Eupatilin was provided by Dong-A Pharmaceutical Co. Ltd (Cheoin-gu, South Korea). Antibodies against PARP, Bax, Bcl-2 and cytochrome c were purchased from Santa Cruz Biotechnology, Inc. (Dallas, TX, USA). Caspase inhibitors Z-DEVE-FMK, Z-IETD-FMK and Z-LEHD-FMK for caspase-3, caspase-8 and caspase-9 were obtained from R&D Systems (Minneapolis, MP, USA). These inhibitors were dissolved in dimethyl sulfoxide (DMSO; Xi'an Chemical Co., Ltd., Boaji, China) and diluted prior to use in cell culture. The present study was approved by the ethics committee board of Mudanjiang Medical University (Mudanjiang, China). The remaining reagents and solvents used were of analytical grade and purchased from Xi'an Chemical Co., Ltd.

The U-2 OS human OS cells were purchased from the Cell Bank of Shanghai Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences (Shanghai, China). The cells were maintained in RPMI-1640 medium supplemented with fetal calf serum (FCS; 10%), glutamine (2 mM), penicillin (100 U/ml) and streptomycin (10 µg/ml) in a humidified atmosphere at 37°C with 5% CO₂.

The viability of U-2 OS cells was determined using an MTT assay. U-2 OS cells were seeded at a density of 5×10⁴ cells/well in 96-well culture plates. Each well contained 100 µl medium supplemented with 10% FCS, and the cells were incubated at 37°C for 24 h prior to treatment with various concentrations of eupatilin. Following 24 h of incubation, the medium was removed and replaced with 10% FCS containing various concentrations of eupatilin (0, 50, 100, 200 and 400 µg/ml) and incubated at 37°C for 96 h. Subsequently, 50 µl MTT solution was added to each well and incubated for a further 4 h as described previously (22). The formazan crystals obtained were dissolved in 100 µl DMSO and the absorbance was recorded using an ELISA microplate reader (model 550; Bio-Rad, Laboratories, Inc., Hercules, CA, USA) at a wavelength of 570 nm.

Cell apoptosis was detected using Annexin V-FITC/PI double staining. In brief, 1×10⁵ untreated (control) or treated cells were harvested following trypsinization and centrifugation (500 × g for 5 min). The cells were then washed twice with ice-cold phosphate-buffered saline (PBS) and stained with an Annexin V-FITC/PI apoptosis kit (BD Pharmingen, San Diego, CA, USA), according to the manufacturer's instructions. Subsequently, the samples were analyzed using the FACS Calibur flow cytometer (BD Biosciences, Franklin Lakes, NJ, USA). Double-staining was used for the quantification of the apoptosis of U-2 OS cells following treatment with eupatilin, by measurement of phosphatidylserine expression on the outer surface of the plasma membrane identified by Annexin V-FITC binding. The results of the assay were analyzed with the exclusion of PI, the plasma membrane integrity probe (23).

Eupatilin-treated cells were harvested using 0.25% trypsin and then washed with PBS twice prior to fixation with 70% ethanol for ~30 min at 4°C. The cells were pelleted by centrifuging at 500 × g for 5 min and resuspended in 1 ml PBS containing 100 µl RNase (10 mg/ml) and 100 µl PI (0.5 mg/ml) for ~20 min at 37°C for cytoplasmic or nuclear DNA staining. The stained cells were then analyzed for DNA content using the FACS Calibur as previously described (24). Annexin V- and PI-positive cells were considered to be necrotic type cells, whereas Annexin-V positive and PI-negative cells were considered to be apoptotic cells.

The ΔΨm of the U-2 OS cells was analyzed by flow cytometry, using rhodamine 123 (Rh123), a fluorescent dye which has been shown to be selectively accumulated within the mitochondria of live cells (25). In brief, the eupatilin-treated and untreated cells were incubated with Rh123 dye (1 mg/ml in DMSO) for 30 min at 37°C in 5% CO₂. Subsequently, the cells were washed twice with PBS, resuspended in PBS, stained with 2 µg/ml PI and immediately subjected to flow cytometric analysis. The loss of ΔΨm was calculated as a percentage using CellQuest™ software (version 5.1; BD Biosciences).

