Hispolon induces apoptosis in human gastric cancer cells through a ROS-mediated mitochondrial pathway

Free Radical Biology & Medicine 45 (2008) 60–72

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Free Radical Biology & Medicine j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / f r e e r a d b i o m e d

Original Contribution

Hispolon induces apoptosis in human gastric cancer cells through a ROS-mediated mitochondrial pathway Wei Chen a, Zhao Zhao a, Ling Li b, Bin Wu c, Shi-fei Chen a, Hong Zhou a, Yong Wang a, Yong-Quan Li a,⁎ a b c

College of Life Science, Zhejiang University, Hangzhou 310058, China Cancer Institute, The Second Affiliated Hospital, Zhejiang University School of Medicine, Hangzhou 310009, China College of Pharmaceutical Sciences, Zhejiang University, Hangzhou 310058, China

A R T I C L E

I N F O

Article history: Received 5 November 2007 Revised 29 February 2008 Accepted 11 March 2008 Available online 29 March 2008 Keywords: Hispolon ROS GSH Apoptosis Mitochondria Gastric cancer Free radicals

A B S T R A C T Severe side effects and complications such as gastrointestinal and hematological toxicities because of current anticancer drugs are major problems in the clinical management of gastric cancer, which highlights the urgent need for novel effective and less toxic therapeutic approaches. Hispolon, an active polyphenol compound, is known to possess potent antineoplastic and antiviral properties. In this study, we investigated the efficacy of hispolon in human gastric cancer cells and explored the cell death mechanism. Hispolon induced ROS-mediated apoptosis in gastric cancer cells and was more toxic toward gastric cancer cells than toward normal gastric cells, suggesting greater susceptibility of the malignant cells. The mechanism of hispolon-induced apoptosis was that hispolon abrogated the glutathione antioxidant system and caused massive ROS accumulation in gastric cancer cells. Excessive ROS caused oxidative damage to the mitochondrial membranes and impaired the membrane integrity, leading to cytochrome c release, caspase activation, and apoptosis. Furthermore, hispolon potentiated the cytotoxicity of chemotherapeutic agents used in the clinical management of gastric cancer. These results suggest that hispolon could be useful for the treatment of gastric cancer either as a single agent or in combination with other anticancer agents. © 2008 Elsevier Inc. All rights reserved.

Gastric cancer is the fourth most frequent cancer and the second leading cause of cancer-related death in the world [1,2]. This aggressive disease is a major global threat to public health, with an estimated 934,000 new cases in 2002 (8.6% of new cancer cases). Almost twothirds of the cases occur in developing countries and 42% in China alone [2]. Of patients presenting with earlier stages of the disease, more than 50% undergo surgery, but even after a curative resection, 60% of these patients eventually relapse and die due to their disease [3,4]. The high risk of relapse after surgery has led to a search for strategies to prevent relapse and to improve survival for gastric cancer patients. Chemotherapy in advanced gastric cancer is an important issue because the majority of patients with gastric cancer develop metastases during the course of their disease [5,6]. However, severe side effects and complications such as gastrointestinal and hematological toxicities because of current anticancer drugs are major problems in the clinical

Abbreviations: PL, Phellinus linteus; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; ROS, reactive oxygen species; MMP, mitochondrial membrane potential; GSH, reduced glutathione; GPX, glutathione peroxidase; PI, propidium iodide; DOX, doxorubicin; 5-FU, 5-fluorouracil; CCCP, carbonyl cyanide m-chlorophenylhydrazone; CIS, cisplatin; MMC, mitomycin C; DCFH-DA, dichlorodihydrofluorescein diacetate; NAC, N-acetyl-L-cysteine; Rh123, rhodamine 123; CSA, cyclosporin A; TFP, trifluoperazine; BSO, buthionine sulfoximine; HEt, hydroethidine; NAO, nonyl acridine orange; DAF, DAF-FM diacetate. ⁎ Corresponding author. Fax: +86 571 88208569. E-mail address: [email protected] (Y.-Q. Li). 0891-5849/$ – see front matter © 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.freeradbiomed.2008.03.013

setting [7]. In particular, the side effects of drugs might be fatal in older patients or in immunocompromised patients, which highlights the urgent need for novel effective and less toxic therapeutic approaches. Chinese medicinal herbs have been used as traditional remedies for thousands of years, and their importance in the prevention and treatment of cancer is becoming increasingly apparent. Recently, experimental and clinical studies reported that several Chinese herbs and their active components were effective in treating and preventing gastric cancer [8–12]. Thus, there is reason to consider the use of other medicinal herbs and active components, perhaps in combination with existing therapies, in the treatment of gastric cancer. Phellinus linteus (PL), a traditional medicinal mushroom, has been widely used in China, Korea, and Japan for the treatment of various diseases, including gastroenteric disorder, peptic ulcers, lymphatic disease, and various cancers. Recent studies have shown that PL has anti-inflammatory, antimutagenicity, and antibacterial effects; stimulates immunity; and inhibits tumor growth and metastasis [13–17]. In our previous study, we found that hispolon, a polyphenol compound isolated from PL, induced apoptosis in human epidermoid KB cells [18]. Other studies also showed that hispolon has antiviral, antiproliferative, and immunomodulatory activities [19–21]. However, there is little information available concerning the ability of hispolon to inhibit gastric cancer. In the present study, we investigated the effects of hispolon on human gastric cancer cells and further examined the cell death mechanism. Our observations demonstrate that gastric cancer cells

