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Abramson Family Cancer Center, University of Pennsylvania, Philadelphia, PA 19104
Received December 26, 2007; accepted April 23, 2008
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Abstract |
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) and is recognized as a contributing factor to resistance of cancer to chemotherapy, including alkylating agents, platinum compounds, and radiation (Vaupel et al., 1989). Many of the adaptive responses set in motion by hypoxic exposure are regulated by hypoxia-inducible factor-1 (HIF-1), an oxygen-sensitive transcription factor that induces expression of genes involved in angiogenesis, proliferation, and metastasis (Semenza, 2002). Other transcription factors, including activator protein-1 (AP-1), are also activated by hypoxia and contribute to the altered gene expression profile of hypoxic cells (Cummins and Taylor, 2005).
Oxaliplatin (L-OHP), a third-generation platinum compound, has a major role in the treatment of colorectal cancer (André et al., 2004; Goldberg et al., 2006). The induction of oxaliplatin resistance by hypoxia has been described in testicular cancer cell lines (Koch et al., 2003) and in a series of hypoxia-conditioned HT29 colon cancer cell lines (Yao et al., 2005a). In previous work, we have demonstrated induction of AP-1 family genes and increased AP-1 transcriptional activity in hypoxic HT29 cells (O'Dwyer et al., 1994; Yao et al., 1994). We have also demonstrated the induction of AP-1 transcriptional activity by oxaliplatin in a panel of colon cancer cell lines (Rakitina et al., 2003). Hence, these pathways may be relevant to tumor responses under hypoxic conditions.
The AP-1 transcription factor consists of homo- and heterodimers of the Jun, Fos, ATF, and MAF protein families that stimulate the expression of a broad range of genes involved in proliferation, differentiation, and apoptosis (Eferl and Wagner, 2003). Activation of AP-1 by various signals, including hypoxia, occurs primarily through parallel three-tier signaling cascades terminating in a group of mitogen-activated protein kinases (MAPKs) (Dérijard et al., 1995). MAP kinases in turn are activated by upstream regulators-MAPK kinases (MEKs or MKKs). MEK1 and MEK2 activate extracellular signal-regulated kinases (ERK1/2) mostly in response to mitogenic stimuli (Zheng and Guan, 1993). MKK3 and MKK6 are responsible for activation of p38 MAPK (Enslen et al., 1998). MKK4 (also known as SEK1) and MKK7 both activate Jun NH2-terminal kinase (JNK). Studies in vitro have shown that MKK4 can also phosphorylate p38 MAPK, whereas MKK7 is specific toward kinases of JNK subgroup (Davis, 2000; Tournier et al., 2001). Activation of MEKs involves multiple kinases (MEKKs or MKKKs), acting on different substrates dependent upon particular extra- or intracellular signals (Fanger et al., 1997). Data on oxaliplatin-induced signaling through MAP kinases and their effects on cytotoxicity of the drug are scarce. Our previous results suggest that both JNK and p38 MAPK contribute to oxaliplatin-induced cell death in colon cancer cell lines (Rakitina et al., 2003).
We previously found that short-term hypoxic exposure activates SEK1 and JNK, leading to up-regulation of c-Jun in HT29 cells (Vasilevskaya and O'Dwyer, 1999). Now we show that JNK activation results from hypoxia-induced signaling through both SEK1 and MKK7, whereas MEK1/2 and MKK3/6 remain constitutively active. In a functional analysis of this induced signaling, we isolated HT29-derived cell lines stably expressing control vector (HTpc3) or dominant-negative constructs for SEK1 or MKK7 (HTS13 and HTM9, respectively). We found that signaling through MKK7/JNK/c-Jun cascade is the major mechanism for hypoxic activation of AP-1 in HT29 cells. We also show that modulation of these pathways affects the sensitivity of hypoxic HT29 cells to oxaliplatin: the inhibition of MKK7 leads to an increase in oxaliplatin resistance upon hypoxic exposure, whereas inhibition of SEK1 renders cells more sensitive under these circumstances. Confirmation of the potential biological relevance of this observation was obtained in in vivo experiments using xenografts of stably transfected cell lines. These data implicate differential effects of signaling through MKK7 and SEK1 in resistance and sensitivity to oxaliplatin in hypoxic tumor cells (i.e., that hypoxic signaling to SEK1 contributes to oxaliplatin resistance, thus identifying SEK1 as a potential target for reversal of hypoxic resistance). Conversely, absence of MKK7 has the potential to render a tumor resistant to agents that require hypoxia signaling to exert their therapeutic effects. These results may further help to address some of the conflicting data regarding the role of signaling through JNK to influence apoptosis (Vasilevskaya and O'Dwyer, 2003).
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Materials and Methods |
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Plasmids and Isolation of Stably Transfected Cell Lines. The FLAG-tagged dominant-negative mutant of MKK7 [pcDNA3-Flag-MKK7 (ala)] was kindly provided by Dr. Roger Davis (University of Massachusetts Medical Center, Worcester, MA); the HA-tagged dominant-negative mutant of SEK1, (HA-SEK-AL/pcDNA3.1) was a gift from Dr. Jim Woodgett (Ontario Cancer Institute, Toronto, ON, Canada); HA-tagged dominant-negative mutants of JNK1 and JNK2 (HA-JNK1-APF and HA-JNK2-APF), cloned into LNCX vector, were kindly provided by Dr. Tomas Berl (University of Colorado, Boulder, CO). To isolate cell lines stably expressing empty vector, dnSEK1 and dnMKK7, transfectants were cultivated at low density in media containing G418 (0.75 mg/ml; Invitrogen); surviving colonies were isolated using glass cylinders, propagated, and assessed for expression of tag protein.
Hypoxic Treatment. Exposure of cells to acute hypoxia was achieved by incubation in anaerobic chamber (Forma Scientific, Inc., Marietta, OH) filled with gas mixture consisting of 5% CO2, 9%H2, and 86% N2. Oxygen content (below 0.5%) was monitored by PROOX 110 oxygen sensor (BioSpherix, Redfield, NY). Cells were plated in 100-mm glass Petri dishes to a density of 2 x 106 cells per dish and subjected to hypoxia within 36 h when the cells had reached 70% confluence. Inhibitors at concentrations of 10 µM were added 2 h before hypoxia treatment; 0.01% DMSO was added to control samples. The cells were harvested at various time points for further isolation of protein extracts.
Protein Extract Preparation. Total protein extracts were prepared as follows: after hypoxia, cells were washed twice with phosphate-buffered saline and lysed inside the chamber in cell lysis buffer (Cell Signaling Technology, Danvers, MA), supplemented with Complete protease inhibitor cocktail (Roche Applied Science, Indianapolis, IN) and 1 mM phenylmethylsulfonyl fluoride (Sigma, St. Louis, MO). The contents of scraped dishes were transferred into microcentrifuge tubes, taken out of hypoxia chamber, and placed in a shaker for 30 min at 4°C. Lysates were then centrifuged for 10 min at 10,000 rpm (4°C) and the protein concentration of cleared cellular extracts was measured using the Bio-Rad Protein Assay (Bio-Rad Laboratories, Hercules, CA).
Western Blotting. For protein electrophoresis in SDS-polyacrylamide gels, protein extracts were used in amounts of 10 µg per lane. Western blotting was carried out according to standard procedures using horseradish peroxidase-conjugated secondary antibodies purchased from Santa Cruz Biotechnology (Santa Cruz, CA) and the ECL+Plus detection system (GE Healthcare, Chalfont St. Giles, Buckinghamshire, UK). The primary antibodies used were as follows: antibodies against MKK7 and phosphorylated forms of JNK, p38, SEK1, and MEK1/2 from Cell Signaling Technology; antibody against phospho-MKK7 from Millipore (Billerica, MA) and c-Fos antibody from Calbiochem (San Diego, CA). The remaining antibodies were purchased from Santa Cruz Biotechnology.
Transfections and Luciferase Reporter Assays. Control and reporter, pTA-Luc and pAP1-Luc, plasmids were from the PathDetect AP-1 Cis-Reporting system (Stratagene, La Jolla, CA). Cells were plated in six-well plates at the density of 1.5 x 105 per well and after 24 h were transfected using FuGENE 6 transfection reagent (Roche) with the mixture of dominant-negative MAPKK or JNK construct (1 µg), control or reporter plasmid (1 µg) and pRL-CMV vector, expressing Renilla reniformis luciferase (40 ng; Promega), unless otherwise noted. Antisense c-jun oligonucleotides and scrambled control were used at a concentration of 100 µM. Twenty-four hours after transfection, cells were subjected to hypoxia for 8 or 24 h, collected, and assayed for luciferase activity using the Dual-Luciferase Reporter Assay System (Promega), according to the manufacturer's protocol. Luciferase activity was measured by TD20/20 tube luminometer (Turner Designs, Sunnyvale, CA). The relative light units were calculated as ratio of firefly luciferase to R. reniformis luciferase luminescence. Nonspecific luciferase activation determined in parallel experiments with control vector was subtracted. All experiments were performed at least twice in duplicate.
Colony-Forming Assays. For clonogenic assays, HTpc3, HTM9, and HTS13 cells were plated in six-well plates at a density of 300 cells per well; after 24 h, 1 µM oxaliplatin was added immediately before transfer of the plates to the hypoxia chamber. After 24 h of hypoxia, plates were returned to normoxic conditions and, after the addition of fresh media, cultivated for 10 to 14 days; colonies were then fixed in 75% ethanol, stained with Coomassie Blue (Sigma, St. Louis, MO), and counted manually. All experiments were performed at least three times in duplicate.
Retroviral Delivery of dnSEK1 Construct into Colon Cancer Cell Lines. The DNA fragment coding for dnSEK1 from HA-SEK-AL/pcDNA3.1 plasmid was cloned into the LPNX plasmid (Clontech Laboratories Inc., Mountain View, CA). The resulting vector (or empty LPNX as a control) was transfected into the PhoenixAmpho packaging cell line, kindly provided by Dr. Anil Rustgi from the University of Pennsylvania, using FuGENE 6 reagent (Roche). Media was exchanged 24 h later and incubation continued further. Supernatants containing control or recombinant virus were collected at 48, 72, and 96 h after transfection, combined, filtered through 0.45-µm sterile filters, and stored at -80°C. Colon cancer cells plated into six-well plates were infected in the presence of polybrene (8 µg/ml; Millipore Bioscience Research Reagents, Temecula, CA); 48 h later, selective media (1 µg/ml puromycin; Sigma) was added. Surviving cells were pooled, evaluated for expression of HA-tag and used in MTT assays to assess oxaliplatin sensitivity under hypoxic conditions.
In Vivo Experiments. To assess the efficacy of oxaliplatin treatment with or without bevacizumab (anti-vascular endothelial growth factor antibody) in HTpc3-, HTS13-, and HTM9-derived tumors, 6- to 8-week old female mice (C.B.17 SCID) were used. To produce tumors, control, HTS13, and HTM9 cells (2 x 106 cells) were injected subcutaneously into the left flank of the mice. When the tumors reached approximately 100 mm3, mice were divided into four groups (six mice per group) for treatment with bevacizumab (Genentech, Inc., San Francisco, CA), oxaliplatin (Eloxatin, Sanofi Aventis, Bridgewater, NJ), the combination of bevacizumab and oxaliplatin, and a vehicle control group. Mice in the bevacizumab treatment group received 5 mg/kg bevacizumab by intraperitoneal injection every 3 days for 30 days. The oxaliplatin treatment group was injected intraperitoneally with 10 mg/kg oxaliplatin per week for 2 weeks. The combination treatment group received bevacizumab (every 3 days, 5 mg/kg for 30 days) and oxaliplatin (weekly for 2 weeks, 10 mg/kg). The control group received saline intraperitoneally every 3 days. Tumor volume and body weight were measured every 3 days. Tumor volume was calculated using the formula V = AB2/2, where A is the largest diameter and B is the smallest diameter. Tumor growth delay was calculated as the difference in the time for control and treated tumors to grow from 100 to 600 mm3. All animal experiments were done according to an IACUC (Institutional Animal Care and Use Committee) protocol approved by the University of Pennsylvania.
Statistical Analysis. Data were analyzed with unpaired Student's test: P < 0.05 was accepted as a statistically significant difference compared with corresponding control.
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Results |
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; Vasilevskaya and O'Dwyer, 1999). The initial approach to identify the MAP kinase signaling pathways involved using pharmacological inhibitors. Concentrations of inhibitors were selected in preliminary model experiments, using known activators of respective pathways (for example, anisomycin to activate JNK pathway, sorbitol for activation of p38MAPK pathway). Chosen concentrations of inhibitors were sufficient to cause detectable responses in signaling (based on Western analysis of the downstream targets), after relatively short exposure (1-3 h). HT29 cells were pretreated for 2 h with inhibitors of ERK (U0126), p38MAPK (SB203580), and JNK (SP600125), all at 10 µM, then subjected to hypoxia for up to 5 h. Total protein extracts were used to assess the activation status of MAPK pathways and the content of AP-1 proteins. 17-AAG (1 µM; 90-kDa heat shock protein inhibitor) was also used to inhibit both ERK and JNK signaling (Vasilevskaya and O'Dwyer, 1999). We found that inhibition of the MEK1/ERK cascade by either U0126 or 17-AAG did not alter c-Fos levels after 5 h of hypoxia and, consequently, had no likely impact on its role in the c-Fos/c-Jun dimer (Fig. 1A). There was no inhibition of activating transcription factor 2 (ATF2) phosphorylation in the presence of SB203580 (Fig. 1B). This inhibitor does not affect phosphorylation of p38 MAPK itself by upstream kinases (Kumar et al., 1999), but its efficacy in our system is supported by hypophosphorylation of cAMP response element-binding protein and of MAPK-activated protein kinase-2, known down-stream targets (Fig. 1B). In contrast, phosphorylation of both c-Jun and ATF2 was inhibited in the presence of SP600125, confirming our previous results (Vasilevskaya and O'Dwyer, 1999) that the activation of these AP-1 proteins in HT29 during hypoxic exposure is dependent mostly on JNK, not p38 MAPK. We also have consistently observed robust phosphorylation of JNK during hypoxia in the presence of SB203580. Activation of JNK upon inhibition of p38 MAPK was described for macrophages (Hall and Davis, 2002) and in our case could also contribute to the active status of ATF2. These data demonstrate that both induction of c-Jun expression (through activation of c-Jun/ATF2) and its activation during hypoxia in HT29 depend predominantly on functional JNK.
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MKK7 Is a Key up-Stream Mediator of Hypoxia Signaling to AP-1 in HT29 Cells. We previously found that SEK1 was activated in HT29 cells subjected to hypoxia for 3 and 6 h, whereas MEK1/2 and MKK3/6 were constitutively active and not further induced (Vasilevskaya and O'Dwyer, 1999). To further dissect hypoxia signaling to c-Jun/AP-1, we examined the role of another up-stream regulatory kinase— MKK7. We subjected HT29 cells to hypoxia for various times (ranging from 30 min to 5 h), collected protein extracts, and assessed activation of MKKs by Western blot analysis. Hypoxia activates both MKK7 and SEK1, with a marked increase in phosphorylation by 3 h, whereas changes in MKK3/6 and MEK1/2 were minimal (Fig. 2A). We therefore focused our further study on the (SEK1/MKK7)/JNK/c-Jun cascade. To assess the effect of modulating the pathway on AP-1 transcriptional activity we carried out luciferase reporter assays: HT29 cells were cotransfected with a reporter plasmid (pAP1-Luc) and plasmid constructs expressing dominant-negative forms of SEK1, MKK7, or JNK1/2. We also used c-jun antisense oligonucleotides (along with corresponding scrambled controls) as a negative control for AP-1 activation. Results of AP-1-Luc assays performed after eight hours of hypoxia (Fig. 2B) demonstrate that although both SEK1 and MKK7 are important for activation of AP-1 by hypoxia, inhibition of MKK7 (64% decrease) affects activation more than inhibition of SEK1 (43% decrease), whereas inhibition of JNKs resulted in 70% decrease in AP-1 activation by hypoxia. These data suggest that the importance of functional MKK7 for the maintenance of basal AP-1 activity and its activation by hypoxia in HT29 cells is comparable with that of JNK. We have to note, however, that because transformation efficiency in HT29 cells is low, it would be appropriate to consider these results more qualitatively than quantitatively. Accordingly, to study the roles of SEK1 and MKK7 further, we opted to isolate cell lines stably expressing the dominant-negative forms of these kinases.
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Differential Effects of Hypoxia on AP-1 Activation in Cell Lines Stably Expressing Dominant-Negative MKKs Confirms Major Role for MKK7. Derivatives of the HT29 cell line stably expressing empty vector (HTpc3), dnSEK1 (HTS13), and dnMKK7 (HTM9) were isolated as described under Materials and Methods. The expression of dominant-negative mutants was confirmed by Western blot analyses for tag-proteins: HA for dnSEK1 and FLAG for dnMKK7 (Fig. 3A, left and right, respectively). When the derivative cell lines were subjected to hypoxia, profound inhibitory effects of dnMKK7 on both JNK and c-Jun phosphorylation were observed (Fig. 3B), whereas the inhibition of SEK1 was less effective, consistent with the results above (Fig. 2B). Again, because the hypoxic activation of ATF2 in HT29 cells depends primarily on JNK, phosphorylation of ATF2 is altered in a manner similar to that of JNK. We then assessed activation of AP-1 in these cell lines using reporter gene assays during prolonged hypoxia: cells were cotransfected with 2.5 µg of control or reporter pAP1-Luc plasmid and pRL-CMV vector, and 24 h after transfection subjected to hypoxia for another 20 h. The importance of SEK1 and MKK7 signaling is shown in the lower levels of AP-1 transcription even under normoxic conditions. With hypoxia, however, significant hypoxic induction of AP1 was observed in SEK1-deficient cells, whereas in MKK7-deficient cell line it was completely abolished (Fig. 3C).
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Signaling through MKK7 Enhances Oxaliplatin Cytotoxicity in Hypoxic HT29 Cells, whereas SEK1 Mediates Protective Responses. We have reported recently that hypoxia-induced transcriptional changes are a factor in the cellular responses of colon cancer cell lines to cisplatin and oxaliplatin (Yao et al., 2005a,b). The importance of the AP-1 transcription factor in cellular resistance to platinum drugs has also been described in various cellular models (Bonovich et al., 2002; Piret et al., 2006). Therefore, we went on to determine whether inhibition of AP-1 signaling through down-regulation of SEK1 or MKK7 could affect cell growth and survival during hypoxia and/or treatment with oxaliplatin. We conducted MTT assays to establish the IC50 for ox aliplatin in hypoxic HT29 cells: oxaliplatin was added immediately before placing the cells into hypoxia chamber for 24 h, followed by cultivation in normoxic conditions for the rest of the 72 h exposure to oxaliplatin. We found differences in oxaliplatin sensitivity of HTpc3, HTS13, and HTM9 cell lines even under normoxic conditions, with IC50 of 0.6, 0.3, and 0.9 µM, respectively (Table 1). Hypoxia alone did not significantly affect the proliferation rate of cell lines: all three lines demonstrated MTT readings comparable with those obtained from cells cultivated in normoxic conditions (data not shown). Differences in drug sensitivity between the lines were more pronounced under hypoxic conditions: although oxaliplatin resistance of HTpc3 and HTS13 cell lines was increased 2-fold, inhibition of MKK7/JNK/c-Jun pathway led to much greater resistance in HTM9 cell line, with IC50 changed from 0.9 to 5 µM (Table 1). The results of MTT assays, however, are influenced by oxaliplatin effects on both survival and cell cycle regulation of HT29-derived cells. The growth inhibitory effects might be especially significant in HTM9 cells, because involvement of MKK7 in the coupling of stress signaling and cell cycle progression has been reported (Wada et al., 2004). Therefore, to establish the impact of SEK1 and MKK7 inhibition on cell survival (and also to separate these from the effects of growth inhibition reflected in the results of MTT assays), we then carried out colony-forming assays. HTpc3, HTS13, and HTM9 cell lines were treated with the range of oxaliplatin concentrations for 72 h, with or without hypoxia for first 24 h similar to previous MTT assays. The sensitivity to oxaliplatin in a panel followed the patterns of MTT assays in general: HTS13 cells exhibited the lowest IC50 concentration and HTM9 cells, the highest (data not shown). To further illustrate the differences in oxaliplatin sensitivity of HTpc3, HTS13, and HTM9 cells lines, we carried out additional colony-forming assays after exposing them to the equimolar concentration of oxaliplatin (1 µM, which falls in a medium range of clinically achievable concentrations) for 24 h with or without simultaneous hypoxic exposure (Fig. 4). Sensitivity to oxaliplatin did not vary dramatically among the cell lines when cultivated in normoxic conditions; however, when cells were subjected to both oxaliplatin treatment and hypoxia, a marked decrease in survival was observed in the HTS13 cell line, whereas HTM9 cells were most resilient. Our data suggest that MKK7-mediated activation of the JNK/c-Jun pathway is essential for cell death of HT29 cells treated with oxaliplatin under hypoxia, whereas SEK1 may be involved in the mechanisms related to oxaliplatin resistance. To determine that the effects we observed are not HT29-specific, we expanded the cellular model to incorporate multiple colon cancer cell lines with varying genetic backgrounds. We used a retrovirus-based delivery system for the introduction of dominant-negative SEK1 into these cells. The results of cytotoxicity assays in SW480 and HCT15 cell lines corroborate these in HT29, whereas BE cells were not sensitized (Table 2).
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Intact MKK7 Is Required for Therapeutic Effects of Oxaliplatin and Bevacizumab in Vivo. Finally, we studied the effects of SEK1 and MKK7 inhibition in a mouse xenograft model. SCID mice were injected subcutaneously with 2 x 106 HTpc3, HTS13, or HTM9 cells and subjected to treatments with bevacizumab and oxaliplatin alone, or in combination. The doses/schedules of the in vivo experiments were selected based on our own previous work (Selvakumaran et al., 2008) and also from other published works in mice models of colon cancer that are known to work in those doses/schedules. Both the anti-vascular endothelial growth factor and oxaliplatin treatments demonstrated growth inhibitory effects on control HTpc3 tumors (Fig. 5, A and B), the combination therapy being the most effective. Figure 5A demonstrates that inhibitory effects of the therapies on the growth of HTM9-derived tumors were modest and differed significantly from control tumors treated with bevacizumab alone (P < 0.01), oxaliplatin alone (P < 0.001), or the combination (P < 0.001). On the other hand, HTS13-derived tumors grew more slowly than control tumors with all treatments (Fig. 5B). These differences in response to therapy were even more evident when we calculated tumor growth delay (Table 3), the time difference between untreated and treated tumors to grow from 100 to 600 mm3. The tumor growth delay in bevacizumab-treated mice was 2.7 and 8 days for HTM9 and HTS13 tumors, respectively, versus 5.4 days for HTpc3 tumors. In the oxaliplatin-treated group, the tumor growth delay was 2 and 10 days versus 7.5 days, respectively. Finally, with the combination treatment, the growth delay for HTM9 tumors was only 1.4 days, dramatically different from 10.9 days for the control tumors, whereas HTS13 tumors demonstrated a growth delay of 13 days. These results show that MKK7-induced activation of JNK provides a direct pro-death signal in hypoxic cells exposed to oxaliplatin, whereas SEK1 has in addition a preponderant survival influence in this context and may be a target for therapeutic intervention.
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Discussion |
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). However, the multitude of other transcription factors activated by hypoxia directly or indirectly (Cummins and Taylor, 2005) may in turn regulate a wide variety of transcriptional responses and may be important for an understanding of the biology of tumor hypoxia in general and of hypoxic resistance to cancer therapies in particular. Activation of the AP-1 transcription factor by hypoxia was reported for multiple cancer cell lines, including human colon adenocarcinoma (Yao et al., 1994; Vasilevskaya and O'Dwyer, 1999) and hepatic and ovarian carcinomas (Xu et al., 1999; Minet et al., 2001), among others. MAP kinase pathways, as major activators of AP-1-regulated transcription, might therefore be considered potential targets to influence cellular responses to hypoxia alone and in combination with chemotherapy. In this article, we have presented results detailing the input of individual MAP kinases and their upstream activators, MAPK kinases, on AP-1 induction by hypoxia in HT29 cells, as well as the effects of selective disruption of the pathways on sensitivity to hypoxia and oxaliplatin.
Our analysis of hypoxia signaling indicates that in HT29 cells, hypoxic induction of AP-1 activity can be abolished by inhibition of JNK. The MEK/ERK pathway seems to play no role in this response: 1) MEK1/2 and ERK1/2 are constitutively active in normoxic HT29 cells, and not affected by hypoxic exposure, and 2) pharmacological inhibition of this pathway by U0126 did not translate into inhibition of the expression or phosphorylation status of the major AP-1 proteins (Fig. 1, A and B). In contrast, dramatic changes occurred in the activity of SEK1 and MKK7 kinases. Hypoxia also strongly induced signaling to p38 and JNK, which have both been shown to be proapoptotic under low oxygen (Betigeri et al., 2006; Sun et al., 2006). A proapoptotic role for these kinases in cisplatin-treated cells has also been demonstrated in various cellular models (Roos and Kaina, 2006), including lung, ovarian, and human colon cancer (Brozovic et al., 2004; Vasilevskaya et al., 2004). Much less is known about the induction of signaling pathways by oxaliplatin, and the involvement of stress-regulated MAP kinases in oxaliplatin-induced cell death. We have reported significant activation of p38 MAPK and JNK by cisplatin and oxaliplatin in a panel of colon cancer cell lines (Rakitina et al., 2003; Vasilevskaya et al., 2003, 2004), and showed that inhibition of JNK and p38 MAPK by pharmacological inhibitors results in decrease of oxaliplatin cytotoxicity in colony-forming assays (Rakitina et al., 2003). Despite the proapoptotic properties of these stress-activated MAPK pathways induced by both hypoxia and platinum drugs, the efficacy of cytotoxic drugs is diminished in hypoxic conditions (Koch et al., 2003; Yao et al., 2005a). Therefore, we have tried to further delineate the role of signaling through these pathways to better understand hypoxic resistance to oxaliplatin.
The pair of kinases activating JNK, SEK1, and MKK7, although acting synergistically (Fleming et al., 2000), display several differences (Wang et al., 2007). One key difference is the ability of SEK1 to phosphorylate p38 MAPK in addition to JNK. They also show variable substrate specificity: SEK1 preferentially phosphorylates the tyrosine residue in ThrPro-Tyr motif of JNK, whereas MKK7 acts on the threonine residue. This is relevant to our results in that although both kinases are necessary for full activation of JNK (Kishimoto et al., 2003), phosphorylation on the threonine residue alone (by MKK7) has been shown to be sufficient to increase JNK activity (Tournier et al., 2001). The results of MTT assays using HT29-derived cell lines HTpc3, HTS13, and HTM9 point to differential functions of SEK1 and MKK7 in HT29 cells during hypoxia and oxaliplatin exposure. Although the survival of none of the cell lines was affected by hypoxia alone, HTM9 cells were 3-fold more resistant to oxaliplatin in normoxic conditions then HTS13 cells, confirming that JNK activation is required for oxaliplatin cytotoxicity. The dramatic difference in survival among HTM9, HTpc3, and HTS13 cell lines after hypoxia in the presence of oxaliplatin demonstrated by both MTT (Table 1) and colony-forming assays suggest that inhibition of SEK1 results in specific alterations of hypoxic signaling rendering cells more sensitive to oxaliplatin. In HT29 cells with impaired MKK7 function, inhibition of the JNK/c-Jun/AP-1 cascade prevents cell death under hypoxia/oxaliplatin treatment, whereas the function of SEK1 seems to be more complex: although the overall activation of JNK and c-Jun in HST13 cell line is lower than in parental cells, they still demonstrate higher sensitivity to all treatments, compared with other two lines. The observed increased mortality in the SEK1-deficient cell line, therefore, could not be solely attributed to the functional MKK7 present in these cells.
Hypoxic induction of p38 MAPK was unaltered in HTM9 and, more important, in HTS13 cell line (Fig. 3B), suggesting that its activation in this situation most likely does not involve SEK1. In colony-forming assays, when these cell lines were treated with hypoxia/oxaliplatin in the presence of a pharmacological inhibitor of p38 MAPK (SB203580), resistance to oxaliplatin was uniformly increased to the same degree in all cell lines, irrespectively of SEK1 and MKK7 status (data not shown). The additional importance of signaling through MKK3/6 to p38 MAPK for oxaliplatin cytotoxicity is evident therefore, and detailed studies are currently under way. Here we focus on SEK1 and MKK7 to demonstrate that their action is not redundant and leads to differential responses to oxaliplatin in cell lines deficient in either of these kinases.
