Human renal cancer cell lines (786-O and Caki-1) were purchased from Bioresource Collection and Research Center (BCRC; Hsinchu, Taiwan) and American Type Culture Collection (ATCC; Manassas, VA, USA), respectively. The cell lines were cultured in RPMI or McCoy's medium and supplemented with 10% fetal bovine serum (FBS) (Gibco, Gaithersburg, MD, USA) and 1% antibiotic antimycotic solution. Cells were incubated at 37°C with 5% CO₂. Escin was purchased from Sigma-Aldrich Co. (St. Louis, MO, USA). Escin was prepared by dissolving the lyophilized powder in dimethyl sulfoxide (DMSO) to a final concentration of 100 mmol/l. The stock solution was stored at −20°C until use.

The cytotoxic effects of escin on renal cancer cell lines were assessed with the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay (Sigma-Aldrich Co.). The cell lines were seeded onto 24-well plates and treated with various concentrations of escin for 24, 48, and 72 h. After incubation for the indicated periods, the medium was removed, and 200 µl of 1X MTT solution was added to each well for 4 h. The medium was aspirated and the formazan product in the cells was solubilized by adding DMSO. An aliquot of 150 µl was measured using a microplate autoreader (L225-0137; PerkinElmer, Taipei, Taiwan) at the wavelength of 540 nm. The IC₅₀ values with dose-dependent curves were calculated by linear interpolation and carried out in triplicates.

Propidium iodide (PI) staining and flow cytometry were used to perform cell cycle analysis. First, 1×10⁶ 786-O cells were seeded on 10 cm dishes for 24 h. The cells were collected after trypsinization and washed with ice-cold phosphate-buffered saline (PBS), fixed and permeabilized with 70% ethanol at −20°C overnight. The next day, after the cells were washed with ice-cold PBS, they were incubated with PI staining solution (0.2 mg/ml RNase, 20 µg/ml PI and 0.1% Triton X-100) for 30 min at room temperature in the dark. Data were collected using a flow cytometer (BD FACSCalibur; BD Biosciences, San Jose, CA, USA) and data were analyzed with WinMDI software (version 2.9). All experiments were performed in triplicate and 10,000 events were counted for each sample.

The BioVision Annexin V-FITC apoptosis detection kit was used for the apoptosis assay (BioVision Inc., Milpitas, CA, USA). First, 786-O cells were seeded onto 10-cm dishes for 24 h and then exposed to different doses of escin for 16 h. Cells were harvested by trypsinization, washed twice with PBS, and resuspended in 500 µl of binding buffer. Cell suspensions were then incubated with 5 µl of Annexin V-FITC and 5 µl of PI for 10 min at room temperature in the dark. The cells were immediately evaluated by flow cytometry (FACSCalibur; Becton-Dickinson, San Jose, CA, USA).

Renal cancer cells (1×10⁶) were seeded in 10-cm culture dishes overnight and treated with the indicated concentrations of escin for 24 h. The cells were harvested, washed twice in PBS, and lysed for 30 min at 4°C with ice-cold RIPA buffer (1% NP-40 in 150 mM NaCl), 50 mM Tris (pH 7.5) and 2 mM EDTA. Protein concentrations were measured using the Bradford protein assay. Equal amounts of protein were loaded on 10–15% sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) gels, transferred to polyvinylidene difluoride (PVDF) membranes, and blocked with 5% non-fat milk in Tris-buffered saline with 0.5% Tween-20 (TBST) buffer (20 mM Tris-HCl, 120 mM NaCl, 0.1% Tween-20) for 1 h. The membranes were incubated with various primary antibodies against cdc-2 (GeneTex, Inc., Irvine, CA, USA), Bcl-2, Bcl-xL, Bax, caspase-3 and −9, poly(ADP-ribose) polymerase (PARP) (1:1,000; Cell Signaling Technology, Boston, MA, USA) and cyclin B1, caspase-8 (p18), Fas, FasL, FADD, β-actin and GAPDH (1:1,000; Santa Cruz Biotechnology, Santa Cruz, CA, USA) at 4°C overnight. After washing, the blots were incubated with HRP-labelled secondary antibodies for 2 h. The signals of the blots were then developed using the enhanced chemiluminescence (ECL) system and analyzed with the LAS3000 system (Fujifilm, Tokyo, Japan).

