Mechanism of colon cancer cell apoptosis mediated by ... - Nature

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Oncogene (2001) 20, 4070 ± 4084 2001 Nature Publishing Group All rights reserved 0950 ± 9232/01 $15.00 www.nature.com/onc

Mechanism of colon cancer cell apoptosis mediated by pyropheophorbide-a methylester photosensitization Jean-Yves Matroule1, Chris M Carthy3, David J Granville3, Olivier Jolois2, David WC Hunt3 and Jacques Piette*,1 1

Laboratory of Virology and Immunology, University of Liege, B-4000 Liege, Belgium; 2Laboratory of Human Histology, University of Liege, B-4000 Liege, Belgium; 3QLT Inc., 887 Great Northern Way, Vancouver, BC, Canada, V5T 4T5

Pyropheophorbide-a methylester (PPME) is a second generation of photosensitizers used in photodynamic therapy (PDT). We demonstrated that PPME photosensitization triggered apoptosis of colon cancer cells as measured by using several classical parameters such as DNA laddering, PARP cleavage, caspase activation and mitochondrial release of cytochrome c. Preincubation of cells with N-acetyl cysteine (NAC) or pyrolidine dithiocarbamate (PDTC) protected against apoptosis mediated by PPME photosensitization showing that reactive oxygen species (ROS) are involved as second messengers. On the other hand, photosensitization carried out in the presence of deuterium oxide (D2O) which enhances singlet oxygen (1O2) lifetime only increases necrosis without aecting apoptosis. Since PPME was localized in the endoplasmic reticulum (ER)/Golgi system and lysosomes, other messengers than ROS were tested such as calcium, Bid, Bap31, phosphorylated Bcl-2 and caspase-12 but none was clearly identi®ed as being involved in triggering cytochrome c release from mitochondria. On the other hand, we demonstrated that the transduction pathways leading to NF-kB activation and apoptosis were clearly independent although NF-kB was shown to counteract apoptosis mediated by PPME photosensitization. Oncogene (2001) 20, 4070 ± 4084. Keywords: photosensitization; photodynamic therapy; pyropheophorbide; apoptosis; caspase; NF-kB Introduction Photodynamic therapy (PDT) is an emerging modality that shows considerable promise for the treatment of solid tumors and a range of non-oncologic disorders (Dougherty et al., 1998; Murphree et al., 1996; Pass, 1993). This approach is predicated on the use of a sensitizing molecule (photosensitizer) that is adminis-

*Correspondence: J Piette, Laboratory of Virology and Immunology, Institute of Pathology B23, University of Liege, B-4000 Liege, Belgium; E-mail: [email protected]. Received 2 January 2001; revised 28 March 2001; accepted 11 April 2001

tered, usually via intravenous injection, and allowed to localize somewhat selectively in cancerous tissue or other sites of therapeutic interest (e.g. atheromatous plaque, neovascular regions). Visible light irradiation is used to activate the photosensitizer yielding primarily singlet oxygen, the reactive oxygen species (ROS) believed responsible for the cytotoxic action of PDT. The speci®city achieved from drug uptake selectivity combined with light targeting makes PDT an appealing approach. The ®rst photosensitizer approved for PDT is a porphyrin oligomer (Photofrin) which is highly eective but exhibits the drawbacks of (i) a tendency to cause prolonged skin photosensitivity, (ii) an activation wavelength lower than that optimal for eective penetration through tissue, and (iii) a poorly de®ned chemical composition which makes a detailed understanding of its mode of action and pharmacokinetics dicult. To address these issues, new photosensitizers are being developed and a number of new agents are now in clinical trials. Several groups have recently reported the antitumor ecacy of pheophorbide-and pyropheophorbide-based PDT (Bellnier et al., 1993; Henderson et al., 1997; Payne et al., 1996). These compounds are chemically well characterized, absorb light above 600 nm and produce less long-term skin photosensitivity than Photofrin. In the pyropheophorbide-a series, either as methylesters or as carboxylic acids, photosensitizing activity increases with the length of the alkyl ether side-chain. These alkyl ether derivatives, although having similar photophysical properties (singlet oxygen and ¯uorescence yields), exhibit remarkable dierences in photosensitizing eciency in biological systems (Pandey et al., 1996). The cellular mechanisms of PDT are still under investigation. Singlet oxygen, the putative cytotoxic agent in PDT, is initially formed by the energy transfer from the excited triplet state of the photosensitizer to ground-state oxygen (Weishaupt et al., 1976). Several factors are thought to work together to achieve complete tumor eradication in vivo. These include direct cellular damage leading to necrosis or apoptosis, vascular destruction, in¯ammatory changes and subsequent immune responses (Korbelik, 1996). While the mechanism responsible for the eective destruction of the malignant tissues by PDT is not completely

Colon cancer cells apoptosis mediated by photosensitization J-Y Matroule et al

elucidated, many reports show that tumor cell death occurred through apoptosis, at least for photosensitizers that distribute to mitochondria (Granville et al., 1998; Kessel and Luo, 1998). In recent years, considerable progress has been made towards a molecular understanding of the processes underlying apoptotic cell death pathway(s) (Evan and Littlewood, 1998; Granville and Hunt, 2000; Jacobson et al., 1997; Steller, 1995). It is becoming clear that dierent cytotoxic stimuli acting through various mechanisms converge on the activation of caspases which instigate the well-conserved execution and terminal stages of apoptosis. However, during the induction phase of apoptosis, multiple signaling pathways in¯uence the central control of the cell death machinery. Mitochondria are situated at the crossroad between the activation and execution phases and their role as orchestrators of apoptosis is now ®rmly established in many systems (Green and Reed, 1998; Halestrap et al., 2000; Kroemer and Reed, 2000). Thus, a variety of apoptosis-related events involve mitochondria, including the release of caspase activators (such as cytochrome c), changes in electron transport, loss of mitochondrial transmembrane potential and altered cellular redox state. Mitochondria are considered the site where photosensitizers are best localized for ecient cell killing with PDT. Understanding the signals that converge on mitochondria and in¯uence these events and their downstream eects are of great signi®cance in biology. We have previously shown that photosensitization with pyropheophorbide-a methylester (PPME) of colon cancer cells activates transcription factor NFkB by mobilization of the IL-1 receptor transduction machinery. NF-kB has been shown to have either antior pro-apoptotic activity depending on the cell type or the nature of the stimulus (Van Antwerp et al., 1996; Wang et al., 1998; Ward et al., 1999). We sought to investigate the mechanism of colon cancer cell apoptosis mediated by PPME photosensitization and to clarify the role of NF-kB in this process. Because PPME mainly localizes in membranes of the endoplasmic reticulum (ER) and in lysosomes, it makes this photosensitizer an interesting compound to study mechanisms of apoptosis triggered by damage generated in organelles other than mitochondria.

