 17 hits from the primary screening were selected for further validation (Fig. 5).

Figure 5 Identification of hits from the traditional Chinese medicine extracts library in HTS assay format. Assays were performed in the optimized automated HTS format. Representative values of the absorbance at 450 nm of 140 samples are presented. The red line indicates the mean of the negative control, and the blue line indicates the mean of negative control ± 3 standard deviations. Samples that exhibited 450 nm absorbance values above the blue line were considered to be hits. HTS, high-throughput screening.

Validation of the primary hits

The increased OD450 values observed in the HTS assay may be due to increased cell viabilities via protection of cells from oxidative damage, reduced H2O2 toxicity by a direct reaction with H2O2 in the culture medium, or extract compounds themselves may absorb at 450 nm. In order to exclude false positives, the current study further validated the primary hits. To determine whether the extract samples absorbed at 450 nm, 0.1 µl primary hit extracts were directly added to the wells of a 96-well plate containing 100 µl pyruvate-free DMEM (no cells), and incubated for 3 h. The results demonstrated that two extracts KY-0520 and KY-0538 had no significant absorption at 450 nm (Fig. 6Aa). In order to test whether H2O2 directly reacts with the extracts in the culture medium, cells were pretreated with 50 µmol/l H2O2 for 3 h, then the culture medium was replaced with pyruvate-free DMEM containing 25 µg/ml primary hit and incubated for additional 3 h. The results indicated that KY-0520 and KY-0538 increased cell viability compared with the H2O2-only treatment when in the culture media with H2O2, and when H2O2/KY-0520 and KY-0538 treatments were performed individually (Fig. 6Ab and c). Taken together, the results indicate that activities of KY-0520 and KY-0538 were due to antioxidative properties. The present study additionally measured the cell viability following treatment with KY-0520 and KY-0538 at different concentrations. H9c2 cell viability was increased by KY-0520 and KY-0538 in a concentration-dependent manner. The concentration-response curves of KY-0520 and KY-0538 demonstrated that the compounds are active between of 0.78 and 100 µg/ml (Fig. 6B), and the EC50 values of KY-0520 and KY-0538 were 11.43 and 19.59 µg/ml, respectively.

Figure 6 Verification of the activities of the primary hits. The two hits KY-0502 and KY-0538 were further confirmed as candidates in the secondary screening assays. (A) To exclude false positive hits, (a) the primary hit molecules were added to the wells only containing sodium pyruvate-free medium without cells; (b) H9c2 cells (100 µl/well) at 3.0×104 cells/ml were plated into 96-well plates, cells were pretreated with 50 µmol/l H2O2 for 3 h, then the medium was replaced with sodium pyruvate-free medium containing 25 µg/ml primary hits, and incubated for an additional 3 h; (c) cells were treated with 50 µmol/l H2O2 for 3 h, 25 µg/ml primary hits were then directly added to wells followed by a 3-h incubation. *P<0.0001 vs. model group. (B) The concentration-dependent effects of KY-0502 and KY-0538 on the viability of H9c2 cells was measured. H9c2 cells (100 µl/well) at 3.0×104 cells/ml were plated into 96-well plates, and treated with 50 µmol/l H2O2 for 3 h. Subsequently, the cells were incubated with different concentrations of KY-0502 and KY-0538 for an additional 3 h, and cell viabilities were tested at 450 nm. Data are presented as the mean ± standard deviation of triplicate measurements. OD, optical density.

Characterization of the cardioprotective activities of KY-0520 and KY-0538

The present study further investigated the antioxidant activity of the TCM extracts. The results demonstrated that H2O2-induced oxidative damage to H9c2 cells significantly increased LDH activity (P<0.0001; Fig. 7A) and MDA levels (P<0.001; Fig. 7B), and decreased SOD activity (P<0.001; Fig. 7C) compared with the control. KY-0520 and KY-0538 treatment (25 µg/ml) significantly reduced the LDH activity (P<0.001; Fig. 7A) and MDA levels (P<0.01; Fig. 7B) compared with H2O2-treated cells. Additionally, KY-0520 and KY-0538 significantly increased the SOD activity levels compared with H2O2-treated cells (P<0.05 and P<0.001, respectively; Fig. 7C). These results suggest that KY-0520 and KY-053 prevent the accumulation of free radicals and attenuate cardiomyocyte damage induced by H2O2.

