Aplidin™, a new antitumoural drug presently in phase II clinical trials, has shown both in vitro and in vivo activity against human cancer cells. Aplidin™ effectively inhibits cell viability by triggering a canonical apoptotic program resulting in alterations in cell morphology, caspase activation, and chromatin fragmentation. Pro-apoptotic concentrations of Aplidin™ induce early oxidative stress, which results in a rapid and persistent activation of both JNK and p38 MAPK and a biphasic activation of ERK. Inhibition of JNK and p38 MAPK blocks the apoptotic program induced by Aplidin™, demonstrating its central role in the integration of the cellular stress induced by the drug. JNK and p38 MAPK activation results in downstream cytochrome c release and activation of caspases -9 and -3 and PARP cleavage, demonstrating the mediation of the mitochondrial apoptotic pathway in this process. We also demonstrate that protein kinase C delta (PKC-δ) mediates the cytotoxic effect of Aplidin™ and that it is concomitantly processed and activated late in the apoptotic process by a caspase mediated mechanism. Remarkably, cells deficient in PKC-δ show enhanced survival upon drug treatment as compared to its wild type counterpart. PKC-δ thus appears as an important component necessary for full caspase cascade activation and execution of apoptosis, which most probably initiates a positive feedback loop further amplifying the apoptotic process.
Aplidin™ (Figure 1), a marine cyclic depsipeptide derived from the Mediterranean tunicate Aplidium albicans (Rinehart, 2000), is a new anticancer agent with potent anti-neoplastic activity in vitro against a wide variety of human tumours. Aplidin™ was chosen for development as an antineoplastic agent because of its antitumour activity against subcutaneous implanted MRI-H254 gastric, PC-3 prostate and Burkitts lymphoma human xenografts as well as HTB-9 bladder carcinoma in the hollow fiber model (Faircloth et al., 1998, 1999). Clinical studies with Aplidin were initiated in early 1999 in different locations of Europe and Canada. Previous results from phase I clinical trials, including more than 200 treated patients, have shown hints of activity, in terms of stabilization of the disease during long-term intervals, in renal, medullary thyroid carcinoma and other neuroendocrine tumours, lung (NSC), head and neck and colorectal cancers. Anti-tumour activity in advanced resistant tumours has also been observed. On this basis, phase II disease oriented clinical studies are ongoing (Anthoney et al., 2000; Armand et al., 2001; Bowman et al., 2001; Ciruelos et al., 2002; Mauroun et al., 2001). Up to date, 17 patients have been already recruited for these studies (unpublished data). Little is known concerning the mechanism of action of Aplidin™. In leukaemia cells, it has been suggested that Aplidin™ causes a G1 blockade and apoptosis, and inhibition of ornithine decarboxylase (ODC) (Erba et al., 1999). Also in leukaemia cells, Aplidin™ seems to cause a reduction in vascular endothelial growth factor (VEGF) secretion and downregulation of its receptor, VEGFR-1 (flt-1), involved in the process of vascularization and growth of certain tumours (Broggini et al., 2000). In addition, it has been described that its homologue didemnin B may interact with elongation factor 1a (Crews et al., 1994) and palmitoyl thioesterase (Crews et al., 1996).
Apoptosis, or programmed cell death, is important for the destruction of undesired cells during development as well as tumour cells and other damaged cells. Consequently, an increase or decrease in apoptosis may contribute to the pathology of a wide range of disorders including those associated with development, autoimmune disease, and cancer (Jacobson and Weil, 1997; Thompson, 1995). Apoptosis is characterized by alterations in plasma membrane, cell shrinkage, depolarization of mitochondria, protein degradation, chromatin condensation, and DNA fragmentation (Wyllie et al., 1980). The critical genes in the apoptotic process have been characterized (Rao and White, 1997). These include the Bcl-2 family of proteins, a family of related regulatory proteins which either promote or suppress apoptosis (Kroemer, 1997), and the caspases, cystein proteases that are responsible for initiation and execution of the apoptotic signal (Nuñez et al., 1998). Caspases catalyse a controlled proteolysis of proteins, including caspases themselves and their downstream substrates. Among them, the effector downstream caspase-3 has been shown to play a pivotal role in the terminal, execution phase of apoptosis induced by a variety of stimuli (Tewari et al., 1995).
