U937, A549, MDA231 and HCT116 cells were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). The culture medium used throughout these experiments was RPMI-1640 (U937, A549, and HCT116) or DMEM (MDA231), containing 10% fetal bovine serum (FBS), 20 mM HEPES buffer and 100 μg/ml gentamicin. Bcl-2-overexpressing U937 cells were generated using a pMAX vector containing the human Bcl-2 gene (provided by Dr Rakesh Srivastava, NIH/NIA). U937 cells (400 μl) in RPMI-1640 (20×10⁶ cells/ml) were transfected by pre-incubation with 15 μg of the Bcl-2 plasmid for 10 min at room temperature and then electroporating at 500 V, 700 μF. The sample was immediately placed on ice for 10 min and then 10 ml complete medium was added and the cells were incubated at 37°C for 24 h. The cells were selected in a medium containing 0.7 μg/ml geneticin (G418) for 4 weeks. Single-cell clones were obtained by limiting dilution and were subsequently analyzed for an increase in Bcl-2 protein expression relative to the identically cloned empty vector control. Chemicals were purchased from Sigma-Aldrich (St. Louis, MO, USA). Anti-Bcl-2, anti-Bcl-xL, anti-Mcl-1, anti-XIAP, anti-cIAP1, anti-cIAP2, anti-cytochrome c, and anti-PARP antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Anti-c-FLIP(L) antibody was obtained from Alexis Corporation (San Diego, CA, USA). Anti-phospho-ERK, anti-phospho-JNK, anti-JNK, anti-phospho-p38 MAPK and anti-p38 antibodies were purchased from Cell Signaling Technology (Beverly, MA, USA). Anti-ERK antibody was obtained from Transduction Laboratories (Lexington, KY, USA). Anti-Qps-2 antibody was purchased from Molecular Probes (Eugene, OR, USA). Anti-actin antibody was obtained from Sigma-Aldrich.

Cells were suspended in 100 μl of phosphate-buffered saline (PBS), and 200 μl of 95% ethanol was added during vortexing. Cells were incubated at 4°C for 1 h, washed with PBS and resuspended in 250 μl of 1.12% sodium citrate buffer (pH 8.4) together with 12.5 μg of RNase. Incubation was continued at 37°C for 30 min. The cellular DNA was then stained by applying 250 μl of propidium iodide (50 μg/ml) for 30 min at room temperature. The stained cells were analyzed by fluorescence-activated cell sorting on a FACScan flow cytometer for the relative DNA content based on red fluorescence.

The cell death detection ELISAPLUS kit (Boehringer Mannheim, Indianapolis, IN, USA) was used for assessing apoptotic activity by detecting fragmented DNA within the nucleus in the RU486-treated cells. Briefly, each culture plate was centrifuged for 10 min at 200 × g, the supernatant was removed, and the pellet was lysed for 30 min. After centrifuging the plate again at 200 × g for 10 min, the collected supernatant containing cytoplasmic histone-associated DNA fragments was incubated with an immobilized anti-histone antibody, and the reaction products were determined by spectrophotometry. Finally, the absorbance at 405 nm and 490 nm (reference wavelength), upon incubating with a peroxidase substrate for 5 min, was determined with a microplate reader. Signals in the wells containing the substrate only were subtracted as background.

After treatment, cells were lysed, and 20 μg of cell lysates was incubated with 100 μl reaction buffer (1% NP-40, 20 mmol/l Tris-HCl (pH 7.5), 137 mmol/l NaCl, 10% glycerol) containing the caspase substrate (DEVD-chromophore p-nitroanilide) at 5 μmol/l. Lysates were incubated at 37°C for 2 h. The absorbance at 405 nm was measured with a spectrophotometer.

Cellular lysates were prepared by suspending 1.2×10⁶ cells in 100 μl of lysis buffer (137 mM NaCl, 15 mM EGTA, 0.1 mM sodium orthovanadate, 15 mM MgCl₂, 0.1% Triton X-100, 25 mM MOPS, 100 μM phenylmethylsulfonyl fluoride, and 20 μM leupeptin, adjusted to pH 7.2). The cells were disrupted by sonication and extracted at 4°C for 30 min. The proteins were electrotransferred to Immobilon-P membranes (Millipore Corp., USA). Detection of specific proteins was carried out with an ECL western blotting kit according to the manufacturer’s instructions.

U937 human leukemia cells (1.2×10⁶ cells/ml) were harvested, washed once with ice-cold PBS and gently lysed for 2 min in 80 μl ice-cold lysis buffer [250 mM sucrose, 1 mM EDTA, 20 mM Tris-HCl (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]. Lysates were centrifuged at 12,000 × g at 4°C for 10 min to obtain the supernatants (cytosolic extracts free of mitochondria) and the pellets (fraction that contains mitochondria). The resulting cytosolic fractions were used for western blot analysis using an anti-cytochrome c antibody.

Rhodamine 123 (Molecular Probes, Inc., Eugene, OR, USA) uptake by mitochondria is directly proportional to its membrane potential. U937 cells subjected to a 4-h treatment were incubated with rhodamine 123 (5 μM) for 30 min in the dark at 37°C. The cells were harvested and suspended in PBS. The mitochondrial membrane potential was subsequently analyzed using a flow cytometer (Becton-Dickinson, Franklin Lakes, NJ, USA).

Data were analyzed with one-way ANOVA, followed by post hoc comparisons (Student-Newman-Keuls) using the Statistical Package for Social Sciences 8.0 (SPSS Inc., Chicago, IL, USA).

