The dried sclerotium of P. cocos was supplied by Dongeui University Oriental Hospital (Busan, Korea) and authenticated by Professor S.H. Hong, Department of Biochemistry, Dongeui University College of Korean Medicine. A voucher specimen (accession no. DEU-27) was deposited at the Natural Resource Bank of Dongeui University College of Korean Medicine. To prepare the EEPC, the dried sclerotium of P. cocos was ground into powder and extracted twice with 10 volumes of 80% ethanol at 85–90°C in a reflux condenser for 3 h. After being filtered through a 0.2-µm filter, the extract was concentrated and lyophilized by vacuum evaporation at 60°C. The solid form of the extract was dissolved in dimethylsulfoxide (DMSO; Sigma-Aldrich, St. Louis, MO, USA) prior to the experiment.

The human leukemia U937 cells and Chang liver cells (an immortalized non-tumor cell line derived from normal liver tissue) were purchased from the American Type Culture Collection (ATCC; Manassas, VA, USA), and maintained at 37°C in a humidified environment (95% air and 5% CO₂), in RPMI-1640 supplemented with 10% heat-inactivated fetal bovine serum (FBS), 2 mM glutamine, 100 U/ml penicillin and 100 µg/ml streptomycin (all from Gibco-BRL, Gaithersburg, MD, USA). Ectopic Bcl-2-overexpressing U937 (U937/Bcl-2) cells were generously provided by Professor T.K. Kwon (Department of Immunology, School of Medicine, Keimyung University, Daegu, Korea) and were maintained in a medium containing 0.7 µg/ml geneticin (G418 sulfate; Calbiochem, San Diego, CA, USA).

For the cell viability assay, cells were seeded at a concentration of 1×10⁵ cells/ml and were treated with the indicated concentrations of EEPC for 24 h or with 90 µg/ml EEPC for the indicated times. After treatments, 3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide (MTT; Sigma-Aldrich) working solution was added to each culture plate and continuously incubated at 37°C for 3 h. The culture supernatant was removed from the wells, and DMSO was added to completely dissolve the formazan crystals. The absorbance of each well was measured at a wavelength of 540 nm with an enzyme-linked immunosorbent assay (ELISA) plate reader (Molecular Devices, Sunnyvale, CA, USA) (30). Cell growth was assessed using the trypan blue dye exclusion assay. In brief, 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). The morphological changes of cells incubated with or without EEPC for 24 h were examined under an inverted microscope.

For 4′,6-diamidino-2-phenylin-dole (DAPI; Sigma-Aldrich) staining, the cells were washed with phosphate-buffered saline (PBS) and fixed with 3.7% paraformaldehyde (Sigma-Aldrich) in PBS for 10 min at room temperature. The fixed cells were washed with PBS and stained with 2.5 µg/ml DAPI solution for 10 min at room temperature. The cells were then washed twice with PBS and analyzed by fluorescence microscopy (Carl Zeiss).

The cells were fixed in 70% ethanol overnight at 4°C, washed in PBS and then resuspended in 1.12% sodium citrate buffer (pH 8.4) together with 12.5 µg of RNase (DNase-free; Sigma-Aldrich). Incubation was continued at 37°C for 30 min. The cellular DNA was then stained by applying a propidium iodide (PI) (10 µg/ml; Sigma-Aldrich) solution for 30 min at room temperature in the dark. The stained cells were analyzed using a FACSCalibur flow cytometer (Becton-Dickinson; San Jose, CA, USA). The level of apoptotic cells containing sub-G1 DNA content was determined as a percentage of the total number of cells (31).

For the preparation of total cellular protein, the cells were gently lysed with lysis buffer [40 mM Tris (pH 8.0), 120 mM, NaCl, 0.5% NP-40, 0.1 mM sodium orthovanadate, 2 µg/ml aprotinin, 2 µg/ml leupeptin and 100 µg/ml phenymethylsulfonyl fluoride] for 30 min. Supernatants were collected and protein concentrations were determined using a Bio-Rad protein assay kit (Bio-Rad, Hercules, CA, USA). In a parallel experiment, the mitochondrial and cytosolic fractions were isolated using a mitochondrial and cytosolic fractionation kit (Active Motif, Carlsbad, CA, USA) according to the manufacturer's protocol. Equal amounts of protein were separated on sodium dodecyl sulfate (SDS)-polyacrylamide gels. Separated protein was transferred to nitrocellulose membranes (Schleicher & Schuell, Keene, NH, USA) and subsequently blocked with tris-buffered saline (10 mM of Tris-Cl, pH 7.4) containing 0.5% Tween-20 and 5% non-fat dry milk for 1 h at room temperature. The proteins were probed with primary antibodies overnight at 4°C. After probing them with the primary antibodies, the membranes were incubated with horseradish peroxidase-conjugated anti-rabbit IgG as a secondary antibody purchased from Amersham Corporation (Arlington Heights, IL, USA). Using an enhanced chemiluminescence (ECL) detection system (Amersham Corporation), immunoreactive bands were detected and exposed to an X-ray film. Primary antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA), Cell Signaling Technology, Inc. (Danvers, MA, USA) and Abcam (Cambridge, UK).

The activities of the caspases were determined by colorimetric assay kits (R&D Systems, Minneapolis, MN, USA), which utilize synthetic tetrapeptides [Asp-Glu-Val-Asp (DEVD) for caspase-3; Ile-Glu-Thr-Asp (IETD) for caspase-8; Leu-Glu-His-Asp (LEHD) for caspase-9] labeled with p-nitroaniline (pNA). Briefly, cells were lysed in the supplied lysis buffer according to the manufacturer's protocol. The supernatants were collected and incubated with the supplied reaction buffer and DEVD-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 plate reader (32).

The MMP of intact cells was measured by a flow cytometer using the lipophilic cationic probe 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetra-ethylbenzimidazolylcarbocyanine iodide (JC-1; Calbiochem). U937 cells were collected and resuspended in PBS and then incubated with 10 µM JC-1 for 20 min at 37°C. The cells were subsequently washed once with cold PBS, suspended and subsequently analyzed using a flow cytometer (33).

All data were derived from at least three independent experiments. Statistical analyses were conducted using SigmaPlot software, and values are presented as mean ± SD. Significant differences between the groups were determined using the unpaired Student's t-test.

We first evaluated the cytotoxic effect of EEPC on U937 cells using MTT assay and trypan blue exclusion method, and found that EEPC significantly reduced the cell viability and proliferation of U937 cells in a concentration- and time-dependent manner (Fig. 1A–C). An additional experiment was conducted using Chang liver cells in order to examine the effect of EEPC on the viability of normal cells (Fig. 1D). EEPC concentrations up to 90 µg/ml did not induce cytotoxicity. Therefore, experiments were performed to determine whether this inhibitory effect of EEPC on U937 cell growth resulted from apoptotic cell death. As shown in Fig. 2A and B, EEPC induced U937 cell death with the characteristic apoptotic features, including cell shrinkage, and chromatin condensation and fragmentation in the nucleus as detected by DAPI staining. We next quantified the apoptotic dead cells using flow cytometric analysis to detect hypodiploid cell populations. As shown in Fig. 2C, treatment of U937 cells with EEPC resulted in a markedly increased accumulation of sub-G1 phase cells, and this response occurred in a concentration-dependent manner. These results suggest an association between the growth inhibition observed in response to EEPC and the induction of apoptosis in U937 cells.

