Sustained Activation of JNK/p38 MAPK Pathways in Response to Cisplatin Leads to Fas Ligand Induction and Cell Death in Ovarian Carcinoma Cells*

Abdellah Mansouri {ddagger} § , Lon D. Ridgway {ddagger} §, Anita L. Korapati {ddagger} §, Qingxiu Zhang {ddagger}, Ling Tian {ddagger}, Yibin Wang ||, Zahid H. Siddik **, Gordon B. Mills {ddagger} and François X. Claret {ddagger} {ddagger}{ddagger}

From the {ddagger} Department of Molecular Therapeutics, University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030, ** Department of Experimental Therapeutics, University of Texas M. D. Anderson Cancer Center, Houston, Texas 77030, || University of Maryland School of Medicine, Baltimore, Maryland 21201

Received for publication, August 8, 2002 , and in revised form, February 26, 2003.
    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
The efficacy of cisplatin in cancer chemotherapy is limited by the development of resistance. Although the molecular mechanisms involved in chemoresistance are poorly understood, cellular response to cisplatin is known to involve activation of MAPK and other signal transduction pathways. An understanding of early signal transduction events in the response to cisplatin could be valuable for improving the efficacy of cancer therapy. We compared cisplatin-induced activation of three MAPKs, JNK, p38, and ERK, in a cisplatin-sensitive human ovarian carcinoma cell line (2008) and its resistant subclone (2008C13). The JNK and p38 pathways were activated differentially in response to cisplatin, with the cisplatin-sensitive cells showing prolonged activation (8–12 h) and the cisplatin-resistant cells showing only transient activation (1–3 h) of JNK and p38. In the sensitive cells, inhibition of cisplatin-induced JNK and p38 activation blocked cisplatin-induced apoptosis; persistent activation of JNK resulted in hyperphosphorylation of the c-Jun transcription factor, which in turn stimulated the transcription of an immediate downstream target, the death inducer Fas ligand (FasL). Sequestration of FasL by incubation with a neutralizing anti-FasL antibody inhibited cisplatin-induced apoptosis. In contrast, chemoresistance in 2008C13 cells was associated with failure to up-regulate FasL. Moreover, in these cells, selective stimulation of the JNK/p38 MAPK pathways by adenovirus-mediated delivery of recombinant MKK7 or MKK3 led to sensitization to apoptosis through reactivating FasL expression. Thus, the JNK > c-Jun > FasL > Fas pathway plays an important role in mediating cisplatin-induced apoptosis in ovarian cancer cells, and the duration of JNK activation is critical in determining whether cells survive or undergo apoptosis.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cisplatin (cis-diamminedichloroplatinum(II) (CDDP)1) is a platinum-based compound that forms intra- and interstrand adducts with DNA (1, 2). CDDP has a broad spectrum of anti-tumor activity and is widely used in the treatment of solid tumors (3). However, one of the major limitations in its efficacy is that many tumors either are inherently resistant or acquire resistance after an initial response (1, 2, 3, 4). The molecular mechanisms that underlie this chemoresistance are largely unknown. Possible mechanisms of acquired resistance to CDDP include decreased platinum accumulation, elevated drug inactivation by metallothionine and glutathione, and enhanced DNA repair activity (5, 6). Increased expression of anti-apoptotic genes and mutations in the intrinsic apoptotic pathway may contribute to the inability of cells to detect DNA damage or to induce apoptosis (1, 7, 8, 9).

Because of the reactivity of CDDP and the complexity of the cellular response to DNA damage, CDDP-induced apoptotic signaling likely involves several pathways. Elucidation of the details of these signaling pathways is important because they may explain why tumor cells exposed to cisplatin often lose sensitivity to this agent and become resistant to apoptotic signals.

Genotoxic stress induces multiple signal transduction pathways, among which are the MAPK pathways. These pathways are parallel cascades of structurally related serine/threonine kinases that play pivotal roles in transducing various extracellular signals to the nucleus. The MAPK signaling cascades regulate a variety of cellular activities, including cell growth, differentiation, survival, and death (10, 11). In mammals, MAPKs are divided into three major groups, ERKs, JNKs/stress-activated protein kinases, and p38, based on their degree of homology, biological activities, and phosphorylation motifs (12). Even though these signaling systems are built from evolutionarily related protein kinases, they produce distinct biological responses. The biological effects of MAPK signaling are executed by phosphorylation of downstream substrates, most notably a number of signal-responsive transcription factors. The broad range of these substrates indicates that MAPKs have pivotal roles in cellular signal transduction and suggests that the extent and duration of MAPK activation play key roles in controlling cell functions.