Mitochondrial-specific cationic dye JC-1 (Invitrogen, Carlsbad, CA, USA), which undergoes potential-dependent accumulation in the mitochondria, was used. When the mitochondrial membrane potential (ΔΨm) is below 120 mV, JC-1 is monomeric and emits green light (540 nm) following excitation with blue light (490 nm). At membrane potentials >120 mV, JC-1 monomer aggregates and emits red light (590 nm) following excitation with green light (540 nm). For the assay, the cells were seeded onto 6-well plates and treated with various concentrations of escin for 24 h, followed by staining with 5 µM JC-1 for 30 min at 37°C. Fluorescence was monitored with a fluorescence plate reader at wavelengths of 490 nm (excitation)/540 nm (emission) and 540 nm (excitation)/590 nm (emission). Changes in the ΔΨm were indicated by changes in the ratio of intensities between the measurements at test wavelengths of 590 nm (red) and 540 nm (green).

A flow cytometric assay of intracellular ROS, which can trigger apoptosis, using dihydroethidium (DHE) (Setareh Biotech LLC, Eugene, OR, USA), a fluorescent superoxide indicator, was previously described (28–31). In the present study, 786-O cells were treated with 0–100 µM escin for 24 h. Subsequently, these cells were incubated with 2 µM DHE in serum-free medium at 37°C for 15 min, washed once with serum-free medium, and then centrifuged at 450 × g to remove extracellular DHE. Finally, the cells were analyzed by flow cytometry.

Experiments were performed three times and the data are presented as mean ± SD. Student's t-test was carried out to assess the statistical differences. p<0.05 was considered to indicate a statistically significant result.

To determine the cytotoxic effect of escin on human renal cancer cells, 786-O and Caki-1 cells were treated with several concentrations of escin (12.5–100 µM) for 24, 48 and 72 h. Normal human peripheral blood mononuclear cells (PBMCs) (as normal control cells) were treated with the indicated concentrations of escin for 24 h. Cell viability of these treated cells was then determined using the MTT assay. As shown in Fig. 1B and C, after 24 h, escin markedly reduced the viability of the 786-O and Caki-1 cells in a concentration-dependent manner with IC₅₀ values of 40.6±1.2 and 35.0±0.8 µM, respectively. The IC₅₀ values of escin were 40.6±1.2, 35.4±0.5 and 26.2±0.5 µM in the 786-O cells at 24, 48 and 72 h, respectively. In contrast, PBMCs treated with escin for 24 h exhibited >75% cell survival (Fig. 1B). At the concentration of 50 µM escin, marked morphological changes such as shrinkage and rounding were noted in the 786-O cells (Fig. 1D). This corroborated the results from the MTT assay (Fig. 1B and C). Taken together, these results suggest that escin induces cell death or apoptosis.

To elucidate the underlying mechanism of escin-induced cell death, we analyzed the distribution of the cell cycle in 786-O cells treated with several concentrations of escin for 24 h. As shown in Fig. 2A and B, the mean percentage of sub-G1 cells, which is an indicator of cell death, in 786-O cells treated with the higher tested concentration (50 µM) of escin (17.7±0.7%) was significantly higher than that in the untreated cells (0.7±0.02%) (p<0.01). In addition, the percentage of cells in the G2/M transition was significantly higher in 786-O cells treated with 25 µM escin (41.6±2%) than in untreated cells (28.3±1.1%) (p<0.01). The levels of cdc-2 protein were decreased relative to the internal control of GAPDH in the cells treated with 25 µM escin, but there were no significant changes in cyclin B1 protein levels following treatment with 25 µM escin (Fig. 2C). These results suggest that escin induces cell cycle G2/M arrest in 786-O cells.

To elucidate the type of cell death caused by escin, we stained escin-treated 786-O cells with Annexin V-FITC and PI to define apoptosis and necrosis, respectively. The percentage of early and late apoptotic, and necrotic cells in the 786-O cell line increased in a concentration-dependent manner (Fig. 3A). At the highest tested concentration of escin, early apoptosis occurred in 77.5% of the 786-O cells compared to the internal control (4.5%) in a concentration-dependent manner (Fig. 3A). Thus, the percentages of live, early apoptotic and late apoptotic cells were calculated and subjected to statistical analysis (Fig. 3B). These results indicate that escin induced apoptosis in the 786-O cells.