In order to investigate whether RU486 induces apoptosis, U937 cells were treated with various concentrations of RU486 for 12 h. We first determined the apoptosis in U937 cells using flow cytometric analysis to detect hypodiploid cell populations. As shown in Fig. 1A, treatment of U937 cells with RU486 resulted in a markedly increased accumulation of sub-G1 phase cells in a dose-dependent manner. Next, we also investigated whether RU486 induces the apoptotic DNA fraction in U937 cells. As shown in Fig. 1B, increasing concentrations of RU486 induced the progressive accumulation of apoptotic DNA. Since cells undergoing apoptosis execute the death program by activating caspases and cleaving PARP (10), we next analyzed PARP cleavage. Exposure to RU486 induced a dose-dependent cleavage of PARP (Fig. 1C). These results suggest that RU486 induced apoptosis in the U937 human leukemia cells. In addition, RU486-induced apoptosis was also observed in a variety of tumor cell types [breast carcinoma cells (MDA231), lung carcinoma cells (A549) and colon cancer cells (HCT116)], demonstrating that RU486-induced apoptosis is a common response in various types of cancer cells (Fig. 1D).

We next analyzed whether treatment with RU486 results in the activation of caspases, a key executioner of apoptosis. Exposure of U937 cells to RU486 strongly stimulated caspase-3 activity in a dose-dependent manner (Fig. 2A). As shown in Fig. 2B, western blot analysis showed that RU486 concentration-dependently induced a marked change in the cleavage of caspase-3. In order to confirm that the activation of caspase-3 is a key step in the RU486-induced apoptotic pathway, U937 cells were pretreated with z-VAD-fmk (25 μM), a cell-permeable pan-caspase inhibitor, followed by treatment with 30 μM RU486 for 12 h. As shown in Fig. 2C, RU486-induced apoptosis was significantly prevented by pretreatment with the potent inhibitor of caspases, z-VAD-fmk, as determined by FACS analysis. We also found that z-VAD-fmk prevented caspase-related events such as cleavage of PARP (Fig. 2C). These results suggest that RU486-induced cell death is associated with caspase-3 activation.

To investigate the underlying mechanisms involved in RU486-induced apoptosis, we analyzed the changes in the expression levels of various anti-apoptotic proteins. As shown in Fig. 3A, protein levels of Bcl-2, Bcl-xL, cIAP1 and cIAP2 were not altered in response to RU486 treatment. However, protein levels of Mcl-1, cFLIP(L) and XIAP were markedly reduced in U937 cells following treatment at the indicated concentrations of RU486 in a dose-dependent manner. Accumulating evidence suggests that mitochondria play an essential role in apoptosis by releasing apoptogenic effectors such as cytochrome c. When we performed western blot analysis using cytosolic fractions to examine the release of mitochondrial cytochrome c in RU486-treated U937 cells, RU486 treatment markedly induced a dose-dependent release of cytochrome c into the cytoplasm (Fig. 3B). In addition, the role of mitochondria in RU486-induced apoptosis of U937 cells was further investigated by examining the effect of RU486 on mitochondrial membrane potential (MMP, Δψm). Exposure of U937 cells to 30 μM RU486 for 4 h led to a significant reduction in the MMP level (Fig. 3C).

In order to evaluate the functional role played by Bcl-2 in preventing RU486-induced apoptosis, cells stably overexpressing Bcl-2 were established. As shown in Fig. 4A, treatment of U937/Vector cells with 30 μM RU486 for 12 h resulted in a markedly increased accumulation of cells in the sub-G1 phase. In contrast, the accumulation of cells in the sub-G1 phase induced by RU486 was inhibited by Bcl-2 overexpression. Subsequent western blot analysis demonstrated that the proteolytic cleavage of PARP in U937/Vector cells was more prominent than that in the U937/Bcl-2 cells when exposed to RU486 (Fig. 4A). Taken together, these results indicate that ectopic expression of Bcl-2 inhibits RU486-induced apoptosis.

Since oxidative stress-mediated cellular changes are induced in cells exposed to cytotoxic drugs and ROS is known to be a mediator of caspase-dependent cell death (11–13), we investigated whether ROS generation induced by RU486 is directly associated with the induction of apoptosis. However, pretreatment with N-acetylcysteine (NAC) did not prevent RU486-induced increase in the sub-G1 cell population and cleavage of PARP (Fig. 4B). These data indicate that ROS generation is not critical for the induction of apoptosis by RU486.

We investigated the effect of RU486 treatment on the expression and activity of MAPKs in order to determine whether this signaling pathway is involved in mediating the observed apoptotic response. As shown in Fig. 5A, RU486 treatment induced a decrease in the phosphorylated ERK levels in a time-dependent manner. However, the phosphorylation of p38 MAPK and JNK gradually increased after RU486 treatment. We then evaluated the possible roles of MAPKs in RU486 treatment-induced apoptosis. As shown in Fig. 5B, pretreatment with SB203580 (a specific inhibitor of p38 MAPK) decreased the sub-G1 phase cell population (17.3%) when compared with the sub-G1 phase cell population following RU486 treatment (28.8%), while treatment of SP600125 (a potent inhibitor of JNK) and PD98059 (a potent inhibitor of ERK) did not significantly decrease the number of cells with sub-G1 DNA content. These results indicate that the activation of the p38 MAPK pathway plays an important role in regulating RU486-induced apoptosis in U937 cells.

Taken together, our results demonstrated that RU486 induced apoptosis through reduction in mitochondrial membrane potential and activation of p38 MAPK in U937 human leukemia cells.