Food and Chemical Toxicology 64 (2014) 157–165

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Sulforaphane induces reactive oxygen species-mediated mitotic arrest and subsequent apoptosis in human bladder cancer 5637 cells Hyun Soo Park a, Min Ho Han b, Gi-Young Kim c, Sung-Kwon Moon d, Wun-Jae Kim e, Hye Jin Hwang b,f, Kun Young Park g, Yung Hyun Choi b,h,⇑ a

School of Korean Medicine, Pusan National University, Yangsan 626-870, Republic of Korea Anti-Aging Research Center & Blue-Bio Industry Regional Innovation Center, Dongeui University, Busan 614-714, Republic of Korea Laboratory of Immunobiology, Department of Marine Life Sciences, Jeju National University, Jeju 690-756, Republic of Korea d School of Food Science and Technology, Chung-Ang University, Ansung 456-756, Republic of Korea e Department of Urology, Chungbuk National University College of Medicine, Cheongju 361-763, Republic of Korea f Department of Food and Nutrition, College of Human Ecology, Dongeui University, Busan 614-714, Republic of Korea g Department of Food and Nutrition, Pusan National University, Busan 609-735, Republic of Korea h Department of Biochemistry, Dongeui University College of Oriental Medicine, Busan 614-052, Republic of Korea b c

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Article history: Received 11 August 2013 Accepted 24 November 2013 Available online 1 December 2013 Keywords: Sulforaphane ROS Mitotic arrest Apoptosis 5637 Cells

a b s t r a c t The present study was undertaken to determine whether sulforaphane-derived reactive oxygen species (ROS) might cause growth arrest and apoptosis in human bladder cancer 5637 cells. Our results show that the reduced viability of 5637 cells by sulforaphane is due to mitotic arrest, but not the G2 phase. The sulforaphane-induced mitotic arrest correlated with an induction of cyclin B1 and phosphorylation of Cdk1, as well as a concomitant increased complex between cyclin B1 and Cdk1. Sulforaphane-induced apoptosis was associated with the activation of caspase-8 and -9, the initiators caspases of the extrinsic and intrinsic apoptotic pathways, respectively, and activation of effector caspase-3 and cleavage of poly (ADP-ribose) polymerase. However, blockage of caspase activation inhibited apoptosis and abrogated growth inhibition in sulforaphane-treated 5637 cells. This study further investigated the roles of ROS with respect to mitotic arrest and the apoptotic effect of sulforaphane, and the maximum level of ROS accumulation was observed 3 h after sulforaphane treatment. However, a ROS scavenger, N-acetyl-L-cysteine, notably attenuated sulforaphane-mediated apoptosis as well as mitotic arrest. Overall, these results suggest that sulforaphane induces mitotic arrest and apoptosis of 5637 cells via a ROS-dependent pathway. Ó 2013 Elsevier Ltd. All rights reserved.

1. Introduction Bladder cancer is the fourth commonest male and ninth commonest female malignant disease in the United States (Abdollah et al., 2013), although the incidence of bladder cancer in Asia is much lower (Cheon et al., 2002; Kakehi et al., 2010). Due to its expanded therapy and the need for lifelong surveillance, bladder cancer is expensive to treat. Despite recent advances in surgical and chemotherapeutic procedures, the 5-year survival rate in patients with invasive and metastatic bladder cancer remains very low (Rosenberg et al., 2005). It is therefore important to develop other effective strategies to improve the survival rate for bladder cancer patients. In light of this, studies on promising dietary ⇑ Corresponding author. Address: Department of Biochemistry, Dongeui University College of Oriental Medicine, Yangjeong-dong San 45, Busanjin-gu, Busan 614-052, Republic of Korea. Tel.: +82 51 850 7413; fax: +82 51 853 4036. E-mail address: [email protected] (Y.H. Choi). 0278-6915/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.fct.2013.11.034

components may offer a new strategy for improvement of bladder cancer prognosis. Recent epidemiologic data indicate that populations with a high dietary intake of cruciferous vegetables have lower incidences of bladder cancer (Liu et al., 2013; Tang et al., 2010; Michaud et al., 1999), which is attributed, in part, to the high content of glucosinolates in many cruciferous vegetables, including broccoli (Tang and Zhang, 2004). Glucosinolates are converted to isothiocyanates by myrosinase, an enzyme which is released when the cell structure of plants has been disrupted through chewing, chopping, or digestion (Shapiro et al., 2001). As a major group of active phytochemicals in cruciferous vegetables, isothiocyanates have shown anticancer activity through the inhibition of carcinogen-activating enzymes, as well as through the induction of carcinogen-detoxifying enzymes, differentiation, cell cycle arrest and apoptosis (Hanlon et al., 2009; Munday et al., 2006). Among isothiocyanates, sulforaphane [1-isothiocyanato-4-(methylsulfinyl)butane] is abundant in broccoli and has been shown to exhibit anti-cancer activities in a

