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Experimental Therapeutics, Molecular Targets, and Chemical Biology |
1 Laboratoire d'Immunologie and 2 Laboratoire de Recherche Chirurgicale, Unité Propre de Recherche de l'Enseignement Supérieur 1833, Faculté Cochin, Université Paris V; 3 Service d'Oncologie Médicale, Hôpital Cochin; 4 Service de Chirurgie Digestive, Centre Hospitalier Universitaire Lariboisière, Université Paris VII, Paris, France; and 5 Service de Chirurgie Digestive, Centre Hospitalier Universitaire Henri-Mondor, Université Paris XII, Créteil, France
Requests for reprints: Frédéric Batteux, Laboratoire d'Immunologie, Pavillon Hardy, Hôpital Cochin, 75679 Paris cedex 14, France. Phone: 33-1-58-41-20-07; Fax: 33-1-58-41-20-08; E-mail: frederic.batteux{at}cch.ap-hop-paris.fr.
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Abstract |
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Key Words: reactive oxygen species • cancer • apoptosis • antioxidant molecule • chemotherapy
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B and activator protein 1, during tumor progression (8). Finally, the transforming capacity of ROS is illustrated by the oncogenic transformation of NIH 3T3 cells by the NADPH oxidase homologue MOX-1, which induces the production of superoxide anions (O2°-; refs. 9, 10). Whereas, under certain circumstances, ROS promote cell proliferation, they can also induce apoptosis. Indeed, most anticancer drugs kill their target cells at least in part through the generation of elevated amounts of intracellular ROS. ROS can stimulate proapoptotic signal molecules, such as apoptosis signal regulating kinase 1, c-Jun-NH2-kinase, and p38 (11, 12); activate the p53 protein pathway; or engage the mitochondrial apoptotic cascade (13). The various ROS can exert different effects according to their nature and to their intracellular level, which is determined by both their production rate and the activity of antioxidant enzymes. Using pharmacologic modulators of the ROS pathways, we first investigated the source and nature of ROS produced in nontransformed cells and in established tumor cell lines and then determined their respective contribution to various cellular signals. Finally, we investigated the consequences of ROS modulation in association with cytolytic drugs in vitro and in vivo in mice with implanted tumors.
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Materials and Methods |
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Chemicals and Cell Lines. All chemicals were from Sigma (Saint Quentin Fallavier, France) except for mangafodipir (Teslascan, Amersham Health, Amersham, United Kingdom), oxaliplatin (Eloxatine, Sanofi-Pharma, Paris, France), and paclitaxel (Taxol, Bristol-Myers Squibb, Rueil Malmaison, France). CT26 (mouse colon carcinoma), Hepa 1-6 (mouse liver hepatoma), and NIH 3T3 (mouse fibroblast) were obtained from American Type Culture Collection (Manassas, VA). Cells were cultured in DMEM/Glutamax-I supplemented with 10% heat-inactivated FCS and antibiotics (Life Technologies, Cergy Pontoise, France). All cell lines were routinely tested to rule out Mycoplasma infection of cells.
Cellular Production of O2°- and H2O2. Cells (2 x 104 per well) were seeded in 96-well plates (Costar, Corning, Inc., Corning, NY) and incubated for 48 hours with various concentrations of copper(II)(3,5-diisopropylsalicylate)2 (CuDIPS), manganese(III)tetrakis(4-benzoic acid) porphyrin (MnTBAP), N-acetyl-L-cysteine (NAC), reduced glutathione (GSH), aminotriazol (ATZ), buthionine sulfoximine (BSO), catalase, or culture medium alone. Cellular production of O2°- was evaluated with the use of nitroblue tetrazolium reduction technique (14). The level of intracellular H2O2 was assessed spectrofluorimetrically (Victor2, Perkin-Elmer, Paris, France) by oxidation of 2',7'-dichlorodihydrofluorescein diacetate (Molecular Probes, Leiden, the Netherlands). O2°- and H2O2 productions were reported to the amount of proteins in each sample (bovine serum albumin microbiuret assay, Pierce, Bezons, France).