Results PPME photosensitization triggers colon cancer cell death through an apoptotic mechanism Incubation of HCT-116 colon cancer cells with PPME at 6 mM followed by irradiation with red light produced a signi®cant level of cytotoxicity as indicated by the Trypan blue exclusion method. Approximatively 10% of the cells survived treament at a 96 kJ/m2 light dose (Figure 1a). Cell killing was clearly in¯uenced by the light dose used as well as the concentration of the photosensitizer applied to the cells. HCT-116 cells

unirradiated or incubated with PPME but not irradiated displayed a 100% survival. In order to determine the nature of cell death, photosensitized cells were assessed for DNA fragmentation by agarose gel electrophoresis. PPME photosensitization produced extensive dose-dependent DNA fragmentation to a degree comparable to that induced by daunomycin (Figure 1b). In a time-course study, DNA laddering was detectable by 3 h with a 96 kJ/m2 light dose (Figure 1b, lower panel). Untreated controls did not exhibit DNA fragmentation. In situ labeling of DNA cleavage products by TUNEL method reinforced these observations. PPME photosensitization gave rise to appearance of TUNEL positive cells as compared to the negative control (Figure 1c). A separate means to measure apoptotic cell death consisted of visualizing DNA condensation along the nuclear membrane by DAPI staining (Figure 1d). This condensation was evident by 3 h after photosensitization and was accompanied by nuclear disruption and the appearance of apoptotic bodies. Since caspase-3 is known to play an executioner role in apoptosis, we evaluated its involvement in PPME-mediated apoptosis. Figure 1e shows a dose-dependent stimulation of caspase-3 activity. This activity was completely abrogated by pre-incubation of HCT-116 cells with a speci®c caspase-3 inhibitor (Z-DEVD-fmk). Although HCT-116 cells expresses a functional p53 protein, stabilization of this protein was not observed in response to PPME photosensitization (data not shown). Further, PPME treatment did not induce expression of p21, a p53-encoded protein involved in the control of cell cycle (data not shown). These results indicate that PPME-mediated cell death is unrelated to p53 activation or to cell cycle arrest. In order to assess the importance of the apoptotic process in PPME-mediated cytotoxicity, HCT-116 cells were analysed by AnnexinV/PI staining method which is commonly used to dierentiate necrotic and apoptotic cell populations. As soon as 4 h after a 64 kJ/m2 or a 96 kJ/m2 irradiation, 4.8% (Figure 2a) and 7.8% (Figure 2b) of cells undergo apoptosis, respectively to reach 19.3% (Figure 2c) and 1.1% (Figure 2d) after 24 h. The ratio necrosis/apoptosis turns out to be dependent on light dose and post-irradiation time. Moreover, dierent PPME concentration used in the treatment turned out also to modulate the ratio between necrosis and apoptosis (data not shown).

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Mitochondria play a central role in PPME-mediated apoptosis To further identify the dierent intermediates transducing the apoptotic signal, it was of interest to assess the contribution of some well-characterized apoptosis factors. Firstly, PPME photosensitization gave rise to the appearance of cytochrome c in the cytosol (Figure 3). This event occurred by 2 h after PDT and was accompanied by pro-caspase-9 cleavage into active caspase-9. Activation of caspase-3, PARP cleavage and DNA fragmentation occurred in temporal proxiOncogene


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Figure 1 PPME photosensitization triggers HCT-116 cells apoptosis. (a) Survival curve for HCT-116 cells incubated with PPME (6 mM) and irradiated with increasing light doses (0 to 96 kJ/m2). Cell survival was determined by Trypan blue exclusion 24 h after photosensitization (-&-, unirradiated cells, -*- irradiation). s.d. were estimated by means of ®ve independent experiments. (b) Apoptosis in HCT-116 cells photosensitized with PPME (6 mM) as measured by DNA laddering. Upper panel: DNA laddering was detected 24 h post-irradiation in HCT-116 cells irradiated with increasing doses of light or irradiated in the absence of PPME. The positive control was HCT-116 cells treated with 2 mM daunomycin for 24 h (Dauno). Lower panel: HCT-116 cells were incubated with 6 mM PPME and then irradiated with 96 kJ/m2 before being incubated for various period of time (0 to 24 h). The state of DNA was analysed by agarose gel electrophoresis. The positive control was as above. (c) DNA cleavage products were detected by the TUNEL assay. HCT-116 cells were incubated with 6 mM PPME and irradiated with 64 kJ/m2. Nuclear DNA fragmentation was visualized by ¯uorescence microscopy (lexc: 495 nm, lem: 520 nm). The negative control was unirradiated cells. Arrows indicate TUNEL positive cells. (d) Nuclear condensation was analysed by DAPI staining. HCT-116 cells were incubated with 6 mM PPME, light-irradiated at 64 kJ/m2 and incubated for 3 h. Cells were then stained with DAPI and analysed by ¯uorescence microscopy (lexc: 340 nm, lem: 388 nm) (e) Caspase-3 activation in cells photosensitized with 6 mM PPME and irradiated with either 32 kJ/m2 (-*-), 64 kJ/m2 (-&-) or 96 kJ/m2 (-~-) of red light. Cells were also pre-treated for 45 min with 50 mM Z-DEVD before being lightirradiated at 96 kJ/m2 (-X-). Caspase 3 activity was measured by ¯uorescence spectroscopy (lexc: 350 nm, lem: 460 nm) at various time after photosensitization (from 0 to 6 h). s.d. were calculated by means of three independent experiments Oncogene