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wide variety of tumors by inducing cell cycle arrest and apoptosis, and by inhibiting metastasis (Cheung and Kong, 2010; Clarke et al., 2008; Gamet-Payrastre, 2006; Lee et al., 2013). This compound has also shown anti-bladder cancer activity in vitro and in vivo; the reported mechanisms of which include modulation of cyclooxygenase-2 expression associated with p38 mitogen activated protein kinase activation in T24 bladder cancer cells (Shan et al., 2010), inhibition of 4-aminobiphenyl-induced DNA damage in RT4 bladder cancer cells and in mouse bladder tissue (Ding et al., 2010), and inhibition of invasion and metastasis by suppressing epithelial-to-mesenchymal transition process (Shan et al., 2013). Many cytotoxic agents and/or DNA damaging agents arrest cell cycling at G1, S or G2/M phase, inducing apoptotic cell death (Naithani et al., 2008; Canavese et al., 2012). Cell cycle checkpoints may function to ensure cells to have time for DNA repair (Schwartz and Shah, 2005; Dean et al., 2012). Therefore, recent studies have offered novel insights into the molecular mechanisms of sulforaphane-induced cell cycle arrest and apoptosis. It has been determined in previous studies that sulforaphane treatment generates reactive oxygen species (ROS) to trigger signal transduction culminating in cell cycle arrest at G2/M phase and/or apoptosis in multiple cancer types (Lee et al., 2012; Singh et al., 2004, 2005; Mi et al., 2011; Cho et al., 2005). We also previously found that cellular ROS generation by sulforaphane plays a pivotal role in the initiation of sulforaphane-triggered apoptotic death in human hepatocellular carcinoma and leukemia U937 cells (Moon et al., 2010; Choi et al., 2008). However, the ROS generation by sulforaphane is tumor cell specific, since normal cell lines were resistant to cell cycle arrest and apoptosis by sulforaphane (Meeran et al., 2010; Kallifatidis et al., 2009; Antosiewicz et al., 2008). Meanwhile, Shan et al. (2006) reported that sulforaphane-induced apoptosis of bladder cancer T24 cells is related to the induction of G1 phase arrest of cell cycle, through up-regulation of cyclin-dependent kinase (Cdk) inhibitor p27 expression. However, sulforaphane treatment leads to accumulation in the G2/M phase of the cell cycle and then apoptotic cell death in other bladder cancer cell lines (RT4, J82 and UMUC3) (Abbaoui et al., 2012). These studies cumulatively indicate that sulforaphane may affect different signaling pathways depending on the cell type or culture conditions used. Nevertheless, the roles of ROS-mediated induction of cell cycle arrest and apoptosis by sulforaphane in human bladder cancer cells has still not been elucidated. Thus, in the present study, the molecular mechanisms of ROS responsible for anti-cancer regulation of sulforaphane were examined in human bladder cancer 5637 cells. Our data demonstrate that sulforaphane induces mitotic arrest, but not G2 phase, and also induces caspase-dependent apoptosis by the accumulation ROS, thereby representing sulforaphane as a promising therapeutic agent for the treatment of bladder cancer.

2. Materials and methods 2.1. Reagents Sulforaphane was purchased from Sigma–Aldrich Chemical Co. (St. Paul, MN) and 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) was obtained from GIBCO-BRL (Gaithersburg, MD). Caspase activity assay kits and enhanced chemiluminescence (ECL) kit were purchased from R&D Systems (Minneapolis, MN) and Amersham (Arlington Heights, IL), respectively. 3-(4,5-dimetylthiazol-2-yl)-2,5-diphenyl-tetrazolium (MTT), 4,6-diamidino2-phenyllindile (DAPI), propidium iodide (PI), 5,50 , 6,60 -tetrachloro-1,10 ,3,30 -tetraethyl-imidacarbocyanine iodide (JC-1) and N-acetyl L-cysteine (NAC) were obtained from Sigma–Aldrich. Pan-caspase inhibitor, z-VAD-fmk, was obtained from Calbiochem (San Diego, CA). 20 ,70 -dichlorofluorescein diacetate (DCFDA) and fluoresceinisothiocyanate (FITC)-Annexin V were purchased from Molecular Probes (Eugene, OR) and PharMingen (SinDiego, CA), respectively. Antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA), Chemicon (Temecula, CA) and Sigma–

Aldrich. Peroxidase-labeled donkey anti-rabbit and sheep anti-mouse immunoglobulin were purchased from Amersham (Arlington Heights, IL). All other chemicals were purchased from Sigma–Aldrich.

2.2. Cell culture and cytotoxicity assay 5637 Cells were purchased from American Type Culture Collection (ATCC, Rockville, MD) and maintained in RPMI-1640 medium supplemented with 10% FBS, 2 mM L-glutamine and penicillin/streptomycin. In order to measure the inhibition of 5637 cell proliferation by sulforaphane, cells were plated in 6-well culture plates (1  105 cells/well) and allowed to adhere overnight, and then treated with different concentrations of sulforaphane for 48 h. After treatment, 0.5 mg/ml MTT solution was added, and the plates were incubated for an additional 3 h at 37 °C. The medium was subsequently removed, and dimethyl sulfoxide (Sigma–Aldrich) was added. Optical density was measured at 540 nm using a microplate spectrophotometer (Dynatech Laboratories, Chantilly, VA).

2.3. Morphological observation of nuclear change After culture with various concentrations of sulforaphane, 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 2.5 lg/ml DAPI solution for 10 min at room temperature. The cells were analyzed using a fluorescence microscope (Carl Zeiss, Oberkochen, Germany).

2.4. Cell cycle analysis Following treatment with sulforaphane, cells were trypsinized, washed with PBS, and fixed in 75% ethanol at 4 °C for 30 min. The cells were suspended in cold 50 l/ml PI solution and 0.1% Triton X-100 (Sigma–Aldrich), and incubated at room temperature in the dark for 30 min. A FACScan flow cytometry system (Becton Dickinson, San Jose, CA) was used for performance of flow cytometry analysis.

2.5. Annexin-V staining To analyze apoptosis, cells were treated with the indicated concentrations of sulforaphane for 48 h, and apoptosis was analyzed by staining phosphatidylserine translocation with 5 lM FITC-annexin V for 30 min at room temperature in the dark according to the manufacturer’s instructions. Flow cytometric analysis was performed as described previously (Li and Gao, 2013).

2.6. Total protein extraction, immunoprecipitation and Western blot assay 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]. For Western blot assay, the total proteins were separated by SDS–polyacrylamide gel and transferred to a nitrocellulose membrane (Schleicher & Schuell, Keene, NH) by electroblotting. After being blocked with blocking solution (1% BSA in PBS plus 0.05% Tween-20) at room temperature for 1 h, each membrane was incubated for 2 h at room temperature or overnight at 4 °C with primary antibodies, and then the membranes were probed for 1 h at room temperature with the peroxidase-labeled secondary antibodies. Detection was performed by the ECL Western blotting detection kit according to the manufacturer’ instructions. In a parallel experiment, interaction of cyclin B1 with Cdk1 was analyzed by immunoprecipitation and subsequent Western blot analysis. For this study, lysates representing 500 lg of protein were precleared by the addition of protein A-Sepharose beads (Sigma–Aldrich), followed by rocking at 4 °C for 1 h. To the precleared lysate, 2 lg of ant-Cdk1 antibody was added and then rocked at 4 °C for 4 h. The antibody-associated complexes were collected by adding protein A-Sepharose beads p and incubating for a further 2 h at 4 °C with constant rotation. The immunoprecipitates were washed three times with lysis buffer and, after removal of as much liquid as possible, were resuspended in SDS sample buffer for Western blot analysis using anti-cyclin B1 and Cdk1 antibodies.

2.7. Analysis of caspase-3, -8 and -9 activities To evaluate caspase activity, cell lysates were prepared after treatment with sulforaphane. Assays were performed in 96-well plates by incubating 20 lg cell lysates in 100 mL reaction buffer containing 5 lM of colorimetric tetrapeptides [Asp-Glu-Val-Asp (DEAD) for caspase-3; Ile-Glu-Thr-Asp (IETD) for caspase-8; Leu-Glu-His-Asp (LEHD) for caspase-9] labeled with p-nitroaniline (pNA) at 37 °C for 2 h according to the manufacturer’s protocol. Thereafter, the optical density of the reaction mixture was measured spectrophotometrically at a wavelength of 405 nm using a microplate spectrophotometer.

H.S. Park et al. / Food and Chemical Toxicology 64 (2014) 157–165 2.8. Measurement of ROS 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 20 lM sulforaphane for various periods. Later, the cells were incubated with 10 mM DCFDA for 20 min at 37 °C in the dark. The ROS production in the cells was monitored with a flow cytometer using the Cell-Quest pro software (Kim et al., 2012).

2.9. Measurement of mitochondrial membrane potential (MMP, Wm) The values of MMP were determined using the dual-emission potential-sensitive probe, JC-1. Briefly, the sulforaphane-treated cells were collected and incubated with 10 lM 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 10,000 cells per sample was measured at 488 nm and 575 nm using a flow cytometer.

2.10. Statistical analysis All values are presented as mean ± standard deviation (SD). We assessed comparisons between groups by one-way analysis of variance (Dunnett’s t-test) and Student’s t-test. P values 60.05 were considered statistically significant.

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3. Results 3.1. Sulforaphane-induced growth inhibition is associated with the induction of apoptosis in 5637 cells In order to determine whether sulforaphane-mediated suppression of 5637 cell proliferation correlates with apoptosis induction, 5637 cells were exposed to different concentrations of sulforaphane and, after 48 h incubation, cell viability was measured by MTT assay. Our results show that exposure of the 5637 cells to sulforaphane decreased cellular viability in a time-dependent manner (Fig. 1A). Under the same conditions, cell morphology was assessed by DAPI staining in order to elucidate whether sulforaphane inhibits cell growth through the induction of apoptosis. As can be seen in Fig. 1B, the nuclear structure of control cells remained intact, while nuclear chromatin condensation and fragmentation, characteristic of apoptosis, was observed in cells treated with 20 lM sulforaphane after 24 h. Furthermore, to measure apoptotic cell death upon sulforaphane treatment, we stained cells for annexin V. As shown in Fig. 1C, after treatment with 20 lM sulforaphane for

Fig. 1. Inhibition of cell viability and induction of apoptosis by sulforaphane treatment in 5637 cells. Cells were seeded at 1  105 cells/ml and were incubated for 12 h. The cells were treated with 20 lM sulforaphane for the indicated times. Cell viability was determined by MTT assay. (B) The cells were fixed and stained with DAPI to visualize DNA. Cells were analyzed by fluorescence microscopy (400). (C) The cells were stained with annexin-V and the percentages of apoptotic cells were then analyzed using flow cytometric analysis. (A and C) Each point represents the mean ± S.D. of three independent experiments. Significance was determined using Student’s t-test (p < 0.05 vs. untreated control).

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24 h and 48 h, the percentages of apoptotic cells increased from 1.9% to 11.7% and 27.6%, respectively. 3.2. Sulforaphane induces cell cycle arrest at the mitotic phase in 5637 cells To evaluate the cell cycle distribution of 5637 cells after sulforaphane treatment, the DNA content was measured by flow cytometry. As shown in Fig. 2A, in control cultures, 15.1% of cells were in the G2/M phase, however, in sulforaphane-treated cells, this percentage was significantly increased (28.7% at 24 h, 35.0% at 36 h and 47.5% at 48 h). Since flow cytometric analysis of DNA content does not allow us to distinguish between G2 and M arrest, we monitored the expressions of M phase progression key proteins in order to establish the precise cell cycle phase in which 5637 cells accumulated. Because, histone H3 is phosphorylated at serine 10 during mitosis by aurora kinase and the phosphorylation status of H3 is considered a marker of mitosis (Hendzel et al., 1997; Murnion et al., 2001), we examined the phosphorylation status of histone H3 (Ser 10) and found that sulforaphane treatment significantly increases the phosphorylation level of histone H3 (Fig. 2B). In addition, sulforaphane treatment resulted in a remarkable increase in the accumulation of cyclin B1 and formation of cyclin B1/Cdk1 complex (Fig. 2C), which is associated with the increased status of phosphorylated levels of Cdk1 at Thr161. These results indicated that sulforaphane treatment caused an arrest at mitosis, but not at the G2 phase, in 5637 cells. 3.3. Sulforaphane induces apoptosis through caspase-dependent cascade in 5637 cells To investigate whether sulforaphane induced apoptosis through the cascade-dependent pathway, immunoblotting and colorimetric caspase activity assays were conducted. Although we could not detect the activated cleavage forms of caspase-8 and -9 (initiator caspases of extrinsic and intrinsic apoptotic pathways, respec-

tively), their pro-forms were concentration-dependently downregulated and their activities were significantly increased in response to sulforaphane (Fig. 3). The increase in apoptosis was also accompanied by the activation of caspase-3, an effector caspase, and cleavage of poly (ADP-ribose) polymerase (PARP), which, has been identified as a substrate for caspase-3. The contribution of the caspase pathway to sulforaphane-induced apoptosis in 5637 cells was next investigated by means of z-VAD-fmk, a pan-caspase inhibitor. For this study, cells were pre-incubated for 2 h with z-VAD-fmk before the addition of sulforaphane, and were then incubated for 48 h. As shown in Fig. 4, z-VAD-fmk pre-treatment strongly reduced sulforaphane-induced apoptotic events, such as chromatin condensation and PARP cleavage, and concomitantly prevented sulforaphane-induced growth inhibition. These results suggest of the involvement of caspase activation in the induction of apoptosis of 5637 cells by sulforaphane treatment. 3.4. Sulforaphane induces apoptosis through ROS-dependent disruption of mitochondrial membrane integrity in 5637 cells To examine the roles of ROS and mitochondria in sulforaphaneinduced apoptosis, flow cytometric analyses were performed using DCFDA and JC-1 dyes, respectively, in 5637 cells treated with sulforaphane. The results indicated that ROS generation was first detected at 30 min, and extension of sulforaphane treatment to 1 h and 3 h resulted in ROS production which was 8.5 and 10.7 times that of the control, respectively (Fig. 5A). Sulforaphane also significantly increased MMP loss in a dose-dependent manner up to 48 h in 5637 cells (Fig. 5B), indicating that sulforaphane induces mitochondrial membrane depolarization. To test the involvement of ROS in mitochondrial dysfunction of sulforaphane-induced apoptosis, the ROS scavenger NAC was utilized. Data summarized in Fig. 5C and D show that NAC pre-treatment significantly inhibited ROS generation as well as MMP loss by sulforaphane. Furthermore, blockade of ROS generation suppressed sulforaphane-mediated chromatin condensation, increases of

Fig. 2. Mitotic arrest of 5637 cells by sulforaphane treatment. (A) After treatment with sulforaphane for the indicated times, cells were harvested and 10,000 events were analyzed for each sample. DNA content is represented on the x-axis and the number of cells counted is represented on the y-axis. Each point represents the mean of two independent experiments. (B) Cells treated with sulforaphane were lysed for protein extraction. Samples (30–50 lg protein/lane) were subjected to SDS–polyacrylamide gels and Western blotting for the detection of the indicated proteins. (C) After treatment with or without 20 lM sulforaphane, total cell lysates were immunoprecipitated with anti-Cdk1 antibody, separated on 10% SDS–polyacrylamide gels, and transferred to nitrocellulose. The levels of cyclin B1 and Cdk1 proteins were detected with anti-cyclin B1 and anti-Cdk1 antibodies, respectively, and ECL detection. (IP, immunoprecipitation) Actin was used as an internal control.

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Fig. 3. Activation of caspases and degradation of PARP proteins by sulforaphane in 5637 cells. (A) Cells were treated with 20 lM sulforaphane for the indicated times. The cells were lysed, and cellular proteins were then separated by SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. The membranes were probed with the indicated antibodies. Proteins were visualized using an ECL detection system. Actin was used as an internal control. (B) The cells were lysed, and aliquots (50 lg protein) were assayed for in vitro caspase-3, -8, and -9 activity using DEVD-pNA, IETD-pNA, and LEHD-pNA as substrates, respectively. The released fluorescent products were measured. Data are expressed as mean ± SD of three independent experiments. Significance was determined by Student’s t-test (p < 0.05 vs. untreated control).

sub-G1 accumulation, PARP cleavage and growth inhibition (Fig. 6). The data indicate that ROS-dependent mitochondrial disruption plays a critical role in sulforaphane-induced apoptosis in 5637 cells. 3.5. Sulforaphane induces mitotic arrest through ROS-dependent processes in 5637 cells We finally investigated whether ROS generation also plays an important role in mitotic arrest induced by sulforaphane. When cells were pre-treated with NAC and then treated with sulforaphane, the increased levels of cyclin B1, Cdk1 and phospho-histone H3 were much lower than those of the sulforaphane-treated cells (Fig. 7A), which was concomitant with the inhibition of sulforaphane-induced accumulation of G2/M cells (Fig. 7B). The results reveal that sulforaphane exerts its anticancer cytotoxicity through a ROS-dependent mitotic arrest in 5637 cells. 4. Discussion In recent years, substantial knowledge has been obtained as to how sulforaphane induces cell cycle arrest and apoptosis in human cancer cells. Previous studies have demonstrated that sulforaphane-reduced cancer cell viability was associated with the induction of G2/M arrest and/or apoptosis through ROS-generation (Lee et al., 2012; Singh et al., 2004, 2005; Mi et al., 2011; Cho et al.,