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are more susceptible to the cytotoxic effects of hispolon than are normal gastric cells. Hispolon-induced cytotoxicity was characterized by ROS-mediated apoptosis, which was associated with massive ROS accumulation and alterations in the mitochondrial membrane potential. Moreover, hispolon potentiated the cytotoxicity of chemotherapeutic agents used in the clinical management of gastric cancer. These preclinical studies suggest that hispolon could be useful for the treatment of gastric cancer. Material and methods Cell culture and reagents Human gastric cancer cell lines SGC-7901, MGC-803, and MKN-45 were obtained from the Institute of Biochemistry and Cell Biology (Chinese Academy of Sciences). The human normal gastric epithelial cell line GES-1 was obtained from the Cancer Research Institute of Beijing, China [22]. All cell lines were routinely cultured in RPMI 1640 medium (Gibco) containing 10% fetal bovine serum (Gibco), 100 units/ ml penicillin, and 100 units/ml streptomycin in a humidified cell incubator with an atmosphere of 5% CO2 at 37°C. Hispolon was synthesized as previously described and its purity was established on the basis of the spectral (1H, 13C NMR and mass) data [23]. Hispolon was dissolved in DMSO and diluted to the desired concentrations before use, with the concentration of DMSO kept below 0.1% in treated groups. Doxorubicin (DOX), 5-fluorouracil (5FU), cisplatin (CIS), mitomycin C (MMC), 3-(4,5-dimethylthiazol-2-yl)2,5-diphenyltetrazolium bromide (MTT), rhodamine 123 (Rh123), carbonyl cyanide 3-chlorophenylhydrazone (CCCP), dichlorodihydrofluorescein diacetate (DCFH-DA), cyclosporin A (CSA), trifluoperazine (TFP), N-acetylcysteine (NAC), catalase, and buthionine sulfoximine (BSO) were purchased from Sigma. Hydroethidine (HEt), nonyl acridine orange (NAO), and DAF-FM diacetate (DAF) were obtained from Molecular Probes. The pancaspase inhibitor z-VAD-fmk was from R&D. All other chemicals were of the highest purity available. Cell viability assay Cell viability was measured by the MTT method as previous described, with some modifications [24,25]. Briefly, cells were grown in 96-well microtiter plates for drug treatment. When incubated for the indicated times, cells were incubated with MTT (0.5 mg/ml) for 4 h. The formazan precipitate was dissolved in 150 μl DMSO, and the absorbance was detected at 490 nm with a Model ELX800 microplate reader (Bio-Tek Instruments). Each test was performed in triplicate experiments. Apoptosis assays Apoptotic rates were analyzed by flow cytometry using an annexin V–FITC/PI kit (Sigma) according to the manufacturer's instructions. Briefly, cells were treated with hispolon for 24 h, and then 1 × 106 cells were harvested, washed twice with ice-cold PBS, and evaluated for apoptosis by double staining with annexin V–FITC and propidium iodide in binding buffer using a FACSCalibur flow cytometer (BD Biosciences). To detect DNA strand breaks, a TUNEL assay was performed using an APO-BRDU kit (BD Biosciences). In brief, cells were fixed and permeabilized by 4% paraformaldehyde and 70% ethanol, followed by incubation with a mixture of FITC–dUTP and TdT for 1 h at 37°C. Stained cells were analyzed with a FACSCalibur flow cytometer. To evaluate activation of caspase-3, flow-cytometric analysis was done using the FITC-Conjugated Monoclonal Active Caspase-3 Antibody Apoptosis Kit I according to the manufacturer's instructions (BD Biosciences).

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Caspase assays Caspase-3, -8, and -9 activity was measured using a colorimetric assay kit (Biovision) according to the manufacturer's instructions. Briefly, cell lysate from 1 × 106 cells was incubated at 37°C for 2 h with 200 μM DEVD-pNA (caspase-3 substrate), IETD-pNA (caspase-8 substrate), or LEHD-pNA (caspase-9 substrate). Samples were read at 405 nm in a microplate reader (Bio-Tek Instruments) and expressed as fold increase on the basal level (DMSO-treated cells). In the caspase inhibitor assay, cells were treated with 0–15 μg/ml hispolon for 24 h, with or without 100 μM z-VAD-fmk pretreatment for 2 h, and then apoptosis was determined by TUNEL assay as described above. Determination of cellular reactive oxygen species Cellular ROS contents were measured by incubating the control or drug-treated cells with 10 μM DCFH-DA at 37°C for 30 min. After incubation with the fluorochrome, cells were washed with PBS and immediately analyzed by flow cytometry. O·-2 and NO were measured by flow cytometry using HEt and DAF as described [26,27]. Briefly, after treatment, cells were incubated with 5 μM DAF-FM diacetate (NO) or 1 μM HEt (O·-2 ) for 30 min and then washed with PBS and immediately analyzed by flow cytometry. Detection of oxidative damage to mitochondrial membranes Mitochondrial membrane lipid peroxidation was detected by measuring the oxidation of cardiolipin, using NAO as a fluorochrome [28]. Briefly, after treatment, cells were collected and incubated with 100 nM NAO for 15 min and analyzed by flow cytometry. The mitochondrial membrane potential (MMP) was measured according to the previously reported methods with some modifications [29,30]. Briefly, after treatment, cells were collected and incubated with 10 μg/ ml Rh123 for 30 min and analyzed by flow cytometry. The decoupling agent CCCP was used as a positive control to induce membrane depolarization. Analysis of cellular GSH and its export to the culture medium A glutathione assay kit (Sigma) was used to measure total cellular glutathione. Briefly, after treatment, cell extracts were prepared according to the manufacturer's instructions. Then total GSH was detected by measuring the product of 5-thio-2-nitrobenzoic acid by colorimetric analysis at 412 nm. The cellular GSH contents were calculated using the standard curve generated in parallel experiments. To determine the effect of hispolon on GSH exported from cells to the culture medium, cells were plated at equal densities and cultured for 24 h. The medium was then replaced with serum-free medium with or without hispolon as indicated and incubated for 1–6 h, and the culture medium was removed for GSH assay as described above. Cellular GPX enzyme activity assay Cellular glutathione peroxidase activity was determined using continuous spectrophotometric rate determination as previous described [31]. Briefly, after treatment, equal amounts of protein extracts from control and hispolon-treated cells were added to a reaction mixture containing 2 mM GSH, 0.1 U glutathione reductase, and 0.2 mg/ml NADPH. Reaction was initiated by adding H2O2 (0.001%), and the kinetics of NADPH oxidation was monitored for 5 min at 340 nm. GPX activity (DA340 nm/min) was calculated by subtracting the slope of the reaction from that of spontaneous oxidation in the control sample.