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Figure 2 Determination of necrotic vs apoptotic cells ratio by Annexin V/PI. HCT-116 cells were incubated with 6 mM PPME and then irradiated with either a 96 kJ/m2 (b and d) or a 64 kJ/m2 (a and c) light dose. Cells were harvested 4 h (a and b) or 24 h (c and d) after photosensitization and stained with Annexin V/PI for FACS analysis. Lower left panel represents living cells, lower right panel represents apoptotic cells, upper right panel represents necrotic cells and upper left panel represents pre-necrotic cells. Staining of control cells after 24 h displayed 85% living cells, 10% necrotic cells and 5% apoptotic cells

mity. Pro-caspase-8 cleavage into its active form was observed concomitant with the activation of the other caspases. However, caspase-8 did not appear to have a primary role in the initiation of the apoptotic signal. Caspase-8 activation was completely blocked by the caspase-3 speci®c inhibitor Z-DEVD (data not shown). Caspase-8 is therefore likely activated downstream of caspase-3. This suggests that PPME photosensitization does not lead to apoptosis through the recruitment of the Fas, TNF-a or related death receptors, cell surface proteins known to induce apoptosis through mobilization of caspase-8 (Schneider and Tschopp, 2000). Moreover, pre-treatment of cells with 10 mM speci®c

caspase-9 inhibitor (LEHD-CHO) completely abrogated caspase-3-like cleavage activity thereby indicating that caspase-9 is likely an initiator caspase for PDTinduced apoptosis (data not shown). Taken together, these results reveal mitochondrial events as key components in the transmission of the apoptotic signal upon PPME photosensitization. NF-kB partially protects HCT-116 cells against PPME-mediated apoptosis The role of NF-kB in apoptosis remains controversial (Kuhnel et al., 2000; Plumpe et al., 2000; Rivera-Walsh Oncogene


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downstream of cytochrome c release. Similarly, under experimental conditions that led to a lower level of apoptotic cell death (i.e., 32 kJ/m2 gave rise to a 50% cell death after 24 h), dierences in apoptosis parameters between these cell lines were also evident (data not shown). Taken together, these results reinforce the hypothesis for a protective role of NF-kB against PDT-induced apoptosis through an activation of anti-apoptotic genes. We previously showed that (i) PPME localized in the cytoplasmic membrane, lysosomes and, the ER but not mitochondria (Matroule et al., 1999a), and (ii) cytochrome c release and caspase activation kinetics were slower than for mitochondrial localizing photosensitizers such as BPD-MA or Pc 4 (Granville et al., 1998; Varnes et al., 1999). Experiments were performed to identify possible intermediates involved in the transduction of the apoptotic signal to mitochondria. Figure 3 Caspase cascade is initiated through mitochondrial changes. HCT-116 cells were treated with 6 mM PPME and then irradiated with a 96 kJ/m2 light dose. Whole cell extracts and cytosolic fractions (S-100) were prepared at various times after irradiation and separated by SDS ± PAGE followed by Westernblotting. Membranes were probed with anti-cytochrome c, anticaspase-3, -8, -9 and anti-PARP antibodies

et al., 2000). We previously demonstrated that PPME triggered NF-kB nuclear translocation by inducing the degradation of its cytoplasmic inhibitor IkBa (Matroule et al., 1999a). It was of interest to investigate the involvement of NF-kB in PPME-mediated apoptosis. The key role of Ser 32 and Ser 36 residues in IkBa degradation produced with PPME photosensitization was revealed using a HCT-116 cell line stably overexpressing a dominant negative IkBa mutant form (S32,S36A). In this line, there is no NF-kB activation after PPME photosensitization (Matroule et al., 1999a). Wild type and mutated HCT-116 cell lines were compared for their sensitivity to PPME-mediated apoptosis. The wild type IkBa overexpressing cells did not exhibit any dierence from wild type HCT-116 cells in terms of proliferation rate or PPME uptake (data not shown). However, survival rate was slightly reduced in S32,S36A cells compared to wild type cells (data not shown). In situ TUNEL labeling performed on both treated cells supported the latter observation. PDTtreated HCT-116 S32,S36A cells displayed a much larger number of TUNEL positive cells as compared to wild type cells (Figure 4a). Similarly, measurement of caspase-3 activity clearly revealed a higher inducibility in the S32,S36A cell line as compared to wild type cells (Figure 4b). When we focused on other intermediates involved in PPME-mediated apoptosis, cleavage of pro-caspase-3, -8, -9 and PARP was more pronounced for the HCT-116 S32,36A cells despite similar cytochrome c release kinetics (Figure 4c). It is possible that caspase activation may be hindered by the NF-kBencoded inhibitor of apoptosis proteins (IAP) that act Oncogene

Neither IL-1 receptor signaling machinery nor ceramide generation are involved in PPME-mediated apoptotic pathway Since IL-1R transduction machinery was shown to mediate NF-kB activation upon PPME photosensitization through the mobilization of IL-1 receptor associated proteins (Matroule et al., 1999a), it was of interest to investigate its possible involvement in the transduction of the apoptotic signal. Treatment with IL-1b (300 U/ml) did not modify basal caspase-3 activity (data not shown). To verify this observation, HCT-116 cells were transiently transfected with wild type (WT-TRAF6) or a dominant negative mutant (DTRAF6) of TRAF6 protein, an adaptator protein associated with the IL-1 receptor required for the transduction of the signal leading to NF-kB activation with PPME. Although expression of the deletion mutant of TRAF6 abrogated PPMEmediated NF-kB activation (Matroule et al., 1999a), it did not aect the activation of caspase-3 by PPME (Figure 5a). Ceramide, a second messenger involved in the IL-1 and TNF-a signaling pathways, also plays a role in NF-kB activation by PPME photosensitization through an activation of acidic sphingomyelinase (Matroule et al., 1999a). Since ceramide was indicated in TNF-induced apoptosis (Gamard et al., 1997; Monney et al., 1998), we evaluated whether ceramide contributed to PPME-mediated apoptosis. Firstly, C2ceramide was added to HCT-116 cells at various concentrations (10 ± 50 mM) and caspase-3 activity was measured 6 and 15 h later. C2-ceramide did not stimulate caspase-3 activity (Figure 5b). Secondly, generation of ceramide was blocked by inhibiting the activity of lysosomal acidic sphingomyelinase. For this purpose, HCT-116 cells were treated with 100 mM chloroquine for 1 h before photosensitization and caspase-3 activity was evaluated at dierent times after treatment (Figure 5c). No eect on caspase-3 stimulation was observed for cells pre-incubated with chlor-


oquine. NH4Cl, a separate inhibitor of acidic sphingomyelinase, also did not aect PPME-mediated caspase3 activation (data not shown).