Taken together, our data suggest that a survival pathway normally induced by hypoxia is not activated when SEK1 is inhibited by overexpression of a dominant-negative mutant. This was somewhat surprising in light of the identification of SEK1 (MKK4) as a tumor suppressor (Wang et al., 2007). Our results, however, are indirectly supported by studies suggesting a pro-oncogenic role for SEK1. Wang et al. (2004) used breast and pancreatic cancer cell lines with different basal levels of SEK1 to modulate kinase expression by introducing either recombinant protein or siRNA construct into cells. They showed that expression of SEK1 (MKK4) in MKK4-negative cell lines stimulated cell proliferation and invasion, whereas inhibition of SEK1 (MKK4) expression in MKK4-positive cells resulted in growth inhibition and higher sensitivity to serum deprivation and suppressed growth in mouse xenograft model. A recent study of MKK4-/-cell lines derived from the pancreatic cell line PL5 (Cunningham et al., 2006), demonstrated that deletion of SEK1 leads to inhibition of tumor growth in vivo and also results in fewer lung metastases when cells were injected intravenously. Our data indicate that SEK1 is involved in the hypoxia-induced signaling pathway mediating increased oxaliplatin resistance. Signaling through this pathway(s) could involve not only targets of SEK1 other than JNK or p38 MAPK, and consequently the AP-1 transcriptional factor, but also upstream mediators of signaling to MAP kinases, which specifically interact with SEK1 but not with MKK7.
The very striking effect of modulation of these signaling pathways in cell culture was demonstrated also in vivo with the SEK1 and MKK7 inhibitory constructs. Signaling through MKK7 was required to elicit the therapeutic effects of bevacizumab, oxaliplatin, and their combination. Inhibition of SEK1 on the other hand, delayed the growth of untreated cells and potentiated the activity of the treatments. These data identify SEK1 as a potential target for reversal of hypoxic resistance to oxaliplatin. We have previously proposed a role for the induction of hypoxia in the action of bevacizumab (Selvakumaran et al., 2008). In HT29 colon adenocarcinoma sublines grown as xenografts (Yao et al., 2005a), hypoxia induction was observed to a comparable degree in all the cell lines, but growth inhibition was proportional to the susceptibility of the cell lines in vitro to hypoxia. The results of the in vivo experiments presented here confirm and extend these findings and show that a determinant of the outcome of bevacizumab treatment in this model is hypoxiainduced signaling through MKK7. The activity of this pathway is similarly required for the action of oxaliplatin: the MKK7-deficient line is resistant to oxaliplatin also. We hypothesize that the integrity of this pathway may be important for the activity of both bevacizumab and oxaliplatin in human colon cancer, and clinical studies are in progress to this end. One implication of this observation is that tumors may exhibit cross-resistance to both bevacizumab and oxaliplatin under conditions of impaired MKK7 signaling. The acquisition of MKK7 mutations would be expected to be favored in resistant colon tumors, whereas mutations of SEK1 would be advantageous for efficacy of oxaliplatin.
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ABBREVIATIONS: HIF-1, hypoxia-inducible factor-1; AP-1, activator protein-1; L-OHP, oxaliplatin; MAPK, mitogen-activated protein kinase; MAP, mitogen-activated protein; MEK or MKK, mitogen-activated protein kinase kinase; ERK, extracellular signal-regulated kinase; JNK, Jun NH2-terminal kinase; MEKK or MKKK, mitogen-activated protein kinase kinase kinase; SEK1 (MKK4), mitogen-activated protein kinase kinase 4; U0126, 1,4-diamino-2,3-dicyano-1,4-bis(methylthio)butadiene;; 17-AAG, 17-allylamino-17-demethoxygeldanamycin; HA, hemagglutinin; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; SB203580, 4-(4-fluorophenyl)-2-(4-methylsulfinylphenyl)-5-(4-pyridyl)1H-imidazole; SP600125, anthra[1-9cd]pyrazol-6(2H)-one; ATF2, activating transcription factor 2.
Address correspondence to: Dr. Irina A. Vasilevskaya, University of Pennsylvania, 1020 BRB II/III, 421 Curie Blvd., Philadelphia, PA 19104. E-mail: vasilevs{at}mail.med.upenn.edu
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