Determination of Enzymatic Activities. The SOD activities of tumor or normal cells and tissues were evaluated with the use of nitroblue tetrazolium reduction technique as previously described by Beauchamp and Fridovich (14). The catalase activities of tumor or normal cells and tissues were determined at 25°C by UV spectroscopy at 240 nm according to Aebi (15). Levels of GSH were measured by the method of Baker et al. (16). Antioxidant enzyme activities and GSH content of cells and tissues were reported to the amount of proteins in each sample.
In vitro Cell Proliferation and Viability Assays. Cells (2 x 104 per well) were seeded in 96-well plates (Costar) and incubated for 48 hours with various amounts of pharmacologic modulators of antioxidant enzymes as indicated in the figure captions. Cell proliferation was determined by pulsing the cells with [3H]thymidine (1 µCi/well) during the last 16 hours of culture. Cell viability was evaluated by the reduction of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide to formazan. The absorbance at 550 nm was recorded in each well with the use of an ELISA microplate reader. Results are expressed as % viable cells ± SE versus cells in culture medium alone (100% viability). Cell viability was controlled in all cases by the crystal violet assay.
In vitro Cytostatic and Cytotoxic Effects of Exogenous H2O2. Cells (5 x 104 cells/well) were seeded as above in culture medium supplemented or not with 400 µmol/L NAC, or 400 µmol/L ATZ (a catalase inhibitor) or 400 µmol/L BSO, which depletes GSH. H2O2 oxidative stress was elicited by the addition of serial dilutions of H2O2 (Sigma) to the cells for 48 hours. The cytostatic and the cytotoxic effects of H2O2 were evaluated by thymidine incorporation and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide reduction.
Measurement of O2°- Production by Flow Cytometry. Cells were treated with either 10 µmol/L rotenone (an inhibitor of mitochondrial complex I), 10 µmol/L antimycin (an inhibitor of mitochondrial complex III), 10 µmol/L diphenyleneiodonium (an inhibitor of NADPH oxidase), or 10 µmol/L allopurinol (an inhibitor of xanthine oxidase) for 30 minutes in culture medium. Cells were washed in HBSS and resuspended in HBSS containing 10 µmol/L DHE (Molecular Probes) at 37°C for 30 minutes. After two washes, cells were resuspended in 500 µL HBSS. Cells (20,000 events per sample) were analyzed by flow cytometry (FACScalibur, Becton Dickinson, Mountain View, CA).
Immunoblotting of Cell Lysates. Cells were treated or not with 400 µmol/L NAC for 24 hours then lysed in ice-cold 10 mmol/L Tris buffer, pH 7.5, with protease inhibitors, 25 mmol/L NaF, 0.5 mmol/L sodium orthovanadate, and 1% Triton X-100. Forty micrograms of cell lysate were analyzed by immunoblotting after 10% SDS-PAGE with the use of anti–extracellular signal-regulated kinase 2 monoclonal antibody from Santa Cruz (Le Perray en Yvelines, France), and anti p-ERK monoclonal antibody from New England Biolabs (Saint Quentin en Yvelines, France).
DNA Strand Break Analysis. The pcDNA3.1 plasmid DNA (Invitrogen) was incubated with oxaliplatin at a molar ratio of 0.50 in a final volume of 50 µL. MnTBAP (5 µmol/L), CuDIPS (5 µmol/L), MnDPDP (5 µmol/L), or NAC (5 mmol/L) was added into the mixture. O2°- was generated with the use of 200 µmol/L xanthine (Sigma) and 1 unit of xanthine oxidase (Sigma) in the dark at 37°C for 24 hours. Then, 10 µL aliquots were electrophoresed on a 0.8% agarose gel and revealed by ethidium bromide staining. The gels were analyzed with the use of a scanner densitometer (Vilber Lourmat, Marnes-la-Vallée, France).