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Reactive oxygen species (ROS) instigate the apoptotic process Since singlet oxygen (1O2) is one of the reactive oxygen species (ROS) generated by PDT and because ROS [superoxide anion (O2.7), hydrogen peroxide (H2O2), hydroxyl radical (OH.)] can also be released by mitochondria as second messenger during the apoptosis process, we decided to investigate the role of ROS in the cell death process induced by PPME photosensitization (Dumont et al., 1999; Katschinski et al., 2000; Shimura et al., 2000; Stridh et al., 1998; Zorov et al., 2000). To test their respective roles, two kinds of experiments were carried out. Firstly, HCT-116 cells were irradiated in PBS where H2O was substituted by D2O since singlet oxygen lifetime (1O2) was known to be speci®cally increased by this isotopic substitution. Cell killing was reproductibly slightly exaggerated when cells were irradiated in the presence of D2O (data not shown). This suggests that 1O2 generated during PPME photosensitization may be involved either in the necrotic or in the apoptotic process. To determine whether apoptosis could be initiated through 1 O2 generation, caspase-3 activity was measured under similar experimental conditions as above (Figure 6a). Unexpectedly, caspase-3 activation was unmodi®ed by the isotopic substitution suggesting that 1O2 generated by PPME photosensitization may control necrosis but not apoptosis. Second, PPME photosensitization was carried out in HCT-116 cells either pre-incubated with 300 mM PDTC or with 20 mM NAC as antioxidants. Survival curves obtained for cells treated with PDTC and NAC indicated a reduced sensitivity to photodynamic killing (data not shown) reinforcing the idea that other ROS than 1O2 are also involved in the cell death process mediated by PPME photosensitization. Importantly, PDTC and NAC completely or partially inhibited caspase-3 activation by PPME, suggesting that ROS others than 1O2 were involved in the apoptosis process (Figure 6b). These data were con®rmed by AnnexinV/PI staining of HCT-116 irradiated in the presence of 300 mM PDTC. We observed a 43% reduction of apoptotic cells as soon as 4 h after a 64 kJ/m2 irradiation and a 61.2% reduction 24 h after irradiation (data not shown). These results indicate that ROS (sensitive to NAC and PDTC) very likely generated through mitochon-

Figure 4 Protective role of NF-kB against PPME-mediated apoptosis. (a) TUNEL assay after a 64 kJ/m2 irradiation with 6 mM PPME in wild type HCT-116 cells (left panel) and S32,S36A IkB-a overexpressing HCT116 cells (right panel). Nuclear DNA fragmentation was visualized by ¯uorescence microscopy (lexc: 495 nm, lem: 520 nm). Arrows indicate TUNEL positive cells. (b) Comparative caspase-3 activation following a 64 kJ/m2 PPME

photosensitization in wild type cells (-&-) and S32,S36A IkB-a overexpressing cells (-*-). s.d. were calculated by means of three independent experiments. (c) Both wild type cells and S32,S36A IkB-a overexpressing cells were treated by PPME and then irradiated with a 64 kJ/m2 light dose. Whole cell lysates were separated by SDS ± PAGE and Western-blotting was performed with anti-cytochrome c, anti-caspase-3, -8, -9 and anti-PARP antibodies Oncogene


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Figure 5 IL-1 signaling pathway is not mobilized in PPMEmediated apoptosis. (a) Caspase-3 activity was measured after a 96 kJ/m2 PPME photosensitization of wild type HCT-116 cells, TRAF6 transiently expressing HCT-116 cells and DTRAF6 transiently expressing cells. (b) HCT-116 cells were treated for 6 and 15 h with various concentrations of C2 ceramide and analysed for caspase-3 activity. (c) After 1 h incubation with 100 mM chloroquine, HCT-116 cells were photosensitized and caspase-3 activity was subsequently determined. In a, b and c, s.d. were calculated by means of three independent experiments

dria contributes to the apoptotic process mediated by PPME photosensitization. On the other hand, 1O2 did not seem to play any role in the apoptotic process. The lack of involvment of 1O2 in the apoptotic pathway Oncogene

could be due to the particular localization of PPME. Indeed, several authors have demonstrated that 1O2 was implicated in apoptosis mediated by plasma membrane localizing photosensitizers like Rose Bengal (Zhuang et al., 1999; Lin et al., 2000), suggesting that the type of photosensitizer and its cellular distribution could be of importance in the role of 1O2 in apoptosis. Cytochrome c release from the mitochondrial interspace is often related to the formation of permeability transition pores which allow small size molecules to cross the mitochondrial membrane (Hirpara et al., 2000; Tafani et al., 2000). To determine whether the permeability transition pore was responsible for the cytochrome c release observed following PPME photosensitization, cells were pre-incubated for 30 min with 5 mM cyclosporin A (CSA), an immunosuppressive agent known to block the pore opening (Lemasters et al., 1998). Although CSA abrogated caspase-3 activation by H2O2, it had no eect on caspase-3 activation by PPME (Figure 6c). Taken together, these results suggested that PPME photosensitization promoted cytochrome c release from mitochondria through a mechanism independent of permeability transition pore but involving ROS production. This mechanism was already observed upon treatment of CEM and HeLas cells with Staurosporin or UV-B (Bossy-Wetzel et al., 1998). We postulate that either PPME induces ROS formation within mitochondria leading to a membrane destabilization enabling the release of cytochrome c or PPME generates ROS which in turn permeabilizes the mitochondrial membrane to release cytochrome c. To further support the role of ROS in the apoptotic process initiated by PPME, several respiratory chain inhibitors including rotenone, antimycine A, myxothiazol and oligomycin were shown to strongly reduce PPME photosensitization induced-caspase 3 activation (data not shown) likely by disrupting ATP synthesis coupled to respiration. Indeed, ATP is required for Apaf-1-mediated caspase-9 activation (Saleh et al., 1999). PPME was shown to be distributed to membranes and in lysosomes, it was of interest to determine whether others secondary messengers than ROS arose from these cellular sites. Bap31, a shuttle protein acting between ER and Golgi apparatus and capable of binding Bcl-2 and caspase-1 and -8, was reported by Granville et al. (1998) to be cleaved into its active form by PDT mediated by BPD-MA in HeLa cells. However, Bap31 cleavage was not observed for HCT116 cells following PPME photosensitization (data not shown). Bid cleavage by caspase-8 has often been observed in response to TNF-a treatment or oxidative stress (Gross et al., 1999; Zhuang et al., 2000). Bid p15 fragments targets mitochondria and triggers cytochrome c release in the cytosol. However, no Bid cleavage was pointed out upon PPME photosensitization despite caspase-8 activation (data not shown). Ca2+ has been shown to play a role in PDT-mediated apoptosis (Ruck et al., 2000; Tajiri et al., 1998). Since ER constitutes an important Ca2+ store, we hypothe-


sized that PPME might trigger Ca2+ release to the cytosol. Therefore, cells were pre-incubated with the Ca2+ chelator, BAPTA-AM, before photosensitization and assessed for subsequent caspase-3 activity. No alteration in caspase-3 activity was observed for BAPTA-AM treated cells (data not shown) indicating that either Ca2+ was not released from ER store or Ca2+ is not a messenger in the apoptotic process mediated by PPME photosensitization. The last intermediate tested was caspase-12 which was recently shown to be involved in ER-speci®c apoptosis and cytotoxicity by amyloid-b in neuronal cells (Nakagawa et al., 2000). In HCT-116 cells photosensitized by PPME, no procaspase-12 cleavage could be visualized and no inhibition of caspase-3 activation or PARP cleavage was visualized after transfecting photosensitized HCT-116 cells with antisense oligonucleotides (data not shown).

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O2 mainly generated from the ER and Golgi membranes is involved in necrosis and not in apoptosis

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Figure 6 ROS participate in PPME-mediated apoptosis. (a) Caspase-3 activity levels were measured after a 96 kJ/m2 irradiation in either PBS-H2O or PBS-D2O. Caspase-3 was determined on whole cell extracts. (b) Caspase-3 activity in HCT-116 cells were pretreated for 1 h with either PDTC (300 mM) or NAC (20 mM) before photosensitization with a 96 kJ/m2 light dose. Caspase-3 activity was measured for whole cell extracts. (c) Caspase-3 activity measurements when 5 mM cyclosporin A (CSA) was added to HCT-116 cells 30 min prior 96 kJ/m2 irradiation. Positive control was made by treating the cells for 24 h with 500 mM H2O2. Caspase-3 activity was measured for whole cell extracts. s.d. were calculated by means of three independent experiments

Since 1O2 lifetime and range of diusion are quite short in cells, it was of interest to identify its production site in order to better correlate the molecular mechanisms to oxidative status. For that purpose, we used dichloro¯uorescein staining method which is commonly used to monitor intracellular ROS production (Hockberger et al., 1999). In the present study, cells preincubated with PPME were subsequently loaded with 6-carboxy-2',7'-dichlorodihydro¯uorescein diacetate di(acetoxymethyl ester) (C-DCDHF-DA-AM) and observed by ¯uorescence microscopy with a FITC ®lter. As shown in Figure 7b ± d, PPME photosensitization under the ®ltered light of the microscope immediately triggered C-DCDHF-DA-AM oxidation as visualized by yellow ¯uorescence. In Figure 7e, the 100-fold magni®cation allowed us to visualize, at early irradiation times, a perfectly similar ¯uorescence pattern between oxidized C-DCDHF-DA-AM (left panel) and PPME (right panel). These data demonstrate that PPME photosensitization triggered the production of 1O2 by photoexcited PPME localized in the ER and the Golgi apparatus. We could not detect any colocalization between C-DCDHF-DA-AM and mitochondria (Figure 7e) reinforcing the idea that the probe detected 1O2 generated by PPME photosensitization and not ROS released by mitochondria. Importantly, the C-DCDHF-DA-AM ¯uorescence was quenched by a pre-incubation with 1 mM mannitol 1 O2 inhibitor (Figure 8a,b), a well-known (k1O24108 M71 s71), but not by PDTC and NAC (k1O25105 M71 s71) (Figure 8c,d). This suggests that PDTC and NAC could not eciently quench the 1O2 production by PPME photosensitization and acted downstream in the oxidative process. From these experiments we can conclude that PPME photosensitization led to 1O2 generation from the ER/Golgi membranes. In order to determine whether or not this 1 O2 production was linked to the apoptosis or to the necrosis process, we used the FACS coupled Annexin Oncogene


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Figure 8 C-DCDHF-DA-AM oxidation is inhibited by mannitol but not by PDTC and NAC. PPME pre-incubated HCT-116 cells were pre-incubated for 2 h with 1 mM mannitol (b), 300 mM PDTC (c), 20 mM NAC (d) or without any antioxidant (a) and then loaded with 37 mM C-DCDHF-DA-AM for 45 min. Coverslips were mounted on glass and observed under FITC ®lter equiped microscope with a 100-fold magni®cation

Figure 7 Singlet oxygen is mainly generated in the ER/Golgi network. PPME pre-incubated HCT-116 cells were loaded with 37 mM C-DCDHF-DA-AM for 45 min in PBS. Coverslips were mounted on glass and observed under ¯uorescence microscopy for 1 s (b), 3 s (c) and 6 s (d). Control cells are represented in (a). Cells shown in e corresponds to a 100-fold magni®cation under FITC ®lter (left panel) or Texas Red ®lter (right panel)

V/PI staining. HCT-116 cells were then photosensitized by PPME and incubated in the presence of mannitol (1 mM) or photosensitized with PPME when water was substituted with D2O in order to increase 1O2 lifetime. Incubation in the presence of 1 mM mannitol reduced by 50% the number of early necrotic cells (upper left panel) and by 20% the whole necrotic cell population (upper left and right panels) 5 h following photosensitization without signi®cantly aecting apoptotic cells (lower right panel) (data not shown). Conversely, replacement of H2O by D2O doubled the number of early necrotic cells and increased by 30% the whole necrotic population without aecting apoptosis (data not shown). These data demontrated that 1O2 produced from the ER/Golgi membranes is mainly involved in necrosis whereas ROS other than 1O2 which were likely produced via the mitochondria are linked to the apoptosis pathway.

By using S70A Bcl-2 mutant and Dloop Bcl-2 deletion mutants, phosphorylation at this residue inhibited the anti-apoptotic function of Bcl-2 (Srivastava et al., 1999). Because several drugs used in PDT were shown to activate MAP kinases (Assefa et al., 1999; Tao et al., 1996), we investigated Bcl-2 phosphorylation to assess a potential link between PPME localization to the ER/ Golgi and subsequent cytochrome c release from mitochondria. Bcl-2 underwent modi®cation following PPME photosensitization as evidenced by the presence of a band with slower electrophoretic mobility corresponding to the phosphorylated form of Bcl-2 (Figure 9a). This phosphorylation was evident within 2 h after PDT and was speci®c for Bcl-2 since no equivalent alteration in electrophoretic mobility of Bcl-XL was observed with the treatment. To further relate this phosphorylation event to PPME-mediated apoptosis, cells were transiently transfected with expression plasmid encoding wild type Bcl-2, dominant negative Bcl-2 S70A or dominant negative Dloop Bcl-2. PPME-induced caspase-3 activation, cytochrome c release and PARP cleavage were assessed in the transfected cells (Figure 9b). Because there was no detectable dierence in the extent of caspase-3 activity, cytochrome c release or PARP cleavage, we concluded that Bcl-2 phosphorylation status likely does not in¯uence PPME-mediated apoptosis.