In addition to caspases, it is becoming increasingly clear that specific protein kinases signal transduction pathways are involved in mediating apoptosis, including members of the mitogen-activated protein kinases family (MAPK) and protein kinase C (PKC) (Chen et al., 1996a; Kummer et al., 1997; Lucas and Sánchez-Margalet, 1995; Osborn and Chambers, 1996). Thus, the integration and balance of these pathways probably do contribute to commitment to apoptosis.
Mitogen-activated protein kinase (MAPK) pathways are parallel cascades of structurally related serine-threonine kinases that serve to integrate numerous extracellular signals, resulting in regulation of cell proliferation, differentiation and cell survival and adaptation (for reviews, see English et al., 1999; Seger and Krebs, 1995). Among the known MAPKs, three groups have been extensively studied: the extracellular signal-regulated kinases (ERK-1/2), which are generally involved in cell growth and differentiation and the c-Jun NH2-terminal kinases (JNK-1/2/3) and p38 MAPK proteins (p38α/β/γ/δ), both activated by environmental stress such as UV light, γ-irradiation, DNA damage, protein synthesis inhibitors, heat shock, osmotic shock, or oxidative stress (English et al., 1999; Hagemann and Blank, 2001; Robinson and Cobb, 1997; Schaeffer and Weber, 1999; Seger and Krebs, 1995). JNK and p38 MAPK have been implicated in the regulation of the apoptotic cell death (Anderson, 1997; Chen et al., 1996a; Xia et al., 1995) induced by different stimuli, including anticancer (Huang et al., 1999; Shtil et al., 1999; Yu et al., 1996b) and chemo preventive agents (Brenner et al., 1997; Chen et al., 1998; Ichijo et al., 1997; Schwenger et al., 1997; Seimiya et al., 1997; Yu et al., 1996a).
Protein kinase C (PKC) represents a family of serine/threonine protein kinases, which presently consists of at least 11 isozymes whose expression varies between cell types. A variety of studies indicate that specific isoforms of PKCs may be either pro-apoptotic or anti-apoptotic, depending on the stimulus or the cell type (Leszczynski, 1995; Lucas and Sánchez-Margalet, 1995).
In the present study, we have examined the molecular events induced by Aplidin™ in human HeLa tumour cells, characterizing the apoptotic program which causes its cytotoxicity. These studies demonstrate that Aplidin™ induces early oxidative stress, JNK and p38 activation, and the induction of the mitochondrial apoptotic pathway including cytochrome c release and caspase cascade activation, and also proteolytic cleavage of PKC-δ, which generates a positive feedback loop amplifying the apoptotic cascade induced by the drug.
Time-dependent effects of Aplidin™ on the viability of HeLa cells
Treatment of HeLa cells with 0.4 μM Aplidin™ caused a fast, time-dependent reduction of cell survival (about 50 and 100% after 6 and 12 h, respectively) (Figure 2a) and typical phenotypic changes associated with apoptosis such as cell shrinkage, loss of cell to cell contact, membrane blebbing and chromatin condensation (not shown). Fluorescent staining of chromatin with Hoechst 33342, revealed the presence of apoptotic bodies in non-viable cells upon drug treatment (Figure 2b). DNA content analysis by flow cytometry showed that Aplidin™ induced a rapid and progressive accumulation of cells in the sub-G1 region (up to 50% at 6 h post-treatment). Interestingly, no significant cell cycle arrest was detected before the onset of apoptosis (Figure 2c). Aplidin™ also induced progressive accumulation of extranuclear DNA, which appeared as a typical ladder due to internucleosomal cleavage associated with apoptosis (Figure 2d).
Oxidative stress on Aplidin™-mediated apoptosis
Oxidative stress and glutathione homeostasis are well established as integral control elements in the cell's decision to enter apoptosis (Poot et al., 1995; Schnelldorfer et al., 2000). To assess whether Aplidin™-induced apoptosis involves generation of cellular oxidative stress, the viability of drug-treated cells was tested in the presence of different radical scavengers (vitamin E, NAC, ebselen, tiron, catalase and DPI) (not shown). Only ebselen (2-phenyl-1,2-benzisoselenazol-3(2H)-one), a permeable organo-selenium compound with potent antioxidant properties and glutathione peroxidase mimetic activity, showed a noteworthy protective effect on Aplidin™-treated cells, as assessed by chromatin morphology and flow cytometry. When cells were preincubated with 40 μM ebselen, the percentage of apoptotic cells decreased significantly (Figure 3a) and most nuclei appeared healthy and uniformly stained (Figure 3b).