In vivo Antitumor Activity of Antioxidant Treatments. CT-26 or Hepa 1-6 tumor cells (2 x 106) were injected s.c. into the back of the neck of BALB/c or C57/BL6 mice, respectively. When the tumors reached a mean size of 200 to 500 mm3, the animals received a single injection of either 20 mg/kg oxaliplatin (Eloxatine) or of vehicle alone. Mice were then injected or not with 10 mg/kg MnDPDP, 10 mg/kg MnTBAP, or 10 mg/kg CuDIPS or with 150 mg/kg NAC i.p. (three injections weekly at the same dosages for 1 month). Tumor size was measured with a vernier caliper every 3 days. Tumor volume was calculated as follows: TV (mm3) = (L x W2)/2, where L is the longest and W the shortest radius of the tumor in millimeters. Results are expressed as means of tumor volumes ± SE (n = 15 in each group). In five mice implanted with CT26 cells, biopsies have been done every 15 days for 45 days. Tumor cells were then compared in vitro with cells from the original CT26 line in terms of ROS production, proliferation rate, intracellular GSH content, and sensitivity to oxaliplatin.
Statistical Analysis. The statistical significance of differences between experimental treated groups and untreated controls was analyzed by 
2 test for incidence data and by Student's t test for comparison of means. A level of P < 0.05 was accepted as significant. * or 
: P < 0.05; ** or : P < 0.02; *** or : P < 0.01; **** or : P < 0.001 versus controls.
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H2O2 production decreased in a dose-dependent manner in all cell types incubated with the thiol-related compound NAC that displays a catalase-like and a glutathione reductase–like activity (Fig. 1B; ref. 19). The concentration of H2O2 decreased in NIH 3T3 cells treated by GSH and increased in NIH 3T3 cells treated with BSO, an agent that depletes the intracellular pool of GSH (Fig. 1B). By contrast, inhibition of catalase by ATZ increased H2O2 production in both tumor cell lines. ATZ had a milder effect (50% increase) in NIH 3T3 cells. Adding exogenous catalase decreased H2O2 level by 37% in CT26 and 33% in Hepa 1-6 tumor cells but had no effect in nontransformed cells (Fig. 1B).
Origin of ROS in Normal and Tumor Cells. Mitochondria are the main source of ROS in nonphagocytic cells (20) but other cytosolic enzymatic systems, such as NADPH oxidase (9) or xanthine oxidase (21), can generate O2°-. Rotenone and antimycin increased the production of O2°- in CT26 or Hepa 1-6 tumor cells but not in NIH 3T3 cells (Fig. 2). By contrast, neither the NADPH-dependent oxidase inhibitor diphenyleneiodonium nor the xanthine oxidase inhibitor allopurinol could inhibit the production of O2°- by tumor cells, whereas diphenyleneiodonium inhibited ROS production in NIH 3T3 fibroblasts (Fig. 2).
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The Effects of Anticancer Drugs Are Modulated by H2O2. In tumor cells cultured with various concentrations of the platinum antitumoral compound oxaliplatin (22), a dose-dependent increase in ROS production associated with a decrease in proliferation was observed upon addition of SOD mimics and BSO. Opposite effects were induced by NAC and GSH (Fig. 4). The catalase pathway was not involved in the antitumoral activity of oxaliplatin. The inhibition of the cytostatic and cytotoxic activities of oxaliplatin by NAC and their increase by SOD mimics was further investigated with the use of other anticancer molecules known to augment intracellular levels of H2O2 in tumor cells, such as paclitaxel (23) and 5-fluorouracil (5-FU; ref. 24). Incubating tumor cells with both drugs in association with increasing concentrations of NAC resulted in a dose-dependent decrease in the cytostatic and cytotoxic effects of paclitaxel or 5-FU on tumor cells versus incubation with paclitaxel or 5-FU alone (Fig. 5A). Reciprocal effects of SOD mimics MnDPDP, MnTBAP, and CuDIPS were observed.