The ERK pathway, which is induced in response to mitogenic stimuli such as peptide growth factors, cytokines, and phorbol esters, involves ERK1 and ERK2, the participation of Raf-1 and Ras oncoproteins, and the activation of MEK1/2 (12). Once activated, ERK phosphorylates several substrates, including Elk-1 (13). The ERK pathway plays a major role in regulating cell proliferation and differentiation (12) and provides a protective effect against apoptosis (14). On the other hand, the signaling cascades involving JNK and p38 are key mediators of stress signals and seem to be responsible mainly for protective responses, stress-dependent apoptosis, and inflammatory responses. These cascades can be stimulated by various stresses such as UV and {gamma}-irradiation, osmotic stress, and heat shock; pro-inflammatory cytokines such as tumor necrosis factor-{alpha} and interleukin-1{beta}; and chemotherapeutic drugs (10, 12).

To understand the molecular basis for the failure of CDDP-based chemotherapy, we compared the cellular responses of the human ovarian carcinoma cell line 2008 and its resistant subclone 2008C13 (15) after treatment with a platinum-based anticancer agent. We found that differences in the duration of the activation of MAPK pathways correlated with CDDP-induced apoptosis. A strong sustained activation of both pathways seemed to be a required priming step for CDDP-induced apoptosis; this activation of both JNK and p38 MAPK in CDDP-sensitive cells correlated with up-regulation of the Fas ligand (FasL), an immediate downstream target of JNK, and was accompanied by the induction of caspase activity and apoptosis. The failure of cisplatin to elicit such a response in the resistant variant indicates that impaired FasL expression could contribute to the development of chemoresistance. Reduction of cisplatin-induced apoptosis by the expression of dominant-negative c-Jun lacking JNK phosphoacceptor sites or by the use of either a small drug inhibitor of JNK/p38 or a neutralizing anti-FasL antibody further underlines the critical role of c-Jun-dependent FasL expression signaling in the induction of apoptosis by genotoxic agents.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Reagents—Cisplatin (Platinol-AQ cisplatin injection) was obtained from Bristol-Myers Squibb Co. Polyclonal antibodies to p38, phospho-p38 (Thr180/Tyr182), ATF-2, phospho-ATF-2 (Thr71), JNK, phospho-JNK (Thr183/Tyr185), c-Jun, phospho-c-Jun (Ser73), ERK, and phospho-ERK (Thr202/Tyr204) were purchased from Cell Signaling (Beverly, MA). The anti-JNK1 monoclonal antibody (clone 333.8), anti-human PARP antibody, anti-caspase-8 antibody, anti-human cytochrome c monoclonal antibody, neutralizing anti-human FasL antibody (NOK-2), and isotype-matched control antibody were obtained from Pharmingen. The anti-Fas monoclonal antibody (CH-11) was purchased from Medical & Biological Laboratories (Watertown, MA). The anti-capase-3/CPP32 antibody was purchased from Transduction Laboratories (Lexington, KY). Anti-{beta}-actin monoclonal antibodies were obtained from Sigma. The caspase inhibitor Z-VAD-fmk and the JNK and p38 kinase inhibitor SB202190 were purchased from Alexis Biochemicals (San Diego, CA). CDDP-sensitive (2008) and CDDP-resistant (2008C13) ovarian cancer cells were kindly provided by Drs. S. B. Howell (University of California at San Diego, La Jolla, CA), S. G. Chaney (University of North Carolina, Chapel Hill, NC), and Z. H. Siddik (M. D. Anderson Cancer Center). The 2008 cell line, established from a patient with serous cystadenocarcinoma of the ovary, and its resistant subclone 2008C13, derived from 2008 cells by in vitro exposure to CDDP, have been characterized by Howell and co-workers (15) and Chaney and co-workers (16). Wild-type c-jun and c-jun/ 3T3 fibroblasts were a gift from Drs. E. F. Wagner (Research Institute for Molecular Pathology, Vienna, Austria) and M. Karin (University of California at San Diego).

Cell Culture and Adenoviral Infection—The CDDP-sensitive human ovarian carcinoma cell line 2008 and its resistant variant 2008C13 were maintained in RPMI 1640 medium supplemented with 10% fetal calf serum (Invitrogen) and 1% penicillin/streptomycin. Wild-type c-jun and c-jun/ 3T3 cells were cultured in Dulbecco's modified Eagle's medium supplemented as described above. Cells were incubated at 37 °C in a humidified atmosphere containing 5% CