Fetal bovine serum (FBS), Dulbecco’s modified Eagle’s medium (DMEM), penicillin, streptomycin and trypsin-ethylenediaminetetraacetic acid (EDTA) were purchased from Gibco-BRL (Gaithersburg, MD, USA). 3-(4,5-Dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT), trypan blue, 4,6-diamidino-2-phenyllindile (DAPI), propidium iodide (PI), paraformaldehyde, 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethyl-imidacarbocyanine iodide (JC-1) and N-acetyl L-cysteine (NAC) were purchased from Sigma-Aldrich (St. Louis, MO, USA). The pan-caspase inhibitor z-VAD-fmk and inhibitors of phosphatidylinositol 3-kinase (PI3K)/Akt and mitogen-activated protein kinase (MAPK) were obtained from Calbiochem (San Diego, CA, USA). DNA ladder size markers were purchased from Invitrogen (Carlsbad, CA, USA). 2′,7′-Dichlorofluorescein diacetate (DCFDA) and fluorescein-isothiocyanate (FITC)-Annexin V were purchased from Molecular Probes (Eugene, OR, USA) and BD Pharmingen (San Diego, CA, USA), respectively. Caspase activity assay kits and an enhanced chemiluminescence (ECL) kit were purchased from R&D Systems (Minneapolis, MN, USA) and Amersham (Arlington Heights, IL, USA), respectively. Antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA), Chemicon (Temecula, CA, USA) and Sigma-Aldrich. Peroxidase-labeled donkey anti-rabbit and sheep anti-mouse immunoglobulin were purchased from Amersham. All other chemicals were purchased from Sigma-Aldrich.

C. rotundus rhizomes were obtained from the Dongeui Oriental Hospital, Dongeui University College of Oriental Medicine, Busan, Republic of Korea. They were cut into small pieces, ground to powder, passed through a 20-mesh sieve, and transferred to a flask in a 1:10 proportion (drug: solvent, w/v), using methanol, ethanol and water. The materials were protected from light with aluminum foil, and the flasks were shaken at 50 rpm for 24 h at 60°C. The extracts were filtered through Whatman paper no. 1 and dried in a Rotavapor at 40°C, resulting in methanol (MECR), ethanol (EECR) and water (WECR) extracts. The extracts were weighed, resuspended into dimethyl sulfoxide to a final concentration of 100 mg/ml (extract stock solution) and then diluted with medium to the desired concentration prior to use.

The MDA-MB-231 human breast carcinoma cell line was purchased from the American Type Culture Collection (Manassas, VA, USA) and maintained in DMEM supplemented with 10% FBS, 1% L-glutamine and penicillin/streptomycin. The cells were cultured in an incubator with 5% CO₂ at 37°C.

Cell growth was assessed using the trypan blue dye exclusion assay. In brief, cells (2×10⁴ cells/well) were seeded in 6-well plates. After treatment with the indicated concentrations of the C. rotundus extracts for 24 h, the cells were trypsinized and viable cells were counted by trypan blue dye exclusion using a hemocytometer under an inverted microscope (Carl Zeiss, Jena, Germany). Cell viability was determined using the MTT assay. In brief, cells (2×10⁴ cells/well) were seeded in 24-well plates and exposed to the extracts of C. rotundus for 24 h. After treatment, 5 mg/ml MTT solution was added, followed by a 3-h incubation at 37°C in the dark. Absorbance of the formazan product was measured at a wavelength of 540 nm with an enzyme-linked immunosorbent assay (ELISA) reader (Molecular Devices, Sunnyvale, CA, USA). Cells were photographed directly for a morphological study using an inverted microscope.

The cells were washed with phosphate-buffered saline (PBS) and fixed with 3.7% paraformaldehyde in PBS for 10 min at room temperature. The fixed cells were permeabilized with 0.1% Triton X-100 for 10 min, the supernatant was removed and the cells were washed three times with PBS. The cells were incubated with 2.5 μg/ml DAPI for 10 min at room temperature in the dark. The cells were then washed twice with PBS and DAPI-stained cell nuclei (apoptotic nuclei stained intensely) and were observed with a fluorescence microscope (Carl Zeiss).

Cells were lysed in a buffer containing 10 mM Tris-HCl, pH 7.4, 150 mM NaCl, 5 mM EDTA and 0.5% Triton X-100 for 1 h at room temperature for the DNA fragmentation assay. Lysates were vortexed and cleared by centrifugation at 19,000 × g for 30 min at 4°C. A 25:24:1 (v/v/v) equal volume of neutral phenol:chloroform:isoamyl alcohol was used to extract DNA from the supernatant, followed by electrophoretic analysis on 1.5% agarose gels containing 0.1 μg/ml ethidium bromide.

A combination of Annexin V-FITC and PI staining distinguishes between early (Annexin V⁺/PI⁻) and late (Annexin V⁺/PI⁺) apoptotic, necrotic and live cells. After the cells were treated with the EECR for 24 h, they were trypsinized, washed with PBS, and resuspended in 100 μl binding buffer containing 5 μl Annexin V-FITC and 5 μl PI for 15 min at room temperature in the dark. The cells were immediately analyzed by flow cytometry (FACSCalibur; Becton-Dickinson, San Jose, CA, USA). The percentages of apoptotic cells (Annexin V⁺ cells) are presented as the means ± standard deviation as previously described (29).