Figure 7 Effects of KY-0502 and KY-0538 on LDH levels, MDA levels and SOD activity in H2O2-treated H9c2 cells. H9c2 cells (1,000 µl/well) at the density of 3.0×104 cells/ml were seeded into 24-well plates and randomly divided into five groups. The cells were treated with dimethyl sulfoxide (control group), 50 µmol/l H2O2 (model group), 50 µmol/l H2O2 + 50 µmol/l quercetin (quercetin group), 25 µg/ml KY-0502 (KY-0502 group) or 25 µg/ml KY-0538 (KY-0538 group). Subsequently, the cells and medium were collected for LDH, MDA and SOD detection using the corresponding assay kits. (A) LDH contents released from the cells, (B) MDA contents of the cells and (C) SOD activities of the cells. *P<0.05, **P<0.01, ***P<0.001 and ΔP<0.0001, comparisons indicated by brackets. LDH, lactate dehydrogenase; MDA, malondialdehyde; SOD, superoxide dismutase.

TCM extracts decrease H2O2-induced apoptosis in H9c2 cells

Based on the cell viability and antioxidative results, the current study further examined whether KY-0520 and KY-0538 exhibited protective effects against H2O2-induced cell apoptosis by western blot analysis of apoptosis-associated proteins (Fig. 8A). As demonstrated in Fig. 8B and C, H2O2 treatment significantly increased the protein expression levels of cleaved caspase-3 (1.31-fold increase; P<0.05;) and Bax (3.19-fold increase; P<0.01) compared with the untreated controls. KY-0520 and KY-0538 (25 µg/ml) significantly decreased the protein expression levels of cleaved caspase-3 (P<0.05) and Bax (P<0.01) in H9c2 cells compared with H2O2 treatment alone. Additionally, the protein levels of Bcl-2, an anti-apoptotic protein, were decreased following H2O2 treatment compared with untreated controls (P<0.05), however, compared with H2O2 treatment alone, Bcl-2 levels were significantly increased following KY-0520 and KY-0538 treatment (P<0.05; Fig. 8D). The results indicated that the extracts inhibited apoptosis by regulation of pro- and anti-apoptotic proteins in H2O2-treated H9c2 cells.

Figure 8 Effect of traditional Chinese medicine extracts on H2O2-induced apoptosis in H9c2 cells. Cells were stimulated with 50 µmol/l H2O2 for 3 h (model group) or treated with 25 µg/ml KY-0502 and KY-0538 for an additional 3 h. Control group were treated with DMSO only. After the cells were lysed, total proteins (50 µg) were separated on 12% SDS-PAGE and detected by immunoblotting. (A) Western blot analysis of apoptotic proteins and quantification of (B) Bcl-2, (C) Bax and (D) cleaved caspase-3 protein expression levels, β-tubulin was used for normalization. Western blot images are representative of three independent experiments and data are presented as the mean ± standard deviation (n=3). *P<0.05, **P<0.01 vs. control group; ΔP<0.05, ΔΔP<0.01 vs. model group. Bcl-2, B-cell CLL/lymphoma 2; Bax, Bcl-2-associated X protein; C-caspase, cleaved caspase.

TCM extracts inhibit Egr-1 protein accumulation in nucleus in H2O2-exposed H9c2 cells

It was previously demonstrated that following exposure of H9c2 cells to 200 µmol/l H2O2, Egr-1 is translocated from the cytoplasm and accumulates in the nucleus (26). The present study treated H9c2 cells with 200 µmol/l H2O2 for 2 h, however, Egr-1 did not translocate from the cytoplasm to nucleus, it accumulated in the cytoplasm and nuclear membrane (Fig. 9A). By contrast, at an H2O2 concentration of 12.5 µmol/l, Egr-1 nuclear staining was increased (Fig. 9B). KY-0520 and KY-0538 (25 µg/ml) markedly decreased Egr-1 immunostaining to near basal levels, and caused Egr-1 redistribution in the nucleus and cytoplasm (Fig. 9B). These results suggested that KY-0520 and KY-0538 may protect H9c2 cells from oxidative damage by regulating Egr-1 activity.