Caspases, such as caspase-9 −8 and −3/7, can be activated by either an intrinsic mitochondrial-mediated pathway or an extrinsic death receptor-mediated pathway (32). Activated caspases cause cleavage of PARP, which is a marker of apoptosis. Thus, to determine whether escin induces apoptosis through the intrinsic or extrinsic pathway, we performed western blot analysis of the pro-form and cleaved forms of caspase-8, −9 and −3, and PARP. As illustrated in Fig. 3C, 786-O cells treated with indicated concentrations of escin for 24 h expressed elevated levels of the cleaved forms of caspase-9 and −3, and PARP. It is important to note that the protein level of caspase-3 in 50 µM escin-treated 786-O cells was not detected. We reasoned that many cells treated with 50 µM escin may be dead and detach from the wells. In addition, the cleaved caspase-3 protein (17 kDa), which is the smallest proteins among all caspase-cleaved protein molecules, is easily further degraded and may not be detected by western blot analysis. However, the cleaved caspase-8 protein level (molecular mass; 18 kDa) was not detected among the concentrations of escin used to treat the cells (Fig. 3C). These results suggest that escin induced intrinsic-pathway apoptosis in these renal cancer cells.

To confirm that escin induces intrinsic apoptosis, we determined the effect of escin on the mitochondrial membrane potential (ΔΨm) in 786-O cells. Loss of ΔΨm is a hallmark of intrinsic apoptosis, since it is associated with the release of pro-apoptotic proteins into the cytosol (33). In escin-treated 786-O cells stained with JC-1 dye, a concentration-dependent decrease in red fluorescence and an increase in green fluorescence were observed (Fig. 4B and C). This suggests that escin reduced ΔΨm. These results support the hypothesis that escin induces mitochondrial-mediated apoptosis in the renal cancer cells.

To further elucidate the mechanism of escin-induced mitochondrial-mediated apoptosis, we determined the effect of escin on the expression of Bcl-2 family proteins which regulate cytochrome c release and caspase activity (33,34). Specifically, we examined three Bcl-2 family proteins, namely Bax, which promotes cytochrome c release, and Bcl-2 and Bcl-xL, both of which inhibit cytochrome c release (33). As shown in Fig. 4A, escin (25 and 50 µM) decreased the expression of Bcl-2 and increased the cleavage fragment (molecular mass; 15 kDa) of Bax, although the Bcl-xL protein level was unchanged in 786-O cells, which differed from the observed expression patterns of the other proteins. Furthermore, the levels of IAP proteins, including XIAP and survivin, which can bind to caspases-9/−3/−7 and prevent apoptosis, were significantly decreased (Fig. 4A). Taken together, these results suggest that escin induced mitochondrial-mediated apoptosis in the renal cancer cells by disrupting anti-apoptotic proteins that regulate the release of caspases and relieve downstream inhibition of apoptosis.

Since ROS can alter the cellular redox state and ΔΨm (35,36), we investigated whether escin-treated 786-O cells produce ROS. DHE staining revealed that red fluorescence intensity (Fig. 5A and B) and ROS production (Fig. 5C) increased in the 786-O cells treated with escin for 24 h. In addition, DHE staining showed that the red fluorescence intensity was significantly increased in the 786-O cells treated at the indicated concentration (≥12.5 µM) of escin (Fig. 5C). This indicates that ROS play a crucial role in escin-induced apoptosis in 786-O cells. Moreover, pretreatment of these cells with two antioxidants, NAC and GSH, reduced escin-induced cell apoptosis (Fig. 5D). Collectively, these results suggest that escin-induced mitochondrial-mediated apoptosis is associated with ROS production in renal cancer cells.

To ascertain whether escin induces extrinsic apoptosis, we determined the effect of escin on the expression of proteins involved in the death receptor pathway in 786-O cells. As shown in Fig. 6, treatment of 786-O cells with >25 µM escin did not increase but decreased expression of the Fas death receptor, Fas-L and Fas-associated death domain (FADD) protein levels as compared to those of internal control β-actin. These results support the hypothesis that escin does not induce death receptor-mediated apoptosis in renal cancer cells.