The cells were collected by trypsin-EDTA and lysed with lysis buffer (20 mM sucrose, 1 mM EDTA, 20 μM Tris-Cl, pH 7.2, 1 mM DTT, 10 mM KCl, 1.5 mM MgCl₂, 5 μg/ml pepstatin A, 10 μg/ml leupeptin and 2 μg/ml aprotinin) containing protease inhibitors for 30 min at 4°C. The mixtures were centrifuged (10,000 × g) for 10 min at 4°C, and the supernatants were collected as whole-cell extracts. Protein content was determined using the Bio-Rad protein assay reagent (Hercules, CA, USA) and bovine serum albumin as the standard according to the manufacturer’s instructions to determine protein concentrations. After normalization, an equal amount of protein was subjected to electrophoresis on sodium dodecyl sulfate-polyacrylamide gels and transferred to nitrocellulose membranes (Schleicher & Schuell, Keene, NH, USA) by electroblotting. The blots were blocked with 5% skim milk for 1 h at room temperature. The blots were incubated overnight with primary antibodies, followed by horseradish peroxidase-conjugated donkey anti-rabbit and sheep anti-mouse immunoglobulin for 1 h. The immunoreactive bands were revealed by enhanced chemiluminescence with a commercially available ECL kit.

MMP values were determined using the dual-emission potential-sensitive probe, JC-1. Briefly, cells were collected and incubated with 10 μM JC-1 for 30 min at 37°C in the dark. After JC-1 was removed, the cells were washed with PBS to remove unbound dye, and the amount of JC-1 retained by 10,000 cells/sample was measured at 488 and 575 nm using a flow cytometer.

Caspase activities were determined with colorimetric assay kits, which utilize synthetic tetrapeptides [Asp-Glu-Val-Asp (DEAD) for caspase-3, Ile-Glu-Thr-Asp (IETD) for caspase-8 and Leu-Glu-His-Asp (LEHD) for caspase-9] labeled with p-nitroaniline (pNA). Briefly, EECR-treated and untreated cells were lysed in the supplied lysis buffer. The supernatants were collected and incubated with the supplied reaction buffer containing dithiothreitol and DEAD-pNA, IETD-pNA or LEHD-pNA as substrates at 37°C. The reactions were measured by changes in absorbance at 405 nm using an ELISA reader.

ROS production was monitored using the stable nonpolar dye DCFDA. The cells were seeded at a density of 1×10⁵ cells/ml, allowed to attach for 24 h and exposed to 200 μg EECR for various periods. The cells were incubated with 10 mM DCFDA for 20 min at 37°C in the dark. ROS production in the cells was monitored with a flow cytometer using CellQuest Pro software.

Data are reported as means ± standard deviation of three independent experiments. A one-way analysis of variance was performed to determine differences. p<0.05 was considered to indicate a statistically significant difference.

To investigate the effects of C. rotundus rhizome extracts on MDA-MB-231 cell growth, the cells were treated with various concentrations of the MECR, EECR and WECR for 24 h, and cell number and viability were measured by trypan blue exclusion and the MTT assay, respectively. As shown in Fig. 1, the MECR and EECR markedly inhibited cell proliferation and viability of MDA-MB-231 cells in a concentration-dependent manner. For example, cell viability was inhibited by >30 or 65% in cells exposed to 150 or 300 μg/ml of the EECR, respectively, compared to that in untreated controls. In contrast, the WECR was not cytotoxic under the same experimental conditions. The EECR possessed stronger cytotoxicity than MECR; thus, we used the EECR for further studies.

Cell morphology was assessed by DAPI staining. The results indicated that the nuclear structure of control cells remained intact, whereas nuclear chromatin condensation and fragmentation, characteristics of apoptosis, were observed in cells treated with the EECR and were associated with numerous cellular morphological changes (Fig. 2A and B). In particular, cell shrinkage and cytoplasm condensation appeared in a dose-dependent manner after EECR treatment. In addition, treatment with the EECR induced progressive accumulation of fragmented DNA, which appeared as a typical ladder pattern of DNA fragmentation due to internucleosomal cleavage associated with apoptosis, in a concentration-dependent manner (Fig. 2C). Furthermore, we evaluated apoptotic induction by flow cytometry analysis with Annexin V and PI staining. As shown in Fig. 2D, treatment with the EECR resulted in an increased accumulation of apoptotic cells compared with that of untreated control cells.

The death receptor and corresponding pro-apoptotic ligands were examined by western blot analysis to determine which apoptosis pathway contributes to EECR-induced apoptosis. The results showed that the levels of TRAIL, Fas and FasL expression were relatively unchanged in response to EECR treatment; however, the levels of death receptor (DR) 4 and DR5 markedly increased in response to EECR treatment (Fig. 3). Next, we examined the effects of the EECR on the levels of Bcl-2 family proteins. The results of immunoblotting revealed downregulation of anti-apoptotic Bcl-2, but not Bcl-xL, in MDA-MB-231 cells (Fig. 3). However, treatment with the EECR caused a marked increase in pro-apoptotic Bax expression, but not Bad. In addition, relative protein expression of anti-apoptotic survivin decreased in a concentration-dependent manner compared to that in control cells, whereas expression of XIAP, cIAP-1 and cIAP-2 was relatively constant in EECR-treated MDA-MB-231 cells (Fig. 3).