Aplidin™ induces JNK, p38 MAPK and ERK activation
The effect of Aplidin™ on MAPKs activity, was assessed by testing the status of representative members of the MAPK pathways, that is, JNK, p38 MAPK and p42/p44-MAPK (ERK). Aplidin™ induced strong and persistent JNK and p38 MAPK phosphorylation with activation of both kinases occurring very rapidly, long before the execution of apoptosis, and full activation within 5–10 min of drug treatment (Figure 4a). Aplidin™ also induced ERK phosphorylation, in a time-dependent biphasic manner with activation peaking first at 15 min and then again 3–4 h post-treatment (Figure 4b). Re-probing blots with antibodies against total JNK1, p38 MAPK and ERK2 revealed no change of proteins upon treatments, indicating that Aplidin™ induced phosphorylation of pre-existing MAPKs rather than promoting de novo protein synthesis.
JNK and p38 MAPK but not ERK signalling pathways mediate Aplidin™-induced apoptosis
To examine the role of JNK, p38 MAPK and ERK in Aplidin™ induced apoptosis, we tested the effect of the MAPK pathways inhibitors curcumin (JNK inhibitor), SB203580 (p38 inhibitor) and PD98059 (MEK inhibitor). After adjusting inhibitor concentrations to assure target specificity (not shown), inhibition of JNK with 30 μM curcumin and p38 MAPK with 30 μM SB203580, showed a significant reduction in the percentage of the apoptotic population as determined by chromatin staining (Figure 5, left) and flow cytometry (Figure 5, right). In contrast, preincubation of cells with 30 μM PD98059 resulted in no detectable protective effect. These results clearly demonstrated that JNK and p38 MAPK activation mediate the cytotoxic action of Aplidin™ in HeLa tumour cells. Given that MAPK pathways are known to be modulated by the intracellular oxidative status, the effect of different anti-oxidants in the activation of these MAPKs and subsequent apoptosis was examined. In line with its effects on apoptosis, 40 μM Ebselen, but not other antioxidants, efficiently block Aplidin™-mediated JNK activation, although it had only a slight effect on p38 MAPK (Figure 6).
Role of caspase cascade on Aplidin™-mediated apoptosis
JNK is known to phosphorylate and inactivate, among others, the anti-apoptotic Bcl-2 protein (Figueroa-Masot et al., 2001), promoting the release of mitochondrial pro-apoptotic factors, such as cytochrome c, Apaf-1 and caspase-9 and subsequent activation of the caspase cascade. Cytochrome c, detected by Western blotting, was found to leak rapidly (5 min) into the cytoplasm of Aplidin™-treated HeLa cells (Figure 7a), suggesting that the drug may induce apoptosis via JNK-mediated mitochondrial stress. In addition, Aplidin™ activation of both the upstream pro-caspase 9 and the downstream effector pro-caspase 3 (shown using pro-caspase antibodies) is also consistent with the mitochondrial apoptotic pathway. Progressive cleavage of the caspase-3 endogenous substrate PARP into an 85 kDa inactive fragment was also observed in Aplidin™ treated HeLa cells and both PARP cleavage and pro-caspases 3 and 9 activation can be inhibited by pre-treatment with the permeable broad spectrum caspase inhibitor z-VAD-fmk. MAPKs were not affected by caspases inhibition, suggesting a role upstream in the signalling cascade (Figure 7b). By contrast, caspase-8 activation was not detected (not shown). Also, preincubation of HeLa cells with 30 μM z-VAD-fmk results in complete blocking of Aplidin™-mediated apoptosis as monitored by chromatin staining with Hoechst 33342 (Figure 7c, right) and flow cytometric analysis (Figure 7c, left). The similarity of the cell cycle distribution of the z-VAD-fmk-pretreated cells to the control cells (low hypodiploidy) confirmed the clear correlation between effector caspase-3 activity and Aplidin™-induced cell death. In addition, Ebselen, which prevented JNK activation and cell death, also blocked Aplidin™-mediated caspase-3 activation (Figure 7d). Similarly, both activation of caspases-9 and -3 and PARP cleavage to the 85 kDa inactive proteolytic fragment can be prevented by preincubation of the cells with SB203580 or curcumin, but not PD98059 (Figure 7e). Together, these results clearly demonstrate that upstream oxidative events and JNK and p38 MAPK activities are mediating the caspase cascade activation and subsequent Aplidin™ mediated apoptosis.
PKC-delta plays an essential role for Aplidin™-induction of apoptosis
Aplidin™-induced cell death can be significantly prevented by inhibition of protein kinase C (PKC) activity. Pre-treatment with either the general PKC inhibitor 10 μM GF-109203X or 100 nM TPA, which leads to PKC down regulation upon long-term exposure, reduced the accumulation of cells in sub-G1 phase maintaining a uniform nuclear staining (Figure 8a). These results link Aplidin™-induced apoptosis with the PKC signalling pathway. These PKC inhibitors had no effect on Aplidin™-mediated JNK and p38 MAPK activation as monitored by Western blotting, suggesting that PKC activity acts downstream of these MAPKs in the apoptotic pathway (Figure 8b).
For the different members of the PKC family studied (α, ζ, λ, δ and μ), Aplidin™ exposure specifically results in concomitant cleavage and activation of PKC-δ as monitored by Western blotting using antibodies against total and phosphorylated PKC-δ as indicated. Phosphorylated PKC-δ on Thr505 appears as a 40-kDa proteolytic product that became apparent around 2 h after Aplidin™ treatment, remaining activated throughout the complete apoptotic process (Figure 8c). Phosphorylation of cleaved PKC-δ at Ser643 was observed and followed the very same pattern above described (not shown).
In order to evaluate the role of PKC-δ in the apoptotic process induced by Aplidin™, we used the PKC-δ inhibitor rottlerin (Gschwend et al., 1994). As shown in Figure 9a, pretreatment with rottlerin efficiently prevented, in a dose-dependent manner, Aplidin™-induced cell death, as monitored by chromatin integrity and flow cytometry. At the molecular level, rottlerin completely blocked cleavage and phosphorylation of PKC-δ as shown by Western blotting. In addition, 10 μM GF-109203X, 100 nM TPA as well as the caspase inhibitor z-VAD-fmk (30 μM) prevented PKC-δ phosphorylation, although GF-109203X did not completely inhibit its proteolytic cleavage as revealed by the presence of a non-phosphorylated 40-kDa fragment. We also examined the effect of PKC-δ inhibition on Aplidin™-mediated caspase cascade activation. Interestingly, pretreatment with rottlerin, GF-109203X and TPA resulted in complete prevention of pro-caspases -9 and -3 activation and PARP cleavage; z-VAD-fmk was used as positive control (Figure 9b).
To further establish the role of PKC-δ in mediating the cytotoxic effect of Aplidin™, we tested a primary culture of murine fibroblasts derived from PKC-δ−/− mice and their wild type counterpart as control. As shown in Figure 10a, no PKC-δ protein was detected in PKC-δ−/− fibroblasts as determined by Western blotting. Treatment of wild type fibroblasts with 0.4 μM Aplidin™ alone for 24 h resulted in a final 10% cell survival with respect to control, untreated cells. Pretreatment of wild type cells with 20 μM rottlerin partially prevented Aplidin™-induced cell death, increasing cell survival up to 26%. Upon Aplidin™ treatment, PKC-δ−/− cells exhibited, compared to wild type cells, significantly higher percentage of cell survival of around 28%. Interestingly, rottlerin pretreatment showed no remarkable protective effect on PKC-δ−/− cells, resulting in a modest increase in cell survival to 31% (Figure 10b). These results clearly associate PKC-δ status and Aplidin™ effects on cell survival.
Although Aplidin™ is effective in vivo against a variety of human cancers, the molecular mechanisms underlying its cytotoxic effects remain largely unknown. Herein, we have explored the intracellular pathways involved in Aplidin™-induced tumour cell death. Our results suggest that Aplidin™ triggers a characteristic apoptotic response without previous cell cycle arrest. We also show that increasing the recovery of the cellular radical-scavenging systems results in complete abrogation of Aplidin™ cytotoxicity, suggesting that it is mediated by early oxidative stress. In line with this, many tumours and multidrug-resistant tumour cells express high levels of GSH and GSH-S-transferase to bypass the cytotoxic effects of metabolic oxidation (Miyanishi et al., 2001; Whelan et al., 1989). Cellular oxidative stress plays an important role in regulating activation of JNK and p38 MAPK (Koji et al., 1999; Luo et al., 1998). In the present study, we show that JNK and p38 MAPK are rapidly activated by Aplidin™, resulting in sustained activation during the apoptotic process. Activation of both kinases is necessary for drug-mediated cytotoxicity, thereby confirming their pivotal role in the suicide program. A similar long lasting stimulation of JNK and p38 MAPK have been described for different forms of stress-induced apoptosis (Bacus et al., 2001; Chen et al., 1996a, 1999; Herr et al., 1997; Luo et al., 1998; Xia et al., 1995), and represent an important component of the cellular response to several structurally and functionally distinct anticancer drugs. It has also been proposed that the timing of JNK and p38 MAPK activation may be of crucial importance, with persistent rather than transient activation being associated with apoptosis (Brenner et al., 1997; Chen et al., 1996a,b). We show here that ebselen, a peroxide scavenger that acts as a mimetic of glutathione peroxidase, not only exerts a potent protective effect against Aplidin™-induced apoptotic death but also completely blocks JNK activation, demonstrating a link between oxidative imbalance, JNK activation and apoptosis. Unpredictably, ebselen had only a slight effect on p38 MAPK activation, suggesting that p38 MAPK may be acting, at least in part, in a parallel pathway from that of JNK.