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Western blot analysis Total cell extracts were prepared using an M-PER Mammalian Protein Extraction Reagent Kit (Pierce) according to the manufacturer's instructions. Cytosolic extracts were prepared as described [32]. The protein concentration of each extract was determined by the Bradford assay. Cell extracts (30 μg protein/lane) were separated by electrophoresis on 10 to 15% standard SDS–polyacrylamide gels, transferred to nitrocellulose membranes, and then detected by the proper primary and secondary antibodies before visualization using a chemiluminescence kit (Pierce). The following antibodies were used: anti-Bcl-2, -Bcl-xL, -Bax, -Bak, -Bad, -XIAP, -cleaved PARP, -cytochrome c, -GPX1, and -β-actin (Cell Signaling Technology). The secondary antibodies were horseradish peroxidase-conjugated anti-rabbit and anti-mouse IgG (Santa Cruz Biotechnology). Statistical analysis Unless otherwise stated, data were expressed as means ± SD and analyzed statistically by one-way ANOVA. p b 0.05 was considered statistically significant. Results SGC-7901 cells are more susceptible to hispolon-induced cell death than are normal GES-1 cells To identify the therapeutic potential of hispolon, SGC-7901 cells and normal GES-1 cells were cultured with the indicated concentrations of hispolon for 24, 48, and 72 h, and then cell viability was determined by MTT assay. Hispolon inhibited the growth of SGC-7901 cells in a dose- and time-dependent manner (Fig. 1B, left), with 50% inhibition (IC50) at 24, 48, and 72 h of 15.6, 11.4, and 7.5 μg/ml, respectively. In contrast, only a small percentage of cell death (16.1%) was found in GES-1 cells after treatment with 20 μg/ml hispolon for 24 h (Fig. 1B, right). These results suggest that SGC-7901 cells are more susceptible to hispolon-induced cell death compared with normal GES-1 cells. To determine whether the cytotoxicity of hispolon toward SGC7901 cells was caused by apoptosis, annexin V–FITC/PI double staining and TUNEL assay were performed. SGC-7901 cells were incubated with the indicated concentrations of hispolon for 24 h, and then apoptosis was assayed using annexin V–FITC/PI double staining. Although there was little increase in apoptosis at the 5 μg/ml concentration, exposure to 10 and 15 μg/ml hispolon resulted in an increase in the percentage of early apoptotic cells (annexin V positive but PI negative) from 1.3% in control compared with 14.4 and 26.2% in treated cells. Further evidence for an increase in apoptosis is the increase in the percentage of late apoptotic cells (annexin V and PI double-positive cells), apparent at both the 10 and the 15 μg/ml concentrations (Fig. 1C). In the TUNEL assay, flow-cytometric analysis showed that hispolon treatment of SGC-7901 cells caused a dose- and time-dependent increase in the percentage of apoptotic cells (TUNEL-positive cells). Treatment of SGC-7901 cells with 10 and 15 μg/ml hispolon for 24 h induced 31.2 and 60.5% of apoptotic cells compared with 1.8% in control (Figs. 1D and E). In contrast, GES-1 cells treated with

Fig. 2. Hispolon induces caspase-dependent apoptosis in SGC-7901 cells. (A) SGC-7901 cells were treated with 0–15 μg/ml hispolon for 24 h, with or without 100 μM z-VADfmk pretreatment for 2 h, and then apoptosis was determined by TUNEL assay. Data represent means ± SD of three independent experiments (⁎p b 0.05 when cells pretreated with z-VAD-fmk and hispolon were compared with cells treated with hispolon alone). (B) SGC-7901 cells were treated with 0–15 μg/ml hispolon for 24 h, and then activation of caspase-3 was determined by flow cytometry. Data represent similar results from three independent experiments. (C) SGC-7901 cells were treated with 0– 15 μg/ml hispolon for 24 h, and then whole-cell lysates were subjected to Western blotting to assess the expression of cleavage of PARP. β-Actin was used as internal control to ensure that equal amounts of proteins were loaded in each lane. Data represent similar results from three independent experiments.

15 μg/ml hispolon for 24 h did not undergo apoptosis (Fig. 1F). These data support the conclusion that SGC-7901 cells are more susceptible to hispolon-induced cell death compared with normal GES-1 cells.

Fig. 1. Hispolon induces more cytotoxicity toward SGC-7901 cells than toward normal GES-1 cells. (A) Chemical structure of hispolon. (B) Dose- and time-dependent effects of hispolon on the cell growth inhibition of SGC-7901 cells (left) and GES-1 cells (right). Cell viability was determined by MTT assay as described in the text. Data represent means ± SD of three independent experiments (⁎p b 0.05 versus control). (C) Hispolon induced apoptosis in SGC-7901 cells. SGC-7901 cells were treated with 0–15 μg/ml hispolon for 24 h, and then apoptosis was determined by flow-cytometric analysis of annexin V–FITC/PI-stained cells. The lower right portion indicates early apoptotic cells (annexin V positive but PI negative), and the upper right portion indicates late apoptotic cells (both annexin V and PI positive). Data represent similar results from three independent experiments. (D) SGC-7901 cells were treated with 0–15 μg/ml hispolon for 24 h, and then apoptosis was determined by TUNEL assay. Data represent similar results from three independent experiments. (E) SGC7901 cells were treated with 15 μg/ml hispolon for 0–24 h, and then apoptosis was determined by TUNEL assay. Data represent means ± SD of three independent experiments (⁎p b 0.05 versus control). H15, hispolon 15 μg/ml. (F) Normal GES-1 cells were incubated with 0–15 μg/ml hispolon for 24 h, and then apoptosis was determined by TUNEL assay. Data represent means ± SD of three independent experiments.

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Hispolon induces caspase-dependent apoptosis in SGC-7901 cells To determine whether hispolon-induced apoptosis was dependent upon caspase activation, SGC-7901 cells were cultured in the presence and absence of the broad-spectrum caspase inhibitor zVAD-fmk and analyzed by TUNEL assay. Pretreatment of SGC-7901 cells with 100 μM z-VAD-fmk resulted in inhibition of hispoloninduced apoptosis (Fig. 2A), indicating that its ability to induce cell death is dependent upon caspases. Then we further checked if caspase-3, the major effector of caspase, and PARP, the main substrate of caspases, were involved in hispolon-

induced apoptosis. Activation of caspase-3 was measured using a FITCconjugated Monoclonal Active Caspase-3 Antibody Apoptosis Kit I (BD Biosciences) according to the manufacturer's protocol. Flow-cytometric analysis revealed that hispolon treatment resulted in a marked increase in caspase-3 activity after 24 h in a dose-dependent manner (Fig. 2B). Correspondingly, the amount of cleaved PARP, as demonstrated by the appearance of the 85-kDa cleavage product, increased after hispolon treatment of SGC-7901 cells in a dose-dependent manner (Fig. 2C). Caspase activity and PARP cleavage are intracellular signs of activation of the apoptotic machinery. Altogether, these data demonstrate that hispolon induces cytotoxicity of SGC-7901 cells through an apoptotic mechanism dependent upon caspase activation. Hispolon-induced apoptosis is mediated through the mitochondrial pathway To further characterize hispolon-induced apoptosis, we examined whether hispolon activates the extrinsic or intrinsic apoptotic pathway in SGC-7901 cells. To determine which apoptotic pathway hispolon activates, we examined the activity of caspase-8 and -9, the apical proteases in the extrinsic and intrinsic pathways, respectively, by colorimetric analysis. As shown in Fig. 3A, hispolon treatment increased the activity of caspase-9 and caspase-3 in a dose-dependent manner compared to vehicle-treated cells. Interestingly, caspase-8 activity in vehicle-treated and hispolon-treated cells remained unaffected. These results indicated that hispolon-induced apoptosis was most likely to occur through the mitochondrial pathway. To confirm whether mitochondrial events were involved in the induction of apoptosis, disruption of the MMP and release of mitochondrial cytochrome c in SGC-7901 cells were investigated by flow cytometry and immunoblotting. Flow-cytometric analysis revealed that exposure of SGC-7901 cells to hispolon decreased the MMP in a dose-dependent manner (Fig. 3B). The mean Rh123 fluorescence of SGC-7901 cells decreased from about 111.8 (control) to 70.5 and 52.6 by 10 and 15 μg/ml of hispolon after 24 h treatment (p b 0.05), suggesting that mitochondria were involved in hispolon-induced apoptosis. Next the release of cytochrome c was investigated by immunoblot analysis. As shown in Fig. 3C, the relative amount of cytochrome c in the cytosol of the cells treated with hispolon was dramatically increased in a dose-dependent manner compared to the vehicle-treated cells. To further examine whether important proapoptotic and antiapoptotic regulatory proteins could be modulated by hispolon in SGC-7901 cells, we determined the expression of Bcl-2 family members and inhibitors of apoptosis proteins by Western blot analysis. As shown in Fig. 3D, Bax and Bak were markedly up-regulated; Bcl-2, Bcl-xL, and XIAP were down-regulated; Bad was unchanged after hispolon treatment. Altogether, these findings demonstrate that hispolon-induced Fig. 3. Hispolon-induced apoptosis is mediated through the mitochondrial pathway. (A) Involvement of caspase activation in hispolon-induced apoptosis. After treatment with hispolon for 24 h, the cytosolic fraction of the cells was analyzed for changes in the activity of caspase-3, -8, and -9 using colorimetric assay. Data represent means ± SD of three independent experiments (⁎p b 0.05 versus control). (B) Effect of hispolon on mitochondrial membrane potential in SGC-7901 cells. After treatment with 0–15 μg/ml hispolon for 24 h, SGC-7901 cells were incubated with Rh123 (10 μg/ml) for 30 min, and then immediately subjected to flow-cytometric analysis. Cells treated with the mitochondrial uncoupler CCCP (10 μM) for 2 h were used for positive control. Results were expressed as mean Rh123 fluorescence (means ± SD of three independent experiments, ⁎p b 0.05 versus control). H5, hispolon 5 μg/ml; H10, hispolon 10 μg/ml; H15, hispolon 15 μg/ml. (C) Determination of cytochrome c release. After treatment with hispolon for 24 h, cytosolic fraction was isolated and the content of cytochrome c was examined by Western blot analysis. β-Actin was used as internal control to ensure that equal amounts of proteins were loaded in each lane. Data represent similar results from three independent experiments. (D) Effect of hispolon on expression of proapoptotic and antiapoptotic proteins. After treatment with 0–15 μg/ml hispolon for 24 h, whole-cell lysates were subjected to Western blotting to assess the expression of Bcl-2 family proteins and XIAP. β-Actin was used as internal control to ensure that equal amounts of proteins were loaded in each lane. Data represent similar results from three independent experiments.

