Sulforaphane was purchased from Sigma-Aldrich Chemical Co. (St. Paul, MN), dissolved in dimethyl sulfoxide (DMSO, Sigma-Aldrich), and then diluted with the medium to the desired concentration prior to use. RPMI-1640 medium and fetal bovine serum (FBS) were obtained from Gibco-BRL (Gaithersburg, MD). 3-(4,5-Dimetylthiazol-2-yl)-2,5-diphenyltetrazolium (MTT), 4,6-diamidino-2-phenylindole (DAPI), propidium iodide (PI), 2,7-dichlorodihydrofluorescein diacetate (DCFH-DA), 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylimidacarbocyanine iodide (JC-1) and N-acetyl L-cysteine (NAC) were obtained from Sigma-Aldrich. Caspase activity assay kits and an enhanced chemiluminescence (ECL) kit were purchased from R&D Systems (Minneapolis, MN) and Amersham Life Science Corp. (Arlington Heights, IL), respectively. Primary antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), Chemicon (Temecula, CA), and Abcam (Cambridge, UK). Peroxidase-labeled donkey anti-rabbit and sheep anti-mouse immunoglobulin were purchased from Amersham Life Science Corp. All other chemicals were purchased from Sigma-Aldrich.

T24 cells were purchased from American Type Culture Collection (Manassas, VA) and maintained in RPMI-1640 medium supplemented with 10% FBS, 2 μm L-glutamine and penicillin/streptomycin under a humidified condition of 5% CO₂ at 37°C. In order to measure the inhibition of T24 cell proliferation by sulforaphane, cells were plated in 6-well culture plates (1×10⁵ cells/well) and allowed to adhere overnight, and were then treated with different concentrations of sulforaphane for 24 h. After treatment, cells were incubated with MTT solution (5 mg/ml) for 3 h, and the medium was then removed. The formazan precipitate was dissolved in DMSO, and the absorbance was read at 540 nm in an ELISA reader (Dynatech Laboratories, Chantilly, VA). Morphological changes were monitored by obtaining photomicrographs under an inverted phase contrast microscope (Carl Zeiss, Oberkochen, Germany).

After culture with various concentrations of sulforaphane for 24 h, cells were washed with phosphate-buffered saline (PBS) and fixed with 3.7% paraformaldehyde in PBS for 10 min at room temperature. Fixed cells were washed with PBS, and stained with DAPI solution for 10 min at room temperature. Then, the nuclear morphology of the cells was examined using a fluorescence microscope (Carl Zeiss).

Cells were harvested, washed with PBS and lysed in a buffer [50 mM Tris (pH 8.0), 0.5% sarkosyl, 0.5 mg/ml proteinase K and 1 mM EDTA] at 55°C for 3 h. Lysates were then treated with RNase A (0.5 mg/ml) for 2 h at 56°C. The lysates were centrifuged at 10,000 g for 20 min. The genomic DNA in the supernatant was extracted with an equal volume of neutral phenol/chloroform/isoamyl alcohol mixture (25/24/1). Approximately 20 μg of DNA was loaded into each well of a 1.5% agarose gel and separated by electrophoresis at 50 V for 120 min in Tris-borate/EDTA electrophoresis buffer (TBE). The DNA was visualized and photographed under ultraviolet (UV) illumination after staining with 0.1 μg/ml ethidium bromide (EtBr).

After treatment with sulforaphane for 24 h, cells were harvested, washed with PBS and fixed with 70% ethanol at −20°C overnight. Cells were washed twice, resuspended in PBS containing 40 μg/ml PI, 0.1 mg/ml RNase A and 0.1% Triton X-100 in a dark room for 30 min at 37°C, and analyzed by flow cytometry (Becton-Dickinson, San Jose, CA). The cell cycle distribution and sub-G1 population (apoptosis) were determined and analyzed.

For isolation of total protein fractions, cells were collected, washed twice with cold PBS, and lysed with cell lysis buffer [20 mM Tris pH 7.5, 150 mM NaCl, 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM EDTA, 0.5 g/ml leupeptin, 1% Na₃CO₄, 1 mM phenylmethane-sulfonyl fluoride]. The cell lysates were centrifuged at 13,000 g at 4°C, and the supernatant was collected for western blot analysis, and measured for protein concentration by using the Bio-Rad protein assay kit (Bio-Rad Laboratories, Hercules, CA). In a parallel experiment, mitochondrial and cytosol cellular fractions were prepared using a Cytosol/Mitochondria Fractionation kit (Calbiochem, San Diego, CA) according to the manufacturer’s protocol. The equal aliquots containing 30–50 μg of each lane were separated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis followed by electro-transfer onto nitrocellulose membranes (Schleicher & Schuell, Keene, NH). The membranes were incubated overnight at 4°C with the primary antibodies and the corresponding horseradish peroxidase-conjugated secondary antibodies. The protein bands were visualized using an ECL kit.