PPME photosensitization triggers Bcl-2 phosphorylation Srivastava and collaborators demonstrated that Paclitaxel treatment gave rise to a MAP kinase dependent Bcl-2 phosphorylation (Srivastava et al., 1999). This phosphorylation occurs in the Dloop region on Ser 70. Oncogene

Discussion In this paper we have shown that PPME, a second generation non-mitochondrial localizing photosensi-


Figure 9 Bcl-2 phosphorylation does not contribute to PPMEinduced apoptosis. (a) Whole cell extracts from photosensitized cells were separated by SDS ± PAGE (12%) and analysed by Western-blotting with either Bcl-2 antibody (upper panel) or BclXL antibody (lower panel). (b) Wild type HCT-116 cells, S70A or Dloop Bcl-2 transiently overexpressing cells were photosensitized at a 96 kJ/m2 light dose. Whole cell extracts and cytosolic fractions (S-100) were prepared 5 h following irradiation and assessed for caspase-3 activity (upper panel), cytochrome c release (middle panel) and PARP cleavage (lower panel). HS indicates herring sperm DNA used as negative control for transfection

tizer, can rapidly trigger apoptosis despite having an ER/Golgi and lysosomal distribution in cells. Apoptosis generated in colon cancer cells by PPME photosensitization does not depend on p53 activation/ stabilisation but leads to the release of cytochrome c from mitochondria and activation of caspase-9 and -3. We have also demonstrated that apoptosis does not originate from neither mobilization of IL-1 receptor transduction machinery nor ceramide generation by acidic sphingomyelinase as shown for NF-kB activation by this compound (Matroule et al., 1999a). The apoptosis initiation mechanism appears classic with the exception of the existence of a critical link between the main localization sites of PPME and mitochondria. This connection is likely based on ROS generation but not on Bid cleavage, Ca2+ release from stores or

cleavage of Bap-31, a pro-apoptotic ER molecule (Figure 10) (Granville et al., 1998). Cellular localization studies with PPME or aminopyropheophorbide (APP) (Matroule et al., 1999a,b) with ¯uorescence microscopy using organelle speci®c probes, revealed that PPME mainly localized in the ER/Golgi apparatus and lysosomes. Lysosomes are believed to quench photodynamic activity by acting as a sink which lowers the active drugs concentrations (MacDonald et al., 1999). It is thus likely that the active cellular site for triggering apoptosis mediated by PPME photosensitization is the ER/Golgi system. The absence of co-localization with the mitochondrial probe rhodamine-123 indicates the likelihood that the ER/Golgi is the site from which the apoptotic signal originates (Matroule et al., 1999b). Others have found that photosensitizers localize to a variety of cellular sites including the outer membrane, lysosomes, mitochondria and the nucleus resulting in damage to these structures upon light exposure (Morgan et al., 2000; Oleinick and Evans, 1998). Although mitochondria are primary targets for some photosensitizers, other organelles were also considered as sites of action (Granville and Hunt, 2000; Lee et al., 1995; Miller et al., 1995). The present data provides evidence that internal membranes, particularly, the ER/Golgi apparatus is a distinct site from which an apoptotic signal can emerge. ROS turns out to be intermediates in the apoptotic process following PPME photosensitization. ROS generated from mitochondria by a perturbation of the electron transport chain may occur subsequent to the primary photodynamic events in the ER/Golgi. The mechanism that causes mitochondria to produce ROS in response to apoptotic stimuli is still unclear but several mitochondrial regulators have been identi®ed such as Bax-like proteins, nitric oxide, ceramide, ATP/ ADP depletion, NADH oxidation (Kroemer and Reed, 2000). Among them, none were clearly identi®ed as emerging from the ER/Golgi where the photosensitization by PPME is triggered. Recently, caspase-12 was shown to mediate ER-speci®c apoptosis and cytotoxicity of neuronal cells by amyloid-b (Nakagawa et al., 2000). Using antibodies directed against caspase-12 and antisense oligonucleotides we were unable to demonstrate processing of pro-caspase-12 and an inhibition of apoptosis mediated by PPME photosensitization ruling out the role of caspase-12 as mediator in our system. The most likely possibility is that other ROS than 1 O2 (O2.7, H2O2 or OH.) produced at the primary site of the photosensitization reaction (in the ER/Goli network) act as second messengers causing mitochondria to release cytochrome c and, via caspase-3 activation, apoptosis. This hypothesis was recently supported by the observation that ROS accumulation in individual mitochondria in isolated cardiac myocytes reproductibly triggered abrupt mitochondrial depolarization which coincided with a burst of mitochondrial ROS generation (Zorov et al., 2000). On the other hand, Jurkat cells treated with oxidants such as H2O2 or tributylin, rapidly caused mitochondria to release

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Figure 10 Model of PPME photosensitization-mediated apoptosis in HCT-116 cells

cytochrome c followed by eector caspase activation and apoptosis (Dumont et al., 1999; Kim et al., 2000). From these reports, it could be anticipated that alternative mechanisms for cytochrome c release exist and these may be due to a direct eect of ROS on mitochondria. Indeed, treatment of isolated mitochondria with tert-butylhydroperoxide leads to opening of the permeability transition pore and release of mitochondrial intermembrane proteins (Kushnareva and Sokolove, 2000). Signi®cantly, CSA which inhibits formation of permeability transition pores, had no eect on the extent of caspase-3 activation by PPME photosensitization. We propose that cytochrome c Oncogene

release following mitochondria injury does not involve pore formation but rather membrane leakage. This conclusion is warranted by the observation that transfection with either wild type or mutated Bcl-2 does not modify cytochrome c release or caspase-3 activation with PPME photosensitization. Indeed, antiapoptotic Bcl-2 members act by preventing the dissipation of the mitochondrial membrane potential, pore formation and cytochrome c release (Gottlieb et al., 2000; Shimizu et al., 1999, 2000). Since overexpression of Bcl-2 does not modify the extent of apoptosis mediated by PPME, cytochrome c release by mitochondria is not likely due to pore formation. A