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ROS Modulate Antitumoral Activity of Cytotoxic Drugs In vivo. When MnTBAP, CuDIPS, or MnDPDP (another SOD mimic; ref. 26) was repeatedly injected into BALB/c mice with CT26 tumors, the volumes of tumors were respectively 59% (P < 0.01), 28% (not significant), and 54% (P < 0.01) smaller than in untreated controls after 1 month (Fig. 6A). In animals treated by the association of oxliplatin and either MnTBAP, CuDIPS, or MnDPDP, the volumes of tumors were respectively 35%, 31%, and 63% smaller after 1 month than with oxaliplatin alone (P < 0.001). Injecting NAC alone resulted in 44% increase in tumor volumes after 1 month compared with untreated mice (P < 0.01). NAC infusion into oxaliplatin-treated mice completely abrogated the beneficial effect of oxaliplatin because, at 1 month, the volumes of tumors were similar to those in the absence of oxaliplatin. The same effects were observed in immunocompetent C57/BL6 mice injected with Hepa 1-6 cells (Fig. 6B). In implanted tumors iteratively biopsied for 45 days, time-dependent increases in GSH content (P < 0.01), O2°- and H2O2 production (P < 0.001 in both cases), and a time-dependent decrease in proliferation rate (P < 0.001) were observed at day 45 versus original cell line.
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Discussion |
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In nontransformed cells, we observed that ROS essentially originate from the cytosolic NADPH and are controlled by the GSH system. Our data are in agreement with the previous observation that Mox-1, a homologue of the gp91 phox, the catalytic moiety of the NADPH oxidase, increases O2°- and H2O2 generation in NIH 3T3 cells, the latter species being responsible for an increased mitotic rate, cell transformation, and tumorigenicity (9, 10). Similarly, as previously described (28), treating nontransformed fibroblasts or human umbilical vein endothelial cells with low amounts of exogenous H2O2 increased the proliferative rate of NIH 3T3 cells. The intracellular targets of H2O2 involved in cell growth are multiple. They include p42/p44 and p38 mitogen-activated protein kinase, p70S6K, signal transducers and activators of transcription, Atk/protein kinase B and phospholipase D signaling pathways, as well as direct inhibition of protein tyrosine phosphatase-1B (1, 8). The effects of ROS on the growth of nontransformed NIH 3T3 cells are finely regulated because detoxification of intracellular H2O2 by NAC decreased the proliferative rate. This already observed phenomenon is related to the suppression of cell cycle progression into G1 phase by inhibition of the mitogen-activated protein kinase pathway (29).
The situation is quite different in established tumor cell lines, such as CT26 and Hepa 1-6, in which the increased generation of ROS resulted from both an elevated mitochondrial production and a profound decrease in the activity of antioxidant enzymes. No increase in the proliferative rates was observed whatever the amounts of exogenous H2O2 added to the cultures. On the contrary, adding increasing amounts of exogenous H2O2 or increasing its intracellular levels with the use of SOD mimics led to a dose-dependent decrease in proliferation and to cell death. Similar data have been obtained in A549 human lung carcinoma cell line and in cell lines with various p53 gene mutations, suggesting that ROS modulation induced by SOD mimics or NAC are not linked with a particular p53 status. Conversely, detoxification of H2O2, especially through the catalase pathway, stimulated tumor cell proliferation. Those data are in line with previous observations that MnSOD or Cu/ZnSOD gene transfection inhibits growth of glioma (30) and pancreatic tumor cell lines (31), a phenomenon that can be reverted by cotransfection of the catalase gene (32, 33). Our data are also in agreement with the report by Hussain et al. (34), showing that p53 is associated with up-regulation of MnSOD resulting in ROS overproduction and apoptosis, a phenomenon inhibited by overexpression of catalase.
Although H2O2 directly controls tumor cell proliferation, OH°, which results from the conversion of H2O2 via the Fenton reaction, is also involved in that control. Indeed, addition of mannitol, a compound known to detoxify OH° (35), stimulated the growth of tumor cell lines. Along with catalase, the glutathione pathway could also be involved in the regulation of the intracellular concentration of H2O2 and tumor cell growth. Indeed, adding seleno-L-methionine, a molecule that stimulates glutathione peroxidase activity, augmented the proliferative rate of the colon carcinoma cell lines as previously described (36). However, adding reduced GSH or depleting intracellular GSH with BSO had weaker effects on the proliferation of tumor cells than exogenous catalase, probably because of the indirect effects of those molecules on the intracellular concentration of H2O2.