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apoptosis in SGC-7901 cells is mediated through the mitochondrial pathway. ROS generation triggers hispolon-induced apoptosis Some previous investigations have reported that generation of ROS is associated with disruption of mitochondrial membrane potential [33,34]. Therefore, we analyzed the production of intracellular ROS in hispolon-treated and untreated cells by flow cytometry. Compared to normal GES-1 cells, SGC-7901 cells exhibited a significant increase (162.8%) in basal ROS content, as quantified by flow cytometry using DCFH-DA (Fig 4A, left). We also used HEt, a relatively specific probe for superoxide (O·-2 ) [35], to compare the basal levels of O·-2 in the two cell lines. SGC-7901 cells exhibited significantly higher HEt fluorescence (102.7) than GES-1 cells (76.6; p b 0.05), suggesting an increase in basal O·-2 in SGC-7901 cells (Fig. 4B). Treatment with hispolon in SGC-7901 cells caused a dose- and time-dependent increase in DCF-reactive ROS (Fig. 4A). The DCF fluorescence of SGC-7901 cells significantly increased as early as 1 h after 15 μg/ml hispolon treatment, indicating rapid generation of ROS production (Fig. 4A, right). In contrast, GES-1 cells were less sensitive to hispolon, showing no significant ROS elevation as the incubation continued (Fig. 4A), suggesting that the normal cells could better cope with hispolon-induced ROS accumulation, likely due to their low basal ROS output (Fig. 4A, left). Interestingly, there was no

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significant change in HEt fluorescence after treatment with hispolon in either cell line, although SGC-7901 cells consistently exhibited a higher basal O·-2 (Fig. 4B), suggesting that ROS induced by hispolon were mainly DCF-reactive species such as hydrogen peroxide (H2O2) and nitric oxide (NO), but not O·-2 . Using DAF, a relatively specific probe for NO [27], we showed that treatment of SGC-7901 cells with 15 μg/ml hispolon did not significantly increase DAF fluorescence (Fig. 4C). These data indicate that hispolon mainly induced accumulation of H2O2 in SGC-7901 cells. The above observations suggest that ROS might mediate the anticancer activity of hispolon. To further test if hispolon could preferentially induce ROS-mediated lipid peroxidation, we treated SGC-7901 and GES-1 cells with 15 μg/ml hispolon for various times and used NAO to detect oxidation of cardiolipin, a mitochondrial membrane lipid component [36]. As shown in Fig. 5A, hispolon caused a massive cardiolipin oxidation in the SGC-7901 cells, evidenced by a time-dependent decrease in NAO fluorescence. Addition of the antioxidant catalase or NAC almost completely reversed the hispolon-induced loss of NAO staining (Fig. 5B), suggesting that the decrease in NAO fluorescence was due to oxidative damage. In contrast, GES-1 cells did not show a significant loss of NAO signal (Fig. 5A). A second assay was used to confirm the preferential induction of mitochondrial membrane damage by hispolon. The functional integrity of mitochondrial membranes was assessed by flow cytometry after cells were labeled with Rh123. A decrease in this fluorescence

Fig. 4. Preferential induction of ROS accumulation by hispolon in SGC-7901 cells. (A) Effect of hispolon on cellular ROS production in SGC-7901 and GES-1 cells. After treatment with 0–15 μg/ml hispolon for 24 h (left) or 15 μg/ml hispolon for 0–24 h (right), cells were incubated with 10 μM DCFH-DA for 30 min and then immediately subjected to flow-cytometric analysis. Results were expressed as mean DCF fluorescence (means ± SD of three independent experiments, ⁎p b 0.05 versus control). H5, hispolon 5 μg/ml; H10, hispolon 10 μg/ml; H15, hispolon 15 μg/ml. (B) Effect of hispolon on cellular O·-2 content in SGC-7901 and GES-1 cells. After treatment with 0–15 μg/ml hispolon for 24 h, cells were incubated with 1 μM HEt for 30 min and then immediately subjected to flow-cytometric analysis. Results were expressed as mean HEt fluorescence (means ± SD of three independent experiments, p b 0.05 versus control). (C) Effect of hispolon on cellular NO content in SGC-7901 and GES-1 cells. After treatment with 0–15 μg/ml hispolon for 24 h, cells were incubated with 5 μM DAF-FM diacetate for 30 min and then immediately subjected to flow-cytometric analysis. Results were expressed as mean DAF fluorescence (means ± SD of three independent experiments, p b 0.05 versus control).

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Fig. 5. Selective killing of SGC-7901 cells by hispolon through ROS-mediated damage. (A) Comparison of oxidative damage to cardiolipin by hispolon in SGC-7901 and GES-1 cells detected by flow cytometry using NAO. After treatment with 15 μg/ml hispolon for 0–24 h, cells were incubated with 100 nM NAO for 15 min and then immediately subjected to flowcytometric analysis. Results were expressed as mean NAO fluorescence (means ± SD of three independent experiments, ⁎p b 0.05 versus control). (B) Effects of catalase, NAC, CSA, and TFP on hispolon-induced cardiolipin oxidation. SGC-7901 cells were pretreated with catalase (2000 U/ml), NAC (1 mM), CSA (5 μM), or TFP (5 μM) for 1 h, followed by incubation with 15 μg/ml hispolon for 24 h, and then immediately subjected to flow-cytometric analysis as above. Data represent means ± SD of three independent experiments; ⁎p b 0.05 when cells incubated with the combination of hispolon and catalase, NAC, CSA, or TFP were compared with cells incubated with hispolon alone. (C) Time-dependent effect of hispolon on the loss of MMP in SGC-7901 and GES-1 cells. After treatment with 15 μg/ml hispolon for 0–24 h, cells were incubated with 10 μg/ml Rh123 for 30 min and then immediately subjected to flow-cytometric analysis. Results were expressed as mean Rh123 fluorescence (means ± SD of three independent experiments, ⁎p b 0.05 versus control). (D–F) Effects of catalase, NAC, CSA, and TFP on the loss of MMP, caspase activation, and apoptosis induced by hispolon. SGC-7901 cells were pretreated with catalase (2000 U/ml), NAC (1 mM), CSA (5 μM), or TFP (5 μM) for 1 h, followed by incubation with 15 μg/ml hispolon for 24 h, and then immediately subjected to detection of MMP (D), caspase-3 activity (E), and apoptosis (F) as described in the text. Data represent means ± SD of three independent experiments; ⁎p b 0.05 when cells incubated with the combination of hispolon and catalase, NAC, CSA, or TFP were compared with cells incubated with hispolon alone.