We next examined the effect of the EECR on caspase activity to determine whether caspase activation occurs during EECR-induced apoptosis. Fig. 4A shows that treatment of MDA-MB-231 cells with the EECR decreased pro-caspase-8 and -9 levels, which are initiator caspases of the extrinsic and intrinsic apoptotic pathways, respectively. In conjunction with the decrease in pro-caspase-8 and -9 expression, western blotting revealed that EECR treatment of MDA-MB-231 cells also resulted in the downregulation of pro-caspase-3. Furthermore, activation of caspases by the EECR was confirmed by measuring enzyme activity using the specific synthetic substrates for each caspase. As indicated in Fig. 4B, we observed a dose-dependent gradual increase in caspase-3, -8 and -9 activities in EECR-treated MDA-MB-231 cells similar to the proteolytic processing of pro-caspases. Activation of caspase-3 was further evidenced from cleavage of PARP from a 116 kDa band to a 89 kDa fragment, a substrate of active caspase-3, which also serves as a marker of cells undergoing apoptosis (30).

Since the EECR activated caspase-9, we investigated whether it would affect EECR-induced apoptosis associated with mitochondrial signaling. Thus, we examined the effects of the EECR on mitochondrial membrane integrity, one of the early events leading to apoptosis, using the JC-1 fluorescent probe. The results in Fig. 4C indicate that treatment with the EECR clearly elicited dissipation of the MMP when compared to that in control cells. The extrinsic apoptotic signaling cascade starts with activation of caspase-8 and truncation of Bid (tBid), a BH3 pro-apoptotic protein, which translocates to the mitochondrial membrane, allowing activation of pro-apoptotic proteins (8,9). Therefore, we further examined the effect of the EECR on the level of Bid. As indicated in Fig. 3, EECR treatment caused a decrease in the amount of the Bid pro-form, which is indirect evidence of protein truncation and activation, suggesting that EECR-induced apoptosis in MDA-MB-231 cells may occur via activation of caspase-8 and Bid truncation.

To further confirm the significance of caspase activation in EECR-induced apoptosis, we examined the effects of the general caspase inhibitor, z-VAD-fmk. As shown in Fig. 5A and B, pre-treatment with z-VAD-fmk completely abrogated the appearance of cells with apoptotic features, such as chromatin condensation and formation of apoptotic bodies, and attenuated the accumulation of fragmented DNA. A flow cytometric analysis and MTT assay indicated that pretreatment of cells with z-VAD-fmk prevented EECR-induced accumulation of apoptotic cell population and growth inhibition (Fig. 5C and D). These results show that EECR-induced apoptosis in MDA-MB-231 cells occurred via a caspase-dependent pathway.

Oxidative stress occurs due to an imbalance in pro-oxidant and antioxidant levels in cells. Excess ROS generation through an imbalance between pro-oxidants and antioxidants leads to damage of cellular macromolecules including DNA, proteins and lipids, and eventually results in physical and chemical damage to tissues that may lead to cell death (31,32). Therefore, we investigated whether ROS generation was also involved in EECR-induced apoptosis. As shown in Fig. 6A, the EECR markedly increased intracellular ROS levels within 15 min, and pretreatment with NAC, a well known ROS scavenger, markedly attenuated EECR-induced ROS generation. However, pretreatment with NAC did not block EECR-induced apoptotic morphological changes or DNA fragmentation (Fig. 6B and C). Moreover, NAC did not inhibit EECR-induced apoptosis and growth inhibition (Fig. 6D and E). Collectively, these results clearly indicate that the EECR-induced apoptosis in MDA-MB-231 cells is not mediated by ROS generation.

Next, we investigated the effect of EECR treatment on the expression and activities of PI3K/Akt and MAPKs to determine whether these signaling pathways play a role in mediating the observed apoptotic response. Western blot analysis showed that total Akt protein and phosphorylated Akt levels did not change significantly in response to EECR treatment. In addition, the EECR did not lead to phosphorylation of the three MAPKs including extracellular signal-regulated kinase (ERK), p38 MAPK and c-Jun NH2-terminal kinase (JNK) (Fig. 7), and there was also no effect on steady-state levels of total MAPK proteins in MDA-MB-231 cells treated with EECR. Moreover, pretreatment with a representative PI3K/Akt inhibitor, LY294002 and inhibitors of MAPKs (PD98059, a potent inhibitor of ERK; SP600125, a potent inhibitor of JNK; SB203589, a specific inhibitor of the p38 MAPK) did not have a significant effect on EECR treatment (Fig. 8). Collectively, these results suggest that the PI3K/Akt and MAPK pathways did not play a role in regulating EECR-induced apoptosis of MDA-MB-231 cells.