similar circumstance was described for apoptosis induced either by UV or staurosporine (Bossy-Wetzel et al., 1998). In these situations, the release of cytochrome c to the cytosol preeceded mitochondrial membrane depolarization which was dependent on caspase activity. Another important feature demonstrated in this paper is a clear separation between the transduction pathways initiated by PPME photosensitization leading to NF-kB activation and apoptosis. NF-kB activation is ROS-independent but requires IL-1 receptor internalization and recruitment of its signal transduction machinery whereas apoptosis is ROS-dependent and does not involve any components of the IL-1R transduction machinery. This is the ®rst description that two very important cellular phenomena initiated by a photosensitization reaction can be clearly separated in terms of the mechanisms involved. However, these two distinct pathways are inter-related because apoptosis is more extensive when NF-kB activation is prohibited by over-expression of a dominant negative form of IkBa. All apoptosis parameters examined except cytochrome c release, were more pronounced for the IkBa mutated cell line. Several reports show that in dierent cell types, NFkB controls genes encoding proteins that exert antiapoptotic eects. For example, binding of TNF-a to its receptor may initiate apoptosis and concomitantly activate NF-kB, which suppresses apoptosis. TRAF1 (TNFR-associated factor 1), TRAF2, and the inhibitor of apoptosis (IAP) proteins c-IAP1, c-IAP2, A1, A20, Bcl-XL have been identi®ed as target genes of NF-kB transcriptional activity (Green, 2000; Hu et al., 1998; Wang et al., 1998, 1999). In cells in which NF-kB is inactive, expression of all the above mentioned proteins have been shown to be required to suppress TNFinduced apoptosis, whereas c-IAP1 and c-IAP2 are sucient to suppress etoposide-induced apoptosis (Wang et al., 1998). One important issue of the work presented above will be to identify the anti-apoptotic genes in¯uenced by NF-kB following PPME-mediated photosensitization. In conclusion, we have shown for the ®rst time that an ER/Golgi-localized photosensitizer can trigger an important apoptosis in colon cancer cells. The eector of apoptosis are very likely to be ROS other than 1O2 causing mitochondria to release cytochrome c initiating the subsequent late stages of apoptosis. This mechanism is clearly separated from the one leading to NF-kB which was shown to be activated by the recruitment of the IL-1 transduction machinery (Matroule et al., 1999a) and demonstrated in the present manuscript to counteract apoptosis. Materials and methods Chemicals and reagents Pyropheophorbide methylester (PPME) was from Sigma (Sigma Chemical Co., St. Louis, MO, USA) and used without modi®cation. A stock solution was made in ethanol

(1 mM) and kept in the dark at 7208C. PPME was diluted in the culture medium just before use and added to exponentially growing cells. Deuterium oxide (99.8% purity) was from Merck (Darmstadt, Germany). Antibodies were obtained from the following sources: rabbit anti-caspase-3 and mouse anti-caspase-8 (Upstate Biotechnology Inc., Lake Placid, NY, USA), mouse anti-caspase-9 and anti-cytochrome c (Pharmingen, Mississauga, Ont., Canada), rat anti-human caspase-12 was a generous gift from J Yuan (Harvard Medical School, USA), mouse anti-poly(ADPribose) polymerase (PARP) (Biomol Research Laboratories, Plymouth Meeting, PA, USA), rabbit anti-PARP p85 fragment (Promega, NL, Canada), mouse anti-Bcl-2 and anti-Bcl-XL (Santa Cruz Biotechnology, USA). Caspase-3 and caspase-9 peptide inhibitors were from Calbiochem (San Diego, USA). Chicken anti Bap-31 antibody was a generous gift from Dr Gordon Shore (McGill University, Quebec, Canada). All other chemicals were of reagent grade.

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Cell culture The human colon carcinoma cell line HCT-116 was grown in McCoy's 5A medium (Biowhittaker, Belgium) supplemented with 10% fetal calf serum (FCS, Gibco-BRL, UK). HCT-116 cells overexpressing IkBa mutated at serines 32 and 36 (S32,36A) were generated as described (Matroule et al., 1999a) and cultivated in the presence of neomycin (250 mg/mL). Plasmids pTRAF6 and pDTRAF6 (289 ± 522) constructs were gifts from D Goeddel (Tularik, San Francisco, USA). pS70A Bcl-2 and pDloop Bcl-2 were generous gifts from D Longo (National Institute on Aging, Baltimore, USA). Wild type Bcl-2 was kindly provided by C Borner (Fribourg University, Switzerland). All plasmids were puri®ed using Qiagen column chromatography (Qiagen, NL, Canada) and their integrity was checked by agarose gel electrophoresis. Exposure of HCT-116 cells to PPME photosensitization Before photosensitization with PPME, HCT-116 cells were cultivated in 25 cm2 ¯asks for three days in McCoys 5A medium with 10% FCS and incubated with 6 mM PPME during the last 20 h in McCoys 5A medium with 2% FCS. Prior to irradiation, HCT-116 cells were washed once with PBS and then irradiated with red light (l4600 nm) at various ¯uence rate covered with PBS. After irradiation, HCT-116 cells were returned in culture at 378C in McCoys 5A medium supplemented with 10% FCS. Cell survival was determined after 24 h using Trypan blue exclusion. Transient transfection assays HCT-116 cells were grown in 25 cm2 ¯ask for 2 days in McCoys 5A medium supplemented with 10% FCS and transfected with 5 mg expression plasmids. Plasmids were mixed in McCoys 5A medium, added to Fugene liposomes (7 ml) (BoehringerMannheim, Germany) for 15 min at room temperature and loaded on cells in 2 ml of McCoys 5A containing 10% FCS for 24 h. Then, HCT-116 cells were incubated with PPME (6 mM) for 24 h and irradiated with red light. Whole cell lysate extraction After photodynamic treatment, cells were collected by scraping, washed once in ice-cold PBS and treated with lysis Oncogene


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buer (1% Nonidet P-40, 20 mM Tris pH 8.0, 137 mM NaCl, 10% glycerol) supplemented with 1 mM phenyl-methylsulfonyl ¯oride, 1 mM sodium orthovanadate and anti-proteases complex (Complete, Boehringer Mannheim, Germany) for 15 min on ice and further centrifuged at 10 000 g for 15 min. Protein concentrations were determined using the bisinconinic acid method (micro BCA, Pierce, USA). For the phosphoBcl-2 assay, a phosphatase inhibitor mix was added to lysis buer as described by Kroll et al. (1999).

conditions followed by Western blotting. Membranes were incubated for 45 ± 120 min with primary antibody at room temperature. Membranes were then probed with horseradish peroxidase-labeled anti-mouse IgG (1/2000 ± 1/5000), antirabbit IgG (1/2000 ± 1/5000) and anti-rat IgG (1/1000) in PBS, 0.05% Tween 20, 5% (w/v) milk powder for 30 ± 60 min at room temperature. Proteins were detected using a chemiluminescence detection system (Amersham, Arlington Heights, IL, USA) and bands visualized by autoradiography.