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Fig. 6. Hispolon causes massive ROS accumulation in SGC-7901 cells by abrogating the GSH antioxidant system. (A) Time-dependent depletion of GSH by hispolon (15 μg/ml,1–6 h) in SGC7901 and GES-1 cells. Cellular GSH was measured by colorimetric analysis (data represent means± SD of three independent experiments; ⁎p b 0.05 versus control). (B) Hispolon-induced GSH efflux from SGC-7901 and GES-1 cells. GSH was measured by colorimetric analysis (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (C) Effects of catalase (CAT) and NAC on hispolon-induced ROS increase. SGC-7901 cells were preincubated with 2000 U/ml CAT or 1 mM NAC for 1 h followed by 15 μg/ml hispolon for 1–6 h. ROS were measured by flow cytometry using DCFH-DA (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (D) Effects of catalase and NAC on hispolon-induced GSH depletion. SGC-7901 cells were preincubated with 2000 U/ml CAT or 1 mM NAC for 1 h followed by 15 μg/ml hispolon for 1–6 h. GSH levels were determined as above (data represent means± SD of three independent experiments; ⁎p b 0.05 versus control). (E) Differential cytotoxic effects of BSO in SGC-7901 and GES-1 cells. SGC-7901 and GES-1 cells were incubated with indicated concentrations of BSO for 24 h. Cell viability was determined by MTT assay. Data represent means± SD of three independent experiments. ⁎Statistically significant difference between the two cell lines (p b 0.05). (F) Effect of hispolon on cellular GPX activity. SGC-7901 cells were incubated with 15 μg/ml hispolon for 0–24 h, and cellular protein extracts were assayed for GPX activity as described in the text (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (G) Effect of hispolon on GPX1 protein expression in SGC-7901 cells after treatment with 15 μg/ml hispolon for 0–24 h. GPX expression was determined by Western blot analysis. β-Actin was used as internal control to ensure that equal amounts of proteins were loaded in each lane. Data represent similar results from three independent experiments.

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Fig. 7. Hispolon effectively induces cytotoxicity and ROS increase in other human gastric cancer cells. (A) Hispolon increases caspase-3 activity in MGC-803 and MKN-45 cells. Cells were treated with 15 μg/ml hispolon for 24 h, with or without preincubation with 1 mM NAC (1 h). Caspase-3 activity was measured by colorimetric assay (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (B) Hispolon induces apoptosis in MGC-803 and MKN-45 cells. Cells were treated with 15 μg/ml hispolon for 24 h, with or without preincubation with 1 mM NAC (1 h). Apoptosis was measured by TUNEL assay (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (C) Hispolon induces cellular GSH depletion in MGC-803 and MKN-45 cells. Cells were treated with 15 μg/ml hispolon for 1–6 h, with or without preincubation with 1 mM NAC (1 h). GSH content was measured by colorimetric analysis (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control). (D) Hispolon induces ROS increase in MGC803 and MKN-45 cells. Cells were treated with 15 μg/ml hispolon for 1–6 h, with or without preincubation with 1 mM NAC (1 h). ROS was measured by flow-cytometric analysis using DCFH-DA (data represent means ± SD of three independent experiments; ⁎p b 0.05 versus control).

indicates a loss of MMP [37]. As shown in Fig. 5C, the loss of MMP was time dependent and occurred significantly as early as 2 h after hispolon treatment. In contrast, GES-1 cells did not exhibited a significant loss of MMP after hispolon treatment. Interestingly, when the time courses for cardiolipin oxidation (Fig. 5A) and loss of MMP (Fig. 5C) in SGC-7901 cells were compared, it appeared that membrane oxidation occurred first, followed by the loss of membrane integrity with a delay of approximately 1 h. To verify the cause–effect relationship between ROS increase and cell death, we tested the effect of the antioxidants catalase (H2O2scavenging enzyme) and NAC (a general free radical scavenger and GSH precursor) on hispolon-induced cell death in SGC-7901 cells. As shown in Fig. 5, catalase and NAC significantly blocked the loss of MMP (Fig. 5D), caspase activation (Fig. 5E), and apoptosis (Fig. 5F) induced by hispolon. In contrast, CSA and TFP, inhibitors of the mitochondrial permeability transition, had little effect on the cardiolipin oxidation (Fig. 5B), loss of MMP (Fig. 5D), caspase activation (Fig. 5E), or apoptosis (Fig. 5F) induced by hispolon. These results suggested that ROS were critical in mediating hispoloninduced cytotoxicity in SGC-7901 cells.

Hispolon causes massive ROS accumulation in SGC-7901 cells by abrogating the GSH antioxidant system As hispolon exerted cytotoxicity by causing excessive ROS accumulation, we then investigated the mechanisms by which hispolon caused ROS increase. Based on the observation that NAC can effectively block hispolon-induced cell death, we postulated that the active ROS generation in SGC-7901 cells would render them highly dependent on GSH to maintain redox balance and that a depletion of GSH by hispolon would result in an excessive accumulation of ROS to a threshold that triggers cell death. To test this possibility, we first examined the effect of hispolon on GSH contents. As shown in Fig. 6A, incubation of SGC-7901 cells with 15 μg/ml hispolon led to a depletion of cellular GSH by 80% in 1 h and almost complete depletion in 2 h. In GES-1 cells, approximately 91 and 83% GSH remained at 1 and 2 h after hispolon treatment, respectively. Analysis of GSH in the medium showed that GSH was rapidly exported from the cells (Fig. 6B). The amount of GSH in the medium was quantitatively accountable for the GSH loss in the cells, suggesting that hispolon-induced GSH efflux was a key mechanism for GSH depletion. Interestingly, catalase effectively

Fig. 8. ROS mediates hispolon potentiating the toxicity of MMC, 5-FU, and DOX, but not CIS. After combined treatment with (A and B) hispolon (2.5 μg/ml) and MMC (0.5–5 μg/ml), (C and D) hispolon (2.5 μg/ml) and 5-FU (1–10 μg/ml), (E and F) hispolon (2.5 μg/ml) and DOX (0.1–0.5 μg/ml), or (G and H) hispolon (2.5 μg/ml) and CIS (1–10 μg/ml) for 24 h, with or without preincubation with 1 mM NAC (1 h), cell viability, ROS production, and cellular GSH content were measured as described in the text. Data represent means ± SD of three independent experiments. ⁎p b 0.05 when cells incubated with the combination of hispolon and MMC, 5-FU, DOX, or CIS were compared with cells incubated with MMC, 5-FU, DOX, or CIS alone.