Activities of caspases were determined by use of 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, sulforaphane-treated and untreated cells were lysed in the supplied lysis buffer. Supernatants were collected and incubated with the supplied reaction buffer containing dithiothreitol (DTT) 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.

The intracellular accumulation of ROS was examined by flow cytometry after being stained with the cell permeable fluorescent probe, DCFH-DA. Briefly, the cells were washed, resuspended in PBS and incubated with 10 μM DCFH-DA for 20 min at 37°C in the dark. The ROS production in the cells was monitored with a flow cytometer (22).

The values of MMP were determined using the dual-emission potential-sensitive probe JC-1. Briefly, cells were collected and resuspended in PBS supplemented with 10 μM JC-1 for 30 min at 37°C in the dark. After the JC-1 was removed, the cells were washed with PBS to remove unbound dye, and the amount of JC-1 retained by cells was immediately analyzed by flow cytometry.

Unless otherwise stated, data are expressed as means ± standard deviation (SD) and analyzed statistically by one-way analysis of variance. A value of p<0.05 was considered statistically significant.

Initially, the effects of sulforaphane on proliferation of T24 cells were measured by an MTT assay. As shown in Fig. 1, sulforaphane treatment inhibited the cell viability of T24 cells in a concentration-dependent manner: sulforaphane at 10 and 20 μM for 24 h inhibited cell proliferation by 25 and 62%, respectively. In addition, sulforaphane stimulation significantly induced morphological changes, including extensive cytosolic vacuolization and the appearance of irregular cell membrane buds (Fig. 2A). To examine whether the cytotoxicity of sulforaphane was due primarily to apoptosis, we assessed the apoptosis parameters of T24 cells in response to sulforaphane treatment. As shown in Fig. 2B, the nuclear structure of the control cells remained intact, whereas nuclear chromatin condensation and fragmentation, both of which are characteristics of apoptosis, were increased in a concentration-dependent manner in cells treated with sulforaphane, which was associated with increased DNA fragmentation (Fig. 2C). Under the same conditions, results of flow cytometry further demonstrated that treatment with sulforaphane induced a concentration-dependent accumulation of apoptotic sub-G1 population of T24 cells (Fig. 1D). These results indicate that the inhibition of cell viability observed in response to sulforaphane was associated with the induction of apoptosis.

Several gene products are known to be important in controlling the apoptotic process. Among them, caspases, a family of cysteine proteases, play essential roles as important mediators in apoptosis and as determinants of general apoptotic morphology through the cleavage of various cellular substrates, including poly ADP-ribose polymerase (PARP), an endogenous substrate of activated caspase-3 (23,24). To elucidate the molecular mechanisms of sulforaphane-induced apoptosis, we assessed whether sulforaphane induces the proteolytic processing of caspases. As illustrated in Fig. 3A, we detected a concentration-dependent upregulation of cleaved caspase-3 and -9, whereas the cleaved forms of caspase-8 were not detected. In addition, to quantify the proteolytic activation of the caspases, we evaluated in vitro caspase activities using fluorogenic substrates (Fig. 3B). Our results indicated that treatment with sulforaphane significantly increased the activities of caspase-3 and -9 compared with control cells, whereas a very low level of caspase-8 activity was detected after the sulforaphane treatment, suggesting the likely involvement of a mitochondria-dependent cascade for caspase activation. To further identify the activation of the caspase cascade, the levels of PARP were examined by western blot analysis. As shown in Fig. 3A, sulforaphane induced a concentration-dependent proteolytic cleavage of PARP, resulting in a reduction of the 116-kDa protein and accumulation of the 85-kDa fragment. In addition, the levels of inhibitor of apoptosis proteins (IAP) family members, including XIAP, cIAP-1 and cIAP-2, were inhibited by sulforaphane treatment in a concentration-dependent manner (Fig. 3A).