Preparation of S-100 fraction (cytosolic extracts)

AnnexinV ± PI staining

Harvested cells were washed once with cold PBS and resuspended in ice-cold buer (20 mM Tris-HCl pH 8.0, 137 mM NaCl, 10% glycerol) supplemented with 1 mM phenyl-methylsulfonyl ¯oride, 1 mM sodium orthovanadate and an anti-protease cocktail (Boehringer Mannheim, Germany). Cells were disrupted using a Kontes dounce homogenizer. Lysates were centrifuged at 10 000 g for 10 min and the supernatant was further ultracentrifuged at 100 000 g for 1 h in a Beckman Airfuge (Analis, Belgium). Protein amounts were measured using Bradford method (Bio-Rad protein assay, Bio-Rad, Germany).

HCT-116 cells were cultivated in six-well plates for 24 h in McCoy's medium supplemented with 10% FCS and then incubated for another 24 h with 6 mM PPME in McCoy's medium supplemented with 2% FCS before photosensitization. Photosensitized cells were gently scraped and pooled with medium ¯oating cells. Cells were individualized in PBSEDTA (10 mM) and stained as described by the manufacturer (Roche, Germany). Brie¯y, cell pellet was resuspended in 500 mL AnnexinV-HEPES solution (10 mM HEPES-NaOH, pH 7.4, 140 mM NaCl, 5 mM CaCl2) and incubated on ice for 30 min in the dark. Cells were then washed once in ice-cold HEPES buer and PI was added just before FACS analysis. Cells were analysed with a FacsCalibur (Becton-Dickinson, Sunnyvale, CA, USA).

DNA laddering After photosensitization, HCT-116 cells were returned to culture for various times as described above and washed once in PBS before being trypsinized, pooled with cells recovered from the supernatant and centrifuged at 5000 g for 2 min. Cells were washed again in PBS and treated with two cycles of lysis buer (1% NP-40 in 20 mM EDTA, 50 mM Tris-HCl, pH 7.5; 10 ml per 106 cells, minimum 50 ml). After centrifugation for 5 min at 1600 g the supernatant was collected and treated with RNAse A for 120 min at 568C and brought to 1% (w/v) SDS, followed by digestion with proteinase K (®nal concentration 2.5 mg/ml) for at least 120 min at 378C. DNA was then precipitated and analysed by gel electrophoresis in 1% agarose gels.

A cell-free caspase-3 assay was performed as described by Granville et al. (1998). Brie¯y, 50 mg of whole cell lysate was incubated at 378C for 4 h with 20 mM Ac-DEVD-AMC as substrate (Calbiochem, San Diego, USA). Fluorescence was measured using a VictorTM 1420 multilabel counter (Wallac, Sweden) or CytoFluor 2350 (PerSeptive Biosystems, Canada) set at 360 and 460 nm for excitation and emission respectively. Z-DEVD-fmk was provided by Enzyme Systems Products (Livermore, USA).

DAPI and TUNEL assays

Fluorescence detection of ROS generation

Various times after photosensitization, HCT-116 cells were washed with PBS and then washed once with 4', 6-Diamine2'-phenylindole (DAPI) (1 mg/ml, Boehringer, Mannheim, Germany) in methanol and incubated for 15 min at 378C. After staining cells washed with methanol and placed on coverslip to be observed by ¯uorescence microscopy (lexc: 340 nm, lem: 388 nm). TUNEL (terminal deoxyuridine nickend-labeling) assays were performed by using TUNEL label mix and tunel enzyme (Boehringer, Mannheim, Germany) according to the procedure described by the manufacturer. Brie¯y, after photosensitization cells were cultured in McCoy's medium supplemented with 10% FCS for dierent periods of time until ®xation. Cells were then washed twice in PBS, ®xed in paraformaldehyde (4%) and permeabilized in 70% ethanol. After two washes in PBS, cells were labeled by ¯uorescein-dUTP in the presence of terminal deoxynucleotidyl transferase (in situ cell death detection kit, Boehringer Mannheim, Germany) and incubated for 60 min at 378C in a humidi®ed atmosphere in the dark. After several washes in PBS, cells were analysed by ¯uorescence microscopy (Nikon Eclipse E800, Japan) using ®lters for ¯uorescein.

HCT-116 cells were grown for 24 h on coverslips in McCoy's medium supplemented with 10% FCS. After a 24 h incubation with 6 mM PPME, cells were loaded at RT in the dark with 37 mM membrane permeable 6-carboxy-2',7'dichlorodihydro¯uorescein diacetate, di(acetoxymethyl ester) (C-DCDHF-DA-AM) (Molecular Probe, USA) for 45 min. Coverslips were rinsed once in PBS to remove the excess of probe and then mounted on glass for microscopic observations. Photosensitization reaction was iniated with the blue light emitting microscope FITC ®lter and C-DCDHF-DAAM ¯uorescence was analysed under the same conditions.

Immunoblot analysis Detergent soluble proteins (30 ± 50 mg) were loaded and separated by SDS ± PAGE (10 ± 12% gels) under reducing Oncogene

Protease assay

Abbreviations PDT, photodynamic therapy; PPME, pyropheophorbide-a methylester; FCS, foetal calf serum; PARP, poly (ADPribose) polymerase; DAPI, 4',6-diamidine-2'-phenylindole dihydrochloride; TUNEL, terminal deoxyuridine nick-endlabeling; ROS, reactive oxygen species; 1O2, singlet oxygen; H2O2, hydrogen peroxide; IL, interleukin; TNF-a, tumor necrosis factor a; ER, endoplasmic reticulum; CSA, cyclosporin A; PDTC, pyrrolidine dithiocarbamate; NAC, N-acetyl cysteine; D2O, deuterium oxide, C-DCDHF-DAAM, 6-carboxy-2',7'-dichlorodihydro¯uorescein diacetate di(acetoxymethyl ester)


Acknowledgments This work was supported by a grant from the Belgian National Fund for Scienti®c Research (NFSR) (Brussels,

Belgium) and TeÂleÂvie (Brussels, Belgium). J-Y Matroule is Research fellow from the FRIA and J Piette is Research Director from the NFSR (Brussels, Belgium).

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