Figure 9 Cellular localization of Egr-1 in H9c2 cells exposed to different concentrations of H2O2 and traditional Chinese medicine extracts. Cells were subjected to immunocytochemical analysis with an antibody directed against total Egr-1 (green). To reveal nuclear morphology, the nuclei were stained with Hoechst 33258 (blue). The images were captured using a fluorescence microscope. Representative images are indicative of ≥3 independent experiments. (A) Cells were untreated (control) or incubated with 200 µmol/l H2O2 for 2 h. (B) Cells were untreated (control) or incubated with either 12.5 µmol/l H2O2 or 12.5 µmol/l H2O2 followed by treatment with 25 µg/ml KY-0502 or KY-0538 for 2 h. Egr-1, early growth response-1.

TCM extracts inhibit ERK1/2 and p38-MAPK in H2O2-exposed H9c2 cells

The MAPK signaling pathways, including ERK1/2, p38 and JNK-MAPK, are critical for the regulation of apoptosis and other cellular processes. Activation of the MAPK signaling pathways is a characteristic feature of oxidant-induced apoptosis (27), and is well established in cardiac myocytes (28). In the present study, the protein levels of p-ERK1/2, p-JNK and p-p38 MAPK were measured in H9c2 cells exposed to 12.5 µmol/l H2O2, and the results are presented in Fig. 10. Western blot analysis demonstrated that the phosphorylation levels of p42- and p44-ERK and p38 MAPK were significantly increased by H2O2 (2.04-, 1.37- and 1.88-fold, respectively) compared with untreated control cells (Fig. 10B and C). However the increased phosphorylation of those kinases was reversed by 25 µg/ml KY-0520 and KY-0538 after 3 h incubation. However, no significant alterations in JNK phosphorylation were observed following 12.5 µmol/l H2O2 or 25 µg/ml active extracts treatment (Fig. 10D). These results indicate that KY-0520 and KY-0538 may regulate the MAPK signaling pathway to protect against H2O2-induced oxidative stress in H9c2 cells.

Figure 10 Effects of traditional Chinese medicine extracts on ERK1/2, p38 and JNK phosphorylation in H2O2-treated H9c2 cells. Cells were stimulated with 12.5 mol/l H2O2 for 3 h (model group) or treated with 25 µg/ml KY-0502/KY-0538 for an additional 3 h. Control cells were treated with dimethyl sulfoxide only. The cells were lysed, total proteins (50 µg) were separated on 12% SDS-PAGE and detected by immunoblotting. (A) Western blot analysis and densitometry measuring (B) ERK1/2, (C) P38 MAPK and (D) JNK phosphorylation levels. β-Tubulin was used for normalization. Western blot images are representative of three independent experiments. Data are presented as the mean ± standard deviation (n=3). *P<0.05 and **P<0.01 vs. control group; ΔP<0.05 and ΔΔP<0.01 vs. model group. ERK, mitogen-activated protein kinase 3; p, phospho; P38, mitogen-activated protein kinase 14; JNK, mitogen-activated protein kinase 8.

Discussion

Increasingly, studies indicate that ROS are associated with the pathogenesis and progression of various cardiovascular diseases. Sensitivity to oxidative stress is greater in the heart compared with other organs due to lower levels of antioxidant enzymes (29). The pathogenesis of cardiac hypertrophy, developed by chronic hypertrophy, is associated with ROS via regulation of the intracellular pathways linked to MAPKs and phosphatidylinositol-4,5-bisphosphate 3-kinase/Akt (30–32). H2O2 is predominantly produced via the dismutation of superoxide anions, it can also swiftly permeate the cell membrane and react with intracellular metal ions to form toxic hydroxyl radicals, which cause DNA damage (33). Previous studies demonstrated that H2O2 is excessively produced during cardiomyocyte apoptosis, leading to caspase-3 activation via mitochondrial dysfunction and cytosolic release of mitochondrial cytochrome c (34).

Numerous studies have investigated natural plant compounds and TCM extracts for their antioxidant activities. Silibinin (the major active component of silymarin extracted from S. marianum) has been demonstrated to have antioxidative, antitumor and anti-inflammatory properties (35). The volatile oil of Nardostachyos Radix et Rhizoma (the root and rhizome of Nardostachys jatamansi DC.) was reported to markedly suppress ROS formation and dose-dependently increase glutathione levels in H9c2 cells following oxidative injury (36). Thus, novel antioxidant agents from natural plants and TCM may be useful for the treatment of cardiac diseases.