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2005). However, with no previous report of on the roles of ROS induced by sulforaphane in the treatment of human bladder cancer cells, we employed a functional study to investigate the meaning of ROS generation of this agent using the 5637 bladder cancer cell line. Our data showed that sulforaphane could induce mitotic arrest, but not G2 phase, and caspase-dependent apoptosis in 5637 cells. We further found that intracellular ROS levels were increased by sulforaphane, and that ROS play a pivotal role in the initiation of sulforaphane-triggered growth arrest and apoptotic death in 5637 cells. Cell cycle dysregulation and apoptosis resistance are hallmarks of tumor cells. Because, many effective anti-cancer agents kill proliferating cancer cells by arresting cell proliferation and inducing apoptosis, the regulation of factors which mediate critical events between these two processes may be a useful anti-tumor target. Cell cycle progression is critically regulated by a sequential activation of Cdks, the activities and specificities of which are determined by phosphorylation of their corresponding catalytic subunits and by their associations with Cdks inhibitors and cyclins, which are differentially expressed during the cell cycle (Canavese et al., 2012; Schwartz and Shah, 2005). Cell cycle arrest allows cells time for DNA repair before entering the S-phase for DNA replication and the M phase for mitosis, which is critical for cells to maintain their genetic integrity. Failure to repair DNA damage would cause a plethora of mutations, and eventually cell death through apoptosis. As a stage of the cell cycle, mitosis is a continuous process to separate the replicated chromosomes into two daughter cells, through prophase, metaphase, anaphase and telophase. Indeed, increases in cyclin B1 expression and the formation of active complex between cyclin B1, with its catalytic subunit Cdk1, a major M-phase kinase, are the rate-limiting steps of mitosis. Cyclin B1/Cdk1 complex is held in an inactive state during interphase by the phosphorylation of Thr14 and Tyr15 residues of Cdk1 subunit (De Souza et al., 2000; Atherton-Fessler et al., 1994). To allow mitotic entry and progression to metaphase, the complex is then activated through the dephosphorylation of these two negative regulatory sites, and concomitant phosphorylation of Cdk1 subunit at Thr161 (Nigg, 2001). However, cyclin B1 degradation by proteasome pathway and inactivation of Cdk1 drives mitotic exit and cytokinesis (Clute and Pines, 1999; Josefsberg et al., 2001). Furthermore, changes in the phosphorylation status of histone H3 are known to occur in mitotic phases (Hendzel et al., 1997; Murnion et al., 2001). While phosphorylation of histone H3 Ser10 is maintained throughout metaphase, dephosphorylation of this site, beginning in anaphase and completed in telophase, is required for further progression of mitotic division. To determine whether sulforaphane induced cell cycle arrest occurs at mitosis or at the G2 phase, we examined the phosphorylation status of histone H3, a hallmark of mitosis, and immunoblotting data revealed a markedly corresponding increase in the levels of phosphorylated histone H3 after sulforaphane treatment (Fig. 2B). As demonstrated in Figs. 2B and C, sulforaphane also up-regulated the expression of cyclin B1, and Thr161-phosphorylated Cdk1, together with enhancing cyclin B1/Cdk1 complex. Consistent with the previous reports in lung, prostate and breast cancer cells, and myeloma, (Xiao et al., 2012; Jakubikova et al., 2011; Mi et al., 2008; Herman-Antosiewicz et al., 2007; Jackson and Singletary, 2004), these results indicate that treatment with sulforaphane causes mitotic blockage, but not in the G2 phase, in 5637 cells. Apoptosis is a highly regulated physiologic process which allows for the reduction of harmful cells during development, tissue homeostasis and disease. Apoptosis is classified into either caspase-dependent or caspase-independent mechanisms. The caspase-dependent pathway can be further divided into two key pathways: the death receptor-mediated pathway (extrinsic pathway) and the mitochondria-initiated apoptotic pathway

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Fig. 4. Effects of a pan-caspase inhibitor on the apoptosis induced by sulforaphane in 5637 cells. (A) Cells were incubated in the presence or absence of 100 lM z-VED-fmk for 2 h before being exposed to 20 lM sulforaphane. After 48 h of incubation, the cells were fixed, stained with DAPI solution, and then observed under a fluorescent microscope (original magnification, 400). (B) Cells grown under the same conditions as (A) were lysed for protein extraction. Samples were subjected to SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. The membranes were probed with anti-PARP and anti-actin antibodies. Proteins were visualized using an ECL detection system. Actin was used as an internal control. (C) Cells were stained with annexin-V, and the percentages of apoptotic cells were then analyzed using flow cytometric analysis. (D) Cell viability was analyzed using MTT assay. The data are expressed as means ± SD of triplicate samples. The significance was determined by Student’s t-test (p < 0.05 vs. untreated control).