In several systems it is known that JNK inactivates the anti-apoptotic Bcl-2 protein, inducing the release of apoptotic factors (Herr and Debatin, 2001). Our results indicate that Aplidin™ induces mitochondrial cytochrome-c release, procaspases-9 and -3 activation, and cleavage of the endogenous caspase-3 substrate PARP, events associated with drug-mediated cell death. In addition, we demonstrate that caspase cascade activation is dependent on JNK and p38 MAPK pathways. Aplidin™ also induces ERK phosphorylation, showing a biphasic pattern of activation. It is generally assumed that transient versus delayed ERK phosphorylation determines the proliferative or differentiating outcome, respectively (Marshall, 1995; Seger and Krebs, 1995). Biphasic ERK phosphorylation has been described for diverse stimuli (Cowan et al., 2000; Hisamoto et al., 2001; Reusch et al., 2001), including apoptosis mediated by cytotoxic drugs (Ding and Templeton, 2000). In the case of taxol or vitamin E, ERK activation is essential for drug cytotoxicity (Bacus et al., 2001; Yu et al., 2001) whilst for cisplatin, induction of ERK, although not essential, serves a putative protective role against the cytotoxic effects of the drug (Persons et al., 1999). This could be the case of Aplidin™ where ERK activation most probably exerts a protective effect against the cytotoxic effect of the drug.
We show that inhibition of PKC activity prevents Aplidin™-mediated apoptosis, indicating that PKC acts as a pro-apoptotic signal, mediating the cytotoxic action of the drug. Pro-apoptotic roles for PKC have been described for several cellular systems and stimuli (Hay et al., 2001; Powell et al., 1996; Zhao et al., 1997). Specifically, PKC-δ is shown to be involved in executing the apoptotic process induced by Aplidin™ as demonstrated by the use of the specific PKC-δ inhibitor rottlerin, which significantly blocks the cytotoxic effect of the drug. PKC-δ is concomitantly cleaved and phosphorylated during Aplidin™-induced apoptosis, giving rise to a phosphorylated (Thr505 and Ser643) proteolytic fragment of 40 kDa, most probably corresponding to the catalytic domain (Gschwendt, 1999). Given the close parallelism between cleavage and phosphorylation, and taking into account that PKC-δ is autophosphorylated in vitro on Ser643 (Stempka et al., 1998), PKC-δ could undergo autophosphorylation in vivo on these residues upon proteolysis. Pretreatment with both rottlerin and GF-109203X prior to the addition of Aplidin™, inhibit partially the accumulation of the processed 40-kDa PKC-δ fragment and completely its phosphorylation, suggesting that the kinase activity of PKC-δ is somehow necessary for its proteolytic cleavage and activation. TPA shows much more effectiveness in the inhibition of both processes, cleavage and phosphorylation, most probably acting by sequestering PKC-δ in a different subcellular compartment, thus preventing the accessibility to the activation machinery. PKC-δ activation occurs late in the apoptotic process, downstream of JNK and p38 MAPK and in a caspase-3 dependent manner, suggesting an effector, rather than an initiator, role in mediating Aplidin™-induced apoptosis. Several studies have reported the involvement of PKC-δ in cells induced to undergo apoptosis by anti-tumoural drugs (Chen et al., 1999; Hofmann, 2001) as well as other different pro-apoptotic stimuli (Krupinski et al., 2000; Reyland et al., 1999; Shao et al., 1997). Furthermore, expression of the PKC-δ catalytic domain in several cell types induces phenotypic changes indicative of apoptosis (Mizuno et al., 1997; Shao et al., 1997). Avoiding the non-verifiable side effects of the inhibitors used, here we show that cells defective in PKC-δ are more resistant to Aplidin™-induced cell death compared to their wild type counterpart. These results unequivocally point to PKC-δ having a specific action in the execution of Aplidin™-mediated cytotoxicity. Different roles for PKC-δ activity during apoptosis have been reported, such as disassembly of nuclear lamina (Cross et al., 2000), inhibition of proliferin expression (Kang et al., 2000) and inactivation of DNA protein kinase (Bharti et al., 1998) among others (Gschwendt, 1999 and references therein). In addition, it has been reported that PKC-δ activates the MEK-ERK pathway (Ueda et al., 1996), which could explain the delayed ERK phosphorylation observed during the Aplidin™-induced apoptotic process, that coincides in time with the proteolytic activation of PKC-δ.
Interestingly, we show that inhibition of PKC-δ activation with rottlerin, GF-109203X or TPA, completely blocks the activation of the caspase cascade upstream of caspase-9. This dependence of PKC-δ activity for full activation of caspase-3 have been reported in other cellular systems (Reyland et al., 1999). Although we cannot rule out the possibility that full-length PKC-δ could have some role in mediating the apoptotic pathway upstream of the caspase cascade, our results could suggest a positive feedback loop whereby caspase-mediated activation of PKC-δ results in amplification of the caspase cascade and subsequent apoptosis. Further supporting this idea, it has been published that cytochrome-c release from mitochondria upon treatment of HL-60 cells with didemnin B, a structural analogue of Aplidin™, is caspase-dependent, suggesting the existence of a large positive feedback amplification loop (Grubb et al., 2001).
Based on the above exposed data, we propose the following mechanistic model for Aplidin™-induced apoptosis (Figure 11). (1) Generation of early oxidative imbalance; (2) Activation of stress kinases JNK and p38 MAPK (and ERK); (3) Possible activation of full-length PKC-δ; (4) Mitochondrial dysfunction and cytochrome c release; (5) Activation of caspase cascade; (6) Target of PKC-δ to be activated by caspases (gain of catalytic competence); (7) Caspase-dependent cleavage of specific substrates such as PARP and PKC-δ; (8) PKC-δ activation (autophosphorylation?); (9) PKC-δ-dependent amplification of the apoptotic machinery.
It would be of great interest to understand the mechanisms that prime PKC-δ to be activated late in the apoptotic process. Since no changes in PKC-δ expression levels are detected in response to Aplidin™, it is conceivable that PKC-δ could undergo some kind of post-translational modification that target it to be processed and activated by caspases. PKC-δ is known to be functionally modulated by tyrosine phosphorylation that might regulate the specificity of the kinase toward a given substrate. Several tyrosine kinases have been reported to phosphorylate PKC-δ such as Src and Abl, although its significance in vivo is not yet unequivocally identified (Gschwendt, 1999 and references therein). Future work in this direction will help to clarify the signalling pathways leading to PKC delta targeting and activation.
Also, although there is good evidence indicating that modulation of PKC-δ may be a good target for inhibiting tumour cell survival (Hofmann, 2001), a more thorough understanding of the potential of activated PKC-δ in antitumour therapy is needed.
Materials and methods
Reagents and antibodies
Aplidin™ was manufactured by PharmaMar, S.A. All other reagents used in this study were of molecular biology grade. Hoechst-33342, propidium iodide, curcumin, PD098059, SB203580, TPA, GF-109203X (bisindolymaleimide I), DPI, tiron, NAC, ebselen, catalase, glutathione and vitamin E were purchased from Sigma. z-VAD-fmk and rottlerin were purchased from Calbiochem.
Anti-JNK1 (sc-474), anti-ERK2 (sc-154), anti-p38 MAPK (sc-535), anti-PARP (sc-7150), anti-(pro)caspase-3 (sc-7148) and anti-nPKCδ (sc-937) polyclonal antibodies were purchased from Santa Cruz Biotechnology. Anti-phospho-p38 (#9211), anti-phospho-JNK (#9251), anti-phospho-PKCδ (Thr550) (#9374), anti-phospho-PKCδ (ser643) (#9376) (polyclonal) and anti-phospho-ERK (#9106) (monoclonal) antibodies were purchased from Cell Signaling Technologies, Inc. Anti-cytochrome c antibody was purchased from Pharmingen-BD (#556433).