Using H2O2 to treat H9c2 rat myocardial cells, the present study established a cell model of oxidative damage for HTS assay, and used the model to identify cardioprotective agents from a library of TCM extracts. The actions of the extract were determined by CCK-8 assay, which is based on dehydrogenase activity detection in viable cells, and is widely used for cell proliferation and cytotoxicity assays. The CCK-8 assay does not require washing or cell lysis, therefore, variability is mini-mized. It has previously been successfully applied in HTS studies as it is inexpensive and easy to operate (37). Therefore, the present study used the CCK-8 assay to evaluate the effect of TCM extracts on the viability of oxidatively damaged cells. Two hits, KY-0520 and KY-0538, were further validated as cardioprotective agents, and attenuated oxidative damage in a concentration-dependent manner (EC50 values, ~11.43 and 19.59 µg/ml, respectively).

The present study used 50 µmol/l H2O2 to induce oxida-tive damage in the model. Various studies have investigated the appropriate working concentration of H2O2, however, results have varied (21,38,39). Sodium pyruvate is commonly supplemented in culture media, however, this compound can nonenzymatically react with H2O2, leading to liberation of CO2, and the conversion of α-keto acid to carboxylic acid (40,41). Therefore, the present study used H2O2 diluted with pyruvate-free DMEM to induce oxidative damage in our cell-based assays.

To establish the HTS assay model, two Z′ factors were required to evaluate the screening method. One was used to evaluate the robustness of the cell model, which indicates whether the cell damage model was successfully established and suitable for HTS. The other Z′ factor was used to evaluate the robustness of the extract screening assay (Figs. 3 and 4). The majority of HTS assays are based on specific targets. Compared with the HTS assays designed to screen for drugs acting on specific targets, the cell damage model has advantages and limitations. Cell-based screening can directly evaluate the protective activities of drugs by measuring cell viability, however, the direct targets of the drugs and the signaling pathways involved are unclear. To understand the mechanisms of action of the drugs, it is necessary to further investigate the potential targets and signaling pathways. The effects of the drug candidates identified in the present study may be mediated by interaction with multiple targets and signaling pathways. Therefore, the cell-protective functions of KY-0520 and KY-0538 may be mediated by their antioxidant activity, and also via interaction with other pathways. Other factors and pathways associated with the effects of KY-0520 and KY-0538 may include Egr-1. Immunofluorescence demonstrated the localization of Egr-1 to be altered by KY-0520 and KY-0538 treatment under H2O2-induced oxidative stress.

H2O2 is a strong oxidant that markedly decreases cell viability and increases apoptosis. The present study measured LDH activity to further investigate the cardioprotective effect of KY-0520 and KY-0538. LDH assays are widely used to quantify the level of LDH release. MDA levels and SOD activity were also measured as indicators of oxidative damage and myocardial function. KY-0520 and KY-0538 demonstrated significant antioxidant activities and protective effects on cardiomyocytes in vitro (Fig. 7). Furthermore, previous studies have demonstrated that H2O2 can decrease the Bcl-2/Bax ratio and increase the level of cleaved caspase-3, therefore inducing apoptosis (42,43). Western blot analysis demonstrated that KY-0520 and KY-0538 regulate the Bcl-2/Bax ratio in H2O2-exposed H9c2 cells, and decrease the H2O2-induced cleaved caspase-3 activation (Fig. 8). These effects may contribute to the antioxidant activity of KY-0520 and KY-0538 and their protection against oxidative stress.

Egr-1 is a transcription factor encoded by an immediate early gene (44). Egr-1 is weakly expressed under normal conditions, and its expression is activated by various environmental stimuli associated with injury and stress, including growth factors, cytokines, T cell receptor ligation, hormones, thrombin, shear stress and mechanical forces, neurotransmitters, ultraviolet light, ROS, ischemia/reperfusion and hypoxia (45–52). Egr-1 mRNA is expressed following cardioplegic arrest and reperfusion in human hearts, and in rat hearts subjected to cold cardioplegia for 40 min followed by 40 min reperfusion. The expression of Egr-1 and the downstream effects on transcription are tightly controlled, and cell-specific upregulation induced by processes such as hypoxia and ischemia, has been previously linked to multiple aspects of cardiovascular injury. Egr-1 regulates cell growth and proliferation (53,54), and positively modulates inflammation, thrombosis and apoptosis, by direct and indirect mechanisms (45,55). It was previously reported that targeting rodent Egr-1 selectively reduced the infarct size following myocardial ischemia/reperfusion. The mechanisms reported to be involved were associated with the attenuation of intercellular adhesion molecule 1-dependent inflammation and inhibition of other Egr-1-dependent molecules, including tumor necrosis factor-α (TNF-α), vascular cell adhesion molecule-1 (VCAM-1), tissue factor (TF), plasminogen activator inhibitor type 1, and p53. Inhibition of functional TF, VCAM-1 and TNF-α has been previously demonstrated to reduce the infarct size in experimental models of myocardial ischemia/reperfusion. The transcription of the pro-apoptotic factor, p53, is inhibited by Egr-1, with an associated reduction in apoptosis and infarct size (56). A previous study demonstrated that Egr-1 represses transcription from the calsequestrin (CSQ) promoter, resulting in reduced expression of CSQ, which is a major calcium storage protein critical for normal cardiac function (57). Additionally, overexpression of Egr-1 directly induced caspase activation and apoptosis in human cardiac fibroblast cultures in vitro (58). These studies indicate that inhibition of Egr-1 activity may be cardioprotective.

The findings of the present study were consistent with a previous study that demonstrated that H2O2 induces the translocation of Egr-1 from the cytoplasm to nucleus, thus promoting the accumulation of Egr-1 in the nuclei of H9c2 cells (Fig. 9B) (26). However, in contrast to the previous study, a high concentration of H2O2 (200 µmol/l) resulted in high levels of Egr-1 in the cytoplasm and nuclear membrane, rather than accumulation in the nucleus (Fig. 9) (26). The different regulatory mechanisms of Egr-1 under different concentrations of H2O2 are not clear. KY-0520 and KY-0538 effectively reversed the translocation of Egr-1 from the cytoplasm to the nucleus induced by 12.5 µmol/l H2O2 (Fig. 9). The cardio-protective activities of KY-0520 and KY-0538 appear to be associated with inhibition of Egr-1 activity and ROS scavenging.

Egr-1 expression is upregulated in response to cardiac ischemia/reperfusion stress (59). Additionally, a previous study demonstrated that Egr-1 mRNA expression in H9c2 cells was upregulated by H2O2 in vitro, and that the upregulation was dependent on MEK/ERK and JNK signaling (26,60). ERK1/2 is a component of the classical MAPK pathway that was previously demonstrated to be directly activated by high levels of ROS (including xanthine oxidase-derived H2O2) leading to transcription of Egr-1 (60–62). Thus, the present study investigated whether these pathways are modified during H2O2-induced oxidative stress. Consistent with previous studies (26,63), the western blot analysis of the present study indicated that the phosphorylation levels of ERK1/2 and p38-MAPK kinase were increased following 12.5 µmol/l H2O2 treatment (Fig. 10). Therefore, it is speculated that ERK1/2 and p38-MAPK may be important upstream regulators that mediated Egr-1 modulation during cardiomyocyte oxidative stress. The results of the present study indicated that the antioxidative effects of KY-0520 and KY-0538 may be mediated by suppression of the ERK1/2, p38-MAPK/Egr-1 signaling pathways in H2O2-induced oxidative stress.

In summary, the hits from the HTS assays may generate novel drugs that have the potential to be used as therapeutics for cardiomyocyte diseases, including heart ischemia/reperfusion, cardiac hypertrophy, cardiac dysfunction and heart failure. The present study established and validated a H2O2-induced cell damage model for use in HTS, however, further mechanistic research is required to understand the effects of the identified hits. Further investigation of the activity of the hits will be performed using primary cardiomyocyte cells or appropriate animal models.

Acknowledgments

The authors of the present study thank the Natural Products Research Department, State Key Laboratory of New-tech for the Chinese Medicine Pharmaceutical Process for providing the TCM extracts library. The current work was supported by the Ministry of Science and Technology of China (no. 2013zx0942203).

References