It has been recognized that the Bcl-2 family members play crucial roles in regulating apoptosis by functioning as promoters (e.g., Bax) or inhibitors (e.g., Bcl-2) of cell death. Mitochondria are specialized organelles containing an outer membrane and an inner membrane separated by an intermembrane space that contains many proapoptotic proteins, such as cytochrome c (25,26). We therefore examined the expression of these molecules after sulforaphane treatment. Western blot analysis data showed that sulforaphane markedly increased protein levels of proapoptotic Bax. In contrast to Bax, anti-apoptotic Bcl-2 expression decreased mildly following sulforaphane treatment. To further characterize the apoptotic effect of sulforaphane in T24 cells, we analyzed the translocation of Bax and the release of cytochrome c using cytosol and mitochondria fractionation. Notably, as shown in Fig. 4, sulforaphane treatment led to a concentration-dependent increase in cytosolic cytochrome c levels and triggered the translocation of Bax to the mitochondria. In addition, the MMP, which is regulated by the Bcl-2 family, was gradually reduced in the sulforaphane-treated cells compared to the control cells (Fig. 5B). These results suggest that sulforaphane-induced apoptosis, which took place by promoting the translocation of Bax to mitochondria, followed by a loss of MMP and release of cytochrome c into the cytosol, results in the activation of caspase-9 and -3, thus leading to apoptosis in the cells. These observations also suggest that sulforaphane induced apoptosis in T24 cells via the mitochondrial pathway.

Evidence has accumulated from many studies that apoptosis associated with endoplasmic reticulum (ER) stress may be responsible for cell death induced by antitumor agents (27,28). ER stress can be characterized by an increase in ER stress-associated molecules. In particular, glucose-regulated protein (GRP) 78 and C/EBP-homologous protein (CHOP) have been considered as vital proteins of ER stress response. To examine whether ER stress was involved in sulforaphane-induced apoptosis, we followed the behavior of these two stress markers in response to sulforaphane treatment. Our data revealed that sulforaphane exposure increased the expression of GRP78 as well as CHOP compared with control (Fig. 4A) in a concentration-dependent manner. These results show that ER stress was involved in sulforaphane-induced apoptosis in T24 cells, as evidenced by upregulation of the expression of ER stress-associated proteins.

Because oxidative stress and ROS generation are directly involved in protease cascades, such as induction of caspases during apoptosis regulation (9,10), we investigated whether the elevated generation of ROS production is related to sulforaphane-induced apoptosis following MMP disruption. To this end, DCFH-DA-based flow cytometric analysis was utilized to measure the amounts of ROS in control and sulforaphane-treated cells. As shown in Fig. 5A, compared with control cells, the production of ROS in sulforaphane-treated cells dramatically increased within 15 min of sulforaphane treatment and peaked at 30 min after treatment; thereafter, the production was attenuated gradually from 1 to 2 h following sulforaphane treatment. In a parallel experiment, pretreatment of the ROS scavenger NAC, along with sulforaphane, drastically decreased ROS generation as compared to the sulforaphane-treated group. Therefore, we proceeded to investigate whether ROS generation was necessary for sulforaphane-induced mitochondrial dysfunction. Fig. 5B shows that the scavenging of ROS by NAC significantly attenuated sulforaphane-induced mitochondrial membrane permeabilization. Furthermore, blockade of ROS generation suppressed sulforaphane-induced cytochrome c release, PARP cleavage, increases of sub-G1 accumulation, and growth inhibition (Fig. 6). These results indicate that ROS play a major role in sulforaphane-induced mitochondrial dysfunction in the T24 cell apoptosis process.

The transcription factor nuclear factor erythroid 2-related factor 2 (Nrf2) is one of the key regulators in the antioxidant response in eukaryotic cells. Under normal physiological conditions, the Nrf2 protein is sequestrated by its cytoplasmic partner, Kelch-like ECH-associated protein 1 (Keap1). ROS induce the release of Nrf2 from Keap1 repression with a subsequent translocation of Nrf2 into the nucleus, where it binds to the antioxidant response element (ARE) for transcription of phase II detoxifying and antioxidant enzymes (29–31). In order to identify the mechanism underlying sulforaphane-induced apoptosis, we assessed the effect of sulforaphane on the Nrf2 pathway. As shown in Fig. 7A, an apparent increase in Nrf2 levels and a reduction in Keap1 protein expression were detected after treatment with sulforaphane. Next, Nrf2 activation was assessed by the accumulation of phosphorylated forms of the Nrf2 protein (p-Nrf2) in the nucleus, and we found that treatment with sulforaphane resulted in a translocation of p-Nrf2 in the nucleus from cytosol (Fig. 7B). Because one of the downstream targets of Nrf2 is the heme oxygenase-1 (HO-1) gene, whose gene product is a sensitive and reliable indicator of cellular stress (32,33), in the next step, we evaluated the effect of sulforaphane on HO-1 protein levels following treatment of T24 cells. As illustrated in Fig. 7A, after treatment of T24 cells with sulforaphane, HO-1 levels increased steadily in a concentration-dependent manner, clearly indicating that sulforaphane treatment activated Nrf2 signaling in T24 cells.