HeLa cells (ATCC# CCL-2) were maintained in DMEM supplemented with 10% FCS and 100 units/ml penicillin and streptomycin at 37°C and 5% CO2. Primary cultures of fibroblasts deficient in PKC-δ (derived from mice homozygous for the null mutation in the PKC-δ gene (PKC-δ−/−) (Miyamoto et al., 2002)) were maintained in DMEM supplemented with 10% FCS, 100 units/ml penicillin and streptomycin, 0.1 mM non-essential aminoacids and 1 mM sodium pyruvate. Nuclei were stained with the permeable bis-benzimide Hoechst-33342 (1 μg/ml in PBS) and examined by fluorescence microscopy.
Determination of cell survival
Cells plated in 35 mm petri dishes were treated with vehicle or 0.4 μM Aplidin™ for different time periods as indicated in the text. Cells were washed twice with PBS, fixed for 10 min in 1% glutaraldehyde solution, rinsed twice in PBS, and stained in 0.2% cressyl violet solution for 15 min. Cells were then rinsed several times with distilled water and air-dried. Cressyl violet dye was extracted with 10% acetic acid solution and cell density indirectly estimated with a spectophotometer by measuring absorbance at 595 nm. Cell survival was expressed as percentage of control cells.
Analysis of DNA fragmentation
Fragmentation of DNA was assessed as previously described (Sánchez et al., 1996). Briefly, cells (3×106) were incubated in the absence or presence of 0.4 μM Aplidin™ for the times indicated, washed with PBS, scraped and pelleted at 4°C. Cells were lysed in 500 μl of 2.5 mM Tris-HCl (pH 8.0), 10 mM EDTA, and 0.25% Triton X-100 and stored at 4°C for 15 min. Intact nuclei were eliminated by centrifugation (500 g for 15 min) and DNA in the supernatant was precipitated, resuspended in 200 μl of TE buffer, and sequentially incubated at 37°C for 30 min with RNAse A (0.1 mg/ml) and for 2 h with proteinase K (0.25 mg/ml). Purified DNA was resolved in 1.5% agarose-TAE gels.
Analysis of nuclear DNA content by flow cytometry
The ploidy determination of nuclei was estimated by flow cytometry DNA analysis. Cells were washed in PBS, fixed by drop wise addition of 1 ml of cold 70% ethanol and allowed to stand on ice for 10 min. Cell pellet was resuspended in 200 μl of PBS containing 10 μg/ml RNase A and incubated at 37°C for 30 min. Propidium iodide (200 μg/ml) was added and the DNA content per nucleus was evaluated in a FACScan flow cytometer (Becton-Dickinson). For analysis, only signals from single nuclei were considered (104 nuclei/assay).
Cells were washed in PBS, collected and resuspended in lysis buffer (20 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1% (v/v) Nonidet P-40, 2 mM EDTA, 1 mM PMSF, 10 μg/ml aprotinin, and 10 μg/ml leupeptin) and kept on ice for 15 min. Cell extracts were cleared by microcentrifugation at 14 000 g for 30 min at 4°C. For cytochrome c detection, cells were harvested by centrifugation, washed in PBS and gently lysed for 30 s in 50 μl of ice-cold buffer containing 250 mM sucrose, 1 mM EDTA, 0.05% digitonin, 25 mM Tris (pH 6.8), 1 mM DTT, 1 μg/ml (each) leupeptin, aprotinin and pepstatin, 1 mM benzimidine and 0.1 mM PMSF. Lysates were centrifuged at 12 000 g at 4°C for 2 min to obtain the cytosolic fraction free of mitochondria (supernatant) and mitochondrial fraction (pellet).
Equal amount of extracts were resolved in SDS–PAGE and electroblotted to activated PVDF membranes (Immobilon-P, Millipore) following standard techniques (Sambrook et al., 1989). Membranes were sequentially probed with primary and appropriate secondary (horseradish-peroxidase-conjugated) antibodies following the manufacturer's instructions. Antibody-antigen complexes were detected using the ECL system (Amersham Pharmacia Biotech).
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We would like to thank Juan M López-Oliva for excellent technical assistance and members of the R+D Department of PharmaMar, S.A. We also thank Drs Simon Munt and José Antonio López-Martín for critical reading of the manuscript and fruitful suggestions.
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