The activities of caspase-3, -8 and -9 were evaluated using a caspase colorimetric assay kit (BioTek Instruments, Inc., Winooski, VT, USA) according to the manufacturer's instructions. U-2 OS cells were seeded in 12-well culture plates at a density of 2×10⁵ cells/well prior to incubation with eupatilin for 48 h. The cells were then harvested, lysed with lysis buffer for ~5 min in an ice bath and then centrifuged at 10,000 × g for 10 min. Subsequently, the reaction buffer was added to the supernatant solutions containing the proteins (100 µg). Caspase-3, -8 and -9 colorimetric substrates (5 µl each) were then added for 2 h at 37°C in a CO₂ incubator, prior to quantification of the optical density (OD) of the mixture at 405 nm using a spectrophotometer. Their respective activities were expressed relative to the theoretical OD value, calculated using the Sigma-Aldrich Caspase 9 Assay Kit, Colorimetric Technical Bulletin (www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/Bulletin/casp8cbul.pdf).

Cells were subjected to eupatilin treatment, and cell proteins were subsequently obtained by incubation for 1 h in 200 µl lysis buffer containing NaCl (300 mM), Tris HCl (50 mM; pH 7.6), Triton X-100 (0.5%), phenylmethanesulfonyl fluoride (2 mM), aprotinin (2 µl/ml) and leupeptin (2 µl/ml) at 4°C. A bicinchoninic acid protein assay kit (Pierce Biotechnology, Inc.; Thermo Fisher Scientific, Rockford, IL, USA) was used to quantify the protein expression levels, according to the manufacturer's instructions. Equal quantities of each protein (20 µg) were separated by 12% SDS-PAGE and then electrotransferred onto polyvinylidene difluoride membranes. The membranes were then incubated for 1 h with PBS containing 5% non-fat milk as a blocking solution at room temperature, prior to incubation with polyclonal rabbit anti-mouse poly(ADP-ribose)polymerase (PARP; 1:1,000 dilution; cat no. 9542), B cell lymphoma-2 (1:1,000 dilution; cat no. 2876), Bcl-2-like protein 4 (Bax; 1:1,000 dilution; cat no. 2772) and cytochrome c (1:1,000 dilution; cat no. 4272) antibodies (all from Cell Signaling Technology, Inc., Danvers, MA, USA) diluted with blocking solution at 4°C overnight. The membranes were subsequently incubated with horseradish peroxidase-conjugated rabbit anti-mouse secondary antibody (1:2,000–5,000) for ~2 h at room temperature. The developed immunoblots were then visualized with an enhanced chemiluminescence detecting system (GE Healthcare Life Sciences, Chalfont, UK). Expression was analyzed relatively, based on the ratio between the target protein and that of polyclonal rabbit β-actin (1:1,000 dilution; cat no. 4967; Cell Signaling Technology, Inc.).

All the quantitative results obtained are presented as the mean ± standard deviation. Student's t-test was used to calculate statistical differences between the control and eupatilin-treated cells, using GraphPad Prism 3.03 software (GraphPad Software, Inc., La Jolla, CA, USA) and P<0.05 was considered to indicate a statistically significant difference. All the studies were performed at least in triplicate.

The effects of eupatilin on U-2 OS cell proliferation were evaluated by MTT assay. The cells were treated with various concentrations of eupatilin (0, 50, 100, 200 and 400 µg/ml) for various time-periods (0, 24, 48, 72 and 96 h) and cell proliferation under these conditions was evaluated. The results revealed a dose-dependent inhibitory effect of eupatilin on U-2 OS cell proliferation 24 h post-treatment (Fig. 1). The inhibitory effect of eupatilin was demonstrated to be significant at a concentration of 100 µg/ml, whereas the most marked inhibition was observed in cells which were treated with eupatilin at concentrations of 200 and 400 µg/ml.

These results confirmed the antiproliferative effect of eupatilin on U-2 OS cells, and in order to elucidate the mechanism underlying this effect, eupatilin at a concentration of 100 µg/ml was selected for use in the subsequent experiments.

To ascertain whether eupatilin inhibited U-2 OS cell proliferation via induction of apoptosis, the cells were treated with eupatilin and then subjected to flow cytometric analysis. The results revealed a significant increase in Annexin V-FITC cell binding following 100 µg/ml eupatilin treatment, compared with that of the control cells, indicating the initiation of apoptosis. Fig. 2A indicates that the apoptotic cell proportion was significantly increased from 3.23% in untreated U-2 OS cells to 49.75% in eupatilin-treated U-2 OS cells. These results confirmed that the cell death observed in U-2 OS cells following eupatilin treatment occurred via the induction of apoptosis. In addition, flow cytometric analysis revealed that eupatilin treatment for 24 h significantly reduced the proportion of cells in S phase, compared with that of the control cells (Fig. 2B). Furthermore, elevated accumulation of cells in the G2/M apoptotic phase was observed, with respect to control cells, while the cell population in G1 phase demonstrated a slight decrease. These results revealed that the eupatilin responses of U-2 OS cells are correlated with apoptotic cell death.

In order to confirm whether mitochondrial dysfunction was involved in eupatilin-induced apoptosis, changes in ΔΨm were evaluated using Rh123 fluorescent dye. Rh123 was used as it is able to cross the mitochondrial membrane and thus accumulates in the mitochondrial matrix. However, this accumulation only occurs when the transmembrane potential is maintained (26). As indicated in Fig. 3A, following exposure of U-2 OS cells to 100 µg/ml eupatilin for 24 h, a significant reduction in ΔΨm was observed. The release of cytochrome c from the mitochondria to the cytosol is typically associated with ΔΨm depolarization (27). In addition, cytochrome c has been demonstrated to have a vital role in apoptosis (28,29). Therefore, the expression of cytochrome c was analyzed by western blotting. Fig. 3B indicates an increase in cytosolic cytochrome c and a decrease in mitochondrial cytochrome c in eupatilin-treated cells, when compared with control cells. These results indicated that eupatilin-induced apoptosis involves mitochondrial dysfunction, associated with a loss of ΔΨm and cytosolic cytochrome c release.

Subsequently, the effects of eupatilin on caspase-3, -8 and -9 activation and PARP cleavage were evaluated, in order to determine the death receptor involvement and mitochondrial pathways in eupatilin-induced apoptosis. The results revealed that eupatilin treatment of U-2 OS cells for 24 h induced activation of caspase-3 and -9, but not caspase-8 (Fig. 4A). Furthermore, as indicated in Fig. 4B, an increase in PARP cleavage was detected following eupatilin treatment. To further investigate the role of caspase activation in eupatilin-induced apoptosis, the effects of caspase inhibitors z-DEVE-FMK, Z-IETD-FMK, and Z-LEHD-FMK for caspase-3, -8 and -9 on apoptosis in U-2 OS cells were evaluated. Significant inhibition of eupatilin-induced apoptosis was observed following pretreatment with caspase-3 and -9 inhibitors, whereas the levels of early apoptosis remained unchanged with respect to caspase-8 inhibition (data not shown). These observations suggested that eupatilin induced caspase-dependent apoptosis in U-2 OS cells via a mitochondrial-dependent pathway.

The effects of eupatilin on Bax and Bcl-2 expression in U-2 OS cells were subsequently examined via analysis of Bax and Bcl-2 protein expression levels following 24 h of treatment. Western blot analysis indicated a marked increase in Bax expression levels in eupatilin-treated cells, whereas a significant decrease was observed in Bcl-2 expression (Fig. 5). This high ratio of Bax/Bcl-2 may contribute to the induction of apoptosis by eupatilin via the mitochondrial-dependent pathway.