We next analyzed whether treatment with EEPC results in the activation of caspases, a cardinal hallmark of apoptosis, including two initiation caspases, caspase-8 and -9, and the executioner caspase-3. As shown in Fig. 3A, the western blot analysis revealed that the expression levels of pro-caspase-8, -9 and -3 were all decreased or the active forms of caspase-8 and -3 were increased in a concentration-dependent manner following EEPC treatment. In addition, quantitative determinations of caspase activities by colorimetric assays consistently showed that the activities of the three caspases were significantly increased by EEPC treatment (Fig. 3B). Furthermore, subsequent immunoblot analysis revealed that progressive proteolytic cleavage products of poly(ADP-ribose) polymerase (PARP), a downstream target protein of activated caspase-3 (34), occurred in U937 cells treated with EEPC. Under the same conditions, levels of the anti-apoptotic inhibitor of apoptosis proteins (IAP) family of proteins, such as XIAP, cIAP-1 and cIAP-2 (Fig. 4A), which bind to caspases and lead to their inactivation (35,36), were markedly inhibited by EEPC treatment in a concentration-dependent manner. These results indicated that EEPC may trigger initiator caspase-8 and -9 initially and subsequently activate the executioner caspase-3.

Given that members of the Bcl-2 protein family are primarily responsible for initiating mitochondrial-mediated apoptosis, we next examined the levels of Bcl-2 family proteins using western blot analysis. As shown in Fig. 4B, EEPC evoked a concentration-dependent reduction in the level of anti-apoptotic Bcl-2 expression, whereas the expression level of pro-apoptotic Bax was markedly induced. The resultant alteration in Bcl-2 family protein expression apparently lowered the ratio of the anti-apoptotic to the pro-apoptotic Bcl-2 family proteins, theoretically causing a disruption in mitochondrial membrane integrity and consequent cytosolic release of cytochrome c to initiate caspase-9 activation (37,38). Therefore, we performed an immunoblot analysis using cytosolic and mitochondrial fractions to examine the release of mitochondrial cytochrome c in EEPC-treated cells and found that EEPC treatment markedly induced a dose-dependent release of cytochrome c into the cytoplasm (Fig. 4C).

Accumulating evidence has demonstrated that Bax protein translocation to the mitochondria is a necessary step in cell activating mitochondrial-mediated apoptosis (39,40). As shown in Fig. 4C, the levels of Bax in the cytosol declined in a concentration-dependent manner after EEPC treatment. In contrast, the Bax levels in the mitochondrial fraction increased, indicating that EEPC treatment markedly led to Bax trans-location into the mitochondria from the cytosol. Moreover, under the same experimental conditions, EEPC caused a concentration-dependent cleavage of BH3-only Bid protein and the formation of its truncated form of Bid, tBid (Fig. 4B), which could translocate to the mitochondria to enhance the mitochondrial-mediated apoptosis pathway (41,42). These results suggest that EEPC reduces the Bcl-2/Bax ratio, inserts Bax from the cytosol into mitochondria, and activates Bid, resulting in mitochondrial dysfunction, release of cytochrome c to the cytosol and apoptosis induction.

Since it has been well established that Bcl-2 plays a critical role in the regulation of the mitochondrial-mediated apoptotic pathway, U937 cells stably overexpressing Bcl-2 were established and we determined the effect of overexpression of Bcl-2 on EEPC-induced apoptosis. As shown in Fig. 5A, western blot analysis revealed that Bcl-2 overexpression inhibited the EEPC-induced cleavage of caspase-3, -9 and PARP. In addition, the role of mitochondria in EEPC-induced apoptosis was further investigated by examining the effect of EEPC on MMP levels. As shown in Fig. 5B, Bcl-2 overexpression was found to significantly block cells from EEPC-induced MMP level reduction when compared to the U937/vector cells. Furthermore, Bcl-2 overexpression was found to significantly protect cells from EEPC-induced morphological changes, accumulation of cells in the sub-G1 phase and growth inhibition (Fig. 6) when compared to U937/vector cells. Taken together, these results indicated that the ectopic expression of Bcl-2 inhibits EEPC-induced apoptosis.