Fig. 5. Increase in ROS generation and loss of MMP by treatment of 5637 cells with sulforaphane. (A and B) Cells were seeded in 6-well plates at an initial density of 1  105 cells per well. After 24 h of stabilization, the cells were challenged with 20 lM sulforaphane for various periods. (A) The medium was discarded and cells were incubated at 37 °C in the dark for 20 min with new culture medium containing 10 lM DCFDA. ROS generation was measured using a flow cytometer. (B) The cells were collected and incubated with 10 lM JC-1 for 20 min at 37 °C in the dark. The cells were then washed once with PBS and the values of MMP were analyzed by flow cytometry. (C and D) Cells were pretreated with 10 mM NAC for 2 h prior to 20 lM sulforaphane treatment. (C) After 3 h of incubation, ROS generation was detected using DCFDA. (D) The loss of MMP was investigated using JC-1 following 48 h treatment with 20 lM sulforaphane. The results are expressed as the mean ± SD of three independent experiments. The statistical significance of the results was analyzed by Student’s t-test (p < 0.05 vs. untreated control; #p < 0.05 vs. sulforaphane-treated cells).

(intrinsic pathway). Both extrinsic and intrinsic pathways involved activation of executioner caspases (-3 and -7) which is important for inducing downstream DNA cleavage molecules (Dean et al.,

2012; King and Cidlowski, 1995). Mitochondria play a central role in both extrinsic and intrinsic apoptotic pathways by releasing cytochrome c into the cytoplasm, leading to the activation of the

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Fig. 6. Induction of ROS-dependent apoptosis by sulforaphane in 5637 cells. (A) Cells were pretreated with 10 mM NAC 2 h prior to 20 lM sulforaphane treatment. After 48 h of incubation, the cells were fixed, stained with DAPI solution, and then observed under a fluorescent microscope (original magnification, 400). (B) Total proteins were subjected to SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. Proteins were visualized using an ECL detection system. Actin was used as an internal control. (C) Cells were stained with annexin-V, and the percentages of apoptotic cells were then analyzed using flow cytometric analysis. (D) Cell viability was analyzed using MTT assay. The data are expressed as the means ± SD of triplicate samples. Significance was determined by Student’s t-test (p < 0.05 vs. untreated control).

Fig. 7. Induction of ROS-dependent mitotic arrest by sulforaphane in 5637 cells. (A) Cells were pretreated with 10 mM NAC 2 h prior to 20 lM sulforaphane treatment. After 48 h of incubation, the cells were lysed and cellular proteins were separated by SDS–polyacrylamide gels and transferred onto nitrocellulose membranes. The membranes were probed with the indicated antibodies. Proteins were visualized using an ECL detection system. Actin was used as an internal control. (B) Cells were collected and their DNA was evaluated by flow cytometry. Each point represents the mean of two independent experiments.

caspase-cascade system (Antico Arciuch et al., 2012; Caroppi et al., 2009). Here, we showed that sulforaphane treatment led to activation of caspase-8 and -9 (Fig. 3), which are critical in extrinsic and intrinsic pathways, respectively. Nevertheless, the reason why suforaphane increases capase-8 known via death receptor-mediated pathway is not understood in this study. Whether sulforaphane-induced caspase-8 activation is linked with G2/M phase

will be studied. In addition, significant change in MMP could be found in cells treated with sulforaphane in a time-dependent manner (Fig. 5B), suggesting that sulforaphane caused a significant change in the transmembrane potential, and subsequently increased the activity of caspase-9. Sulforaphane also activated effector caspase-3, followed by the cleavage of PARP, a nuclear enzyme which is involved in DNA repair in response to various stresses (Fig. 3). Moreover, the pan-caspase inhibitor z-VAD-fmk suppressed sulforaphane-induced apoptosis and growth inhibition (Fig. 4). Although further studies are needed, this information indicates that ROS required, at least in part, for both the mitotic arrest and apoptosis induced by sulforaphane treatment in 5637 cells The redox state of cells has been shown to be involved in cell cycle regulation and cell death/survival (Schumacker, 2003; Antico Arciuch et al., 2012). The results of several recent studies suggest the possibility that ROS-mediated oxidative DNA damage and depolarization of mitochondrial membranes play an important role in sulforaphane-induced cancer cell death and cell cycle arrest (Lee et al., 2012; Singh et al., 2004, 2005; Mi et al., 2011; Cho et al., 2005). However, the oxidative stress in human bladder cancer cells caused by sulforaphane lacks a mechanistic study. In this study, to examine whether ROS are related to mitotic arrest and apoptosis induced by sulforaphane, we measured changes of DCFDA fluorescence in 5637 cells exposed to sulforaphane. As indicated in Fig. 5A, we found that intracellular ROS levels were markedly increased within 30 min by sulforaphane treatment in 5637 cells, and the quenching of ROS generation with antioxidant NAC, a widely used ROS scavenger, conferred significant protection against sulforaphane-elicited mitochondrial dysfunction and apoptosis (Figs. 5D and 6), demonstrating that ROS play a pivotal role in the initiation of sulforaphane-triggered apoptotic death in 5637 cells. Furthermore, the increase in the expression of cyclin B1 and the phosphorylation of Cdk1 on treatment with sulforaphane was reversed by NAC pretreatment. We also found that NAC inhibited both the phosphorylation of histone H3 and mitotic arrest in-

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duced by sulforaphane (Fig. 7). Interestingly, no apoptotic G1 phase was seen in response to sulforaphane when sulforaphane-induced G2/M phase disappeared in the presence of NAC, indicating that sulforaphane-induced G2/M arrest and apoptosis is mediated by ROS generation. Here, we cannot rule out the possibility that G2/M arrest and apoptosis are separately regulated because ROS inhibitor inhibits both G2/M arrest and apoptosis. Presumed previously, if G2/M arrest induces delayed apoptosis, the inhibition of G2/M arrest might promote early apoptosis in response to sulforaphane. We need to conduct further studies to evaluate the relationship between sulforaphane-induced G2/M arrest and apoptosis Collectively, these findings may suggest that the generation of ROS is required for both mitotic arrest and apoptosis induced, at least in a part, by sulforaphane treatment of 5637 cells. 5. Conclusions In summary, mitotic arrest by sulforaphane treatment likely mediated the ROS-dependent activation of cyclin B1/Cdk1 complex and phosphorylation of histone H3. Apoptosis by sulforaphane treatment was induced via mitochondrial disruption, including both that which occurred through the caspase-dependent pathway, and ROS generation. Although in vivo studies are required to evaluate the biological efficacy of treatment with sulforaphane, our results describe a potent molecular mechanism for sulforaphane-induced mitotic arrest in the 5637 human bladder cancer line, by sequentially triggering apoptosis through ROS accumulation. Conflict of Interest The authors declare that there are no conflicts of interest. Acknowledgements This research was supported by Grants from the Globalization of Korean Foods R&D Program (912001-1), funded by the Ministry of Food, Agriculture, Forestry and Fisheries and the National Research Foundation of Korea (NRF) Grant funded by the Korea government (2008-0062611), Republic of Korea. References Abbaoui, B., Riedl, K.M., Ralston, R.A., Thomas-Ahner, J.M., Schwartz, S.J., Clinton, S.K., Mortazavi, A., 2012. Inhibition of bladder cancer by broccoli isothiocyanates sulforaphane and erucin: characterization, metabolism, and interconversion. Mol. Nutr. Food Res. 56, 1675–1687. Abdollah, F., Gandaglia, G., Thuret, R., Schmitges, J., Tian, Z., Jeldres, C., Passoni, N.M., Briganti, A., Shariat, S.F., Perrotte, P., Montorsi, F., Karakiewicz, P.I., Sun, M., 2013. Incidence, survival and mortality rates of stage-specific bladder cancer in United States: a trend analysis. Cancer Epidemiol. 37, 219–225. Antico Arciuch, V.G., Elguero, M.E., Poderoso, J.J., Carreras, M.C., 2012. Mitochondrial regulation of cell cycle and proliferation. Antioxid. Redox. Signal. 16, 1150– 1180. Antosiewicz, J., Ziolkowski, W., Kar, S., Powolny, A.A., Singh, S.V., 2008. Role of reactive oxygen intermediates in cellular responses to dietary cancer chemopreventive agents. Planta Med. 74, 1570–1579. Atherton-Fessler, S., Liu, F., Gabrielli, B., Lee, M.S., Peng, C.Y., Piwnica-Worms, H., 1994. Cell cycle regulation of the p34cdc2 inhibitory kinases. Mol. Biol. Cell 5, 989–1001. Canavese, M., Santo, L., Raje, N., 2012. Cyclin dependent kinases in cancer: potential for therapeutic intervention. Cancer Biol. Ther. 13, 451–457. Caroppi, P., Sinibaldi, F., Fiorucci, L., Santucci, R., 2009. Apoptosis and human diseases: mitochondrion damage and lethal role of released cytochrome C as proapoptotic protein. Curr. Med. Chem. 16, 4058–4065. Cheon, J., Kim, C.S., Lee, E.S., Hong, S.J., Cho, Y.H., Shin, E.C., Lee, W.C., Yoon, M.S., 2002. Survey of incidence of urological cancer in South Korea: a 15-year summary. Int. J. Urol. 9, 445–454. Cheung, K.L., Kong, A.N., 2010. Molecular targets of dietary phenethyl isothiocyanate and sulforaphane for cancer chemoprevention. AAPS J. 12, 87– 97.

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