Cytotoxic drugs induce cellular stress responses and the generation of ROS. However, whether this phenomenon participates in the antitumoral activity of those drugs has remained unclear thus far. To address this issue, we have examined the combined effects of cytotoxic molecules with several modulators of ROS pathways. As already observed with cisplatin (37), oxaliplatin increased ROS production and decreased CT26 and Hepa 1-6 cell proliferation in a dose-dependent manner. Whereas the GSH system was not involved in the control of the basal proliferative rate of tumor cells, it modulated the ROS production and the antitumoral activity of oxaliplatin as shown by the decrease in ROS levels and the increase in tumor cell proliferation following addition of exogenous GSH. Several lines of evidence suggest that the antagonistic effects of GSH and oxaliplatin involve intracellular mechanisms and do not result from extracellular interaction of GSH with oxaliplatin. Indeed, in our study, tumor cells depleted of GSH by BSO had an increased sensitivity to oxaliplatin. Moreover, when exogenous GSH was added with oxaliplatin to tumor cells in culture, the initial increase in ROS levels was similar to that observed when cells were incubated with oxaliplatin alone. Those data confirm that platinum-induced oxidative stress involves oxidation of cellular components and depletion of intracellular GSH (37, 38). The inhibition of the cytostatic and cytotoxic activities of oxaliplatin by NAC and their increase by SOD mimics was further investigated with the use of other anticancer molecules known to augment intracellular levels of H2O2 in tumor cells. Incubating CT26 and Hepa 1-6 cells with the taxane-related compound paclitaxel (23) or with 5-FU (24), with increasing concentrations of NAC, resulted in a dose-dependent decrease in the cytostatic and cytotoxic effects of paclitaxel and 5-FU on tumor cells versus incubation with individual drugs alone. Reciprocal effects were observed with SOD mimics MnTBAP and CuDIPS that generate H2O2 through O2°- dismutation.
DNA is the main target of platinum-related antitumor drugs. H2O2 generated by those drugs in association with that produced by metalloporphyrin SOD mimics can alter DNA structure (25). Indeed, the damages caused to DNA by oxaliplatin in vitro were increased by the addition of SOD mimics, whereas the association of NAC and oxaliplatin was antagonistic. The same was observed in vivo in murine models of colon cancers. During tumor growth, H2O2 production increased whereas ex vivo proliferation rates decreased, and the GSH content augmented whereas an ex vivo resistance to oxaliplatin developed. Consequently, the three SOD mimics studied decreased, whereas NAC increased tumor growth when given alone in mice with implanted tumors. In addition, when oxaliplatin was associated, the three SOD mimics potentiated, whereas NAC abrogated the antitumoral effect of oxaliplatin. This observation should stimulate further clinical investigations because NAC is widely used after surgery as a mucolytic agent in patients operated for cancers. Our observation is in line with a previous large human study that showed an increased incidence in lung cancer in heavy smokers treated with antioxidant molecules (39). In conclusion, the growth of normal cells is triggered when those cells are submitted to oxidant signals directed to growth-related genes up to a critical threshold beyond which ROS become cytotoxic. Because in tumor cells, the level of endogenous ROS is close to that threshold, the simultaneous exposure to ROS-generating agents and to cytotoxic drugs dramatically increases the rate of cell death. Those data have clinical implications in humans. Indeed, antioxidant molecules probably play a protective role against cancer in healthy individuals by preventing DNA damages linked to the oxidative stress. However, once cancer cells have emerged, a cancer-promoting effect can result from the administration of agents that decrease intracellular H2O2 levels, whereas an anticancer activity is exerted by agents that tend to increase that level.
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Acknowledgments |
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The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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Footnotes |
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Received 5/21/04. Revised 11/ 2/04. Accepted 11/29/04.
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is mediated by ceramide. Am J Respir Cell Mol Biol 2001;24:762–8.
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