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abrogated hispolon-induced ROS increase (Fig. 6C) but did not prevent GSH depletion (Fig. 6D), indicating that the depletion of GSH was not secondary to the ROS increase. In contrast, pretreatment of SGC-7901 cells with 1 mM NAC effectively prevented hispolon-induced GSH depletion (Fig. 6D) and ROS accumulation (Fig. 6C). Combined with the data that NAC suppressed the cytotoxicity of hispolon (Fig. 5F) these results revealed that depletion of cellular GSH is an important mechanism responsible for hispolon-induced ROS accumulation and cell death. To further demonstrate the important role of GSH in the survival of SGC-7901 cells, we used BSO, an inhibitor of GSH synthesis, to determine if this agent could also kill SGC-7901 cells. As shown in Fig. 6E, BSO exhibited greater cytotoxic effect on SGC-7901 cells compared to GES-1 cells, although a longer incubation and higher concentrations were required to achieve significant cell death. Because GPX is the major enzyme that uses GSH as the substrate to scavenge peroxides, we examined if hispolon could affect GPX activity and impair the cell's ability to use the remaining GSH. As shown in Figs. 6F and G, the reduction of GPX enzyme activity was time dependent and occurred significantly as early as 1 h after hispolon treatment (Fig. 6F), whereas the expression of GPX protein did not decrease even after 24 h incubation of hispolon (Fig. 6G). These results indicated that hispolon not only depleted the cellular GSH pool but also inhibited GPX enzyme to abrogate the GSH antioxidant system, leading to massive ROS accumulation in SGC7901 cells. Hispolon is effective in killing other gastric cancer cells The ability of hispolon to induce severe ROS-mediated damage in the SGC-7901 cells prompted us to test its effectiveness in killing other gastric cancer cells. As shown in Figs. 7A and B, hispolon was very effective in inducing apoptosis of both MGC-803 and MKN-45 gastric cancer cells as assayed by caspase-3 (Fig. 7A) and TUNEL assays (Fig. 7B). Flow-cytometric analysis showed that hispolon (15 μg/ml) caused 36.2 and 48.8% cell death in MGC-803 and MKN-45 cells, respectively, after 24 h treatment, which was largely abrogated by 1 mM NAC (Fig. 7B). Consistently, hispolon caused a depletion of cellular GSH and a substantial ROS increase, which were also effectively prevented by 1 mM NAC (Figs. 7C and D). Low-dose hispolon augments the cytotoxicity of MMC, 5-FU, DOX, and CIS Because some previous investigations have reported that a combination of different agents proved to be more effective than a single agent in killing cancer cells [38,39], we considered the possibility that low doses of hispolon might potentiate the effects of chemotherapeutic agents used in the clinical management of gastric cancer. SGC-7901 cells were incubated with MMC (0.5–5 μg/ml), 5-FU (1–10 μg/ml), DOX (0.1–0.5 μg/ml), or CIS (1–10 μg/ml), in the presence or absence of a low dose of hispolon (2.5 μg/ml). Then cell viability, ROS production, and cellular GSH content were determined. As shown in Fig. 8, the combination of hispolon and MMC (Fig. 8A), hispolon and 5-FU (Fig. 8C), or hispolon and DOX (Fig. 8E) caused a progressive increase in cytotoxicity over that seen with MMC, 5-FU, or DOX alone. These cytotoxic effects were largely abrogated by NAC. Consistently, these combination treatments caused a substantial ROS increase and GSH depletion, which were also effectively prevented by NAC (Figs. 8B, D, and F). These observations suggest that ROS might mediate hispolon potentiating the toxicity of MMC, 5-FU, and DOX. In contrast, the potentiation of CIS toxicity by hispolon (Fig. 8G) seemed to be much less than that of the other three drugs (MMC, 5-FU, and DOX), and there was no significant ROS increase or GSH depletion in SGC7901 cells after the combination of hispolon and CIS treatment (Fig. 8H). Moreover, NAC cannot prevent the cytotoxicity caused by the combination of hispolon and CIS (Fig. 8G). These results indicate that

the potentiation of CIS toxicity by hispolon was unlikely to be triggered by ROS. Discussion Various chemotherapy drugs, including doxorubicin, 5-fluorouracil, cisplatin, and mitomycin C, have been used to treat gastric cancer [5]. Unfortunately, all of these anticancer drugs affect not only pathological tumor cells, but also normal cells [7]. Therefore, the search for new chemopreventive and antitumor agents that are more effective but less toxic has become a matter of great interest. In this study, we found that hispolon, a polyphenol compound, acts directly on human gastric cancer cells to induce cytotoxicity in a manner that causes apoptosis. Hispolon induced cytotoxicity toward gastric cancer cells at concentrations that are minimally toxic to normal gastric cells. Investigating the mechanism by which gastric cancer cells undergo apoptosis in response to hispolon treatment, we found that hispolon abolished the glutathione antioxidant system through depletion of cellular GSH, and inhibition of GPX enzyme activity resulted in severe ROS accumulation. Excessive ROS caused oxidative damage to the mitochondrial membranes and impaired the membrane integrity, leading to cytochrome c release, caspase activation, and apoptosis. The correlation between ROS increase and apoptosis induced by hispolon in gastric cancer cells and the suppression of cytotoxicity by NAC or catalase suggest the critical role of ROS in hispolon-induced cell death. Moreover, the observations that BSO caused much greater cytotoxicity in gastric cancer cells compared to normal gastric cells further support the role of GSH in maintaining redox balance and gastric cancer cells survival. Reactive oxygen species, such as H2O2 and O·-2 , are constantly produced during metabolic processes in all living species. Under physiological conditions, the maintenance of an appropriate level of intracellular ROS is important in keeping redox balance and cell proliferation [40–43]. However, excessive ROS accumulation will lead to cellular injury, including lipid peroxidation, protein oxidation, enzyme inactivation, and oxidative DNA damage [44–47]. One common feature of cancer cells is the increase in ROS generation. Compelling evidence suggests that most cancer cells are under oxidative stress associated with increased metabolic activity and production of ROS [48]. These biochemical characteristics make the cancer cells more vulnerable to damage by additional ROS stress, either through inhibiting ROS elimination or by adding exogenous ROS [49,50]. The cell-damaging property of ROS and the increased ROS generation in cancer cells may provide an opportunity to exploit the cell killing potential of ROS by using exogenous ROS-stressing agents to increase the intracellular ROS to a toxic level, or the threshold that triggers cell death. In the present study, we found that hispolon effectively induced apoptosis in human gastric cancer cells, which was associated with a significant increase in the levels of intracellular ROS, whereas there is no obvious cell death or ROS increase in human normal gastric cells after hispolon treatment. The degrees of ROS accumulation and cell death induced by hispolon were dependent on endogenous ROS generation. The heightened basal level of ROS in gastric cancer cells seems responsible for their high sensitivity to hispolon. Because gastric cancer cells depend on GSH to impair the active ROS output, abolishment of the GSH antioxidant system by hispolon would severely affect these cells, leading to oxidative damage and cell death. In contrast, it is less likely to induce such severe ROS stress in normal gastric cells, due to their low basal ROS output. Our study indicated that hispolon induced the ROS increase by two mechanisms, depleting GSH by promoting its efflux and inhibiting GPX enzyme activity, which together effectively abrogate the GSH antioxidant system. Although it has been widely recognized that cancer cells normally produce more ROS than do normal cells and are intrinsically under oxidative stress [48,51–53], the mechanism responsible for the increase in ROS generation in cancer cells is largely unclear. Oncogenic signals, mitochondrial dysfunction, and

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active metabolism are likely factors contributing to the increased production of ROS in cancer cells [26,28,54–56]. An important observation of our study is that hispolon dramatically enhanced the cytotoxic effects of other chemotherapeutic agents (such as MMC, 5-FU, and DOX) commonly used in the treatment of gastric cancer, whereas the potentiation of CIS toxicity by hispolon seemed to be much less than that of the other three drugs. A possible explanation for this difference might be that the combination of hispolon and MMC, 5-FU, or DOX synergistically caused severe ROS accumulation in SGC-7901 cells, leading to massive cell death. A similar phenomenon was found in a recent study: a combination of two agents synergistically induced massive ROS increase and cell death in cancer cells [26]. In contrast, the combination of hispolon and CIS did not cause significant ROS increase in SGC-7901 cells, resulting in less cell death. This is in agreement with a recent study that the cytotoxic effect of CIS in some cancer cells was unlikely to be triggered by ROS [28]. Taken together, these findings may provide a possibility that the combination of hispolon and other ROS-generating agents would be a new strategy to treat gastric cancer. In addition to the preferential cytotoxicity in gastric cancer cells, the safety of hispolon is indirectly reflected by the facts that PL, a traditional medicinal mushroom from which hispolon can be extracted [18], has been widely used as a medication with multiple functions for hundreds of years in China and that an adverse effect of PL associated with the treatment was not reported. Furthermore, a recent investigation reported a case of advanced prostate cancer that markedly improved after administration of the PL extract [57]. To the best of our knowledge, the extract of PL, which can be obtained from some dietary supplement companies in China, is being consumed widely in China, especially among people with malignant disease. A growing number of people take at least 30 g PL extract a day for maintenance of their health, which is equivalent to ~30 mg of pure hispolon. The stomach surface area of an average human adult is ~900 cm2, equivalent to the total culture area of 15 petri dishes (100 mm) [58]. If we apply 30 mg of hispolon to 15 dishes of gastric cancer cells, it would kill them definitely. However, it should be noted that human body is a much more complex system, and an in vitro dosage is not directly comparable to the amount of hispolon uptake by PL consumers. In conclusion, our study suggests that hispolon, used either alone or in combination with other drugs, may represent a promising novel targeted approach in the treatment of gastric cancer. Further investigation of hispolon in mouse models will contribute to the additional understanding of its in vivo activity toward malignant cells and its potential toxicity toward normal tissues. Acknowledgments This work was supported by grants from the Natural Science Fund of Zhejiang Province (No. R207609), the National High Technology Research and Development Program of China (No. 2007AA021506), and the Research Project of Science and Technology of Zhejiang Province, China (No. 2005C23027). References [1] Parkin, D. M. Global cancer statistics in the year 2000. Lancet Oncol. 2:533–543; 2001. [2] Parkin, D. M.; Bray, F.; Ferlay, J.; Pisani, P. Global cancer statistics, 2002. CA Cancer J. Clin. 55:74–108; 2005. [3] De Vita, F.; Giuliani, F.; Galizia, G.; Belli, C.; Aurilio, G.; Santabarbara, G.; Ciardiello, F.; Catalano, G.; Orditura, M. Neo-adjuvant and adjuvant chemotherapy of gastric cancer. Ann. Oncol. 18:vi120–vi123; 2007. [4] Ajani, J. A. Evolving chemotherapy for advanced gastric cancer. Oncologist 10:49–58; 2005. [5] Wohrer, S. S.; Raderer, M.; Hejna, M. Palliative chemotherapy for advanced gastric cancer. Ann. Oncol. 15:1585–1595; 2004. [6] Ridwelski, K.; Gebauer, T.; Fahlke, J.; Kroning, H.; Kettner, E.; Meyer, F.; Eichelmann, K.; Lippert, H. Combination chemotherapy with docetaxel and cisplatin for locally advanced and metastatic gastric cancer. Ann. Oncol. 12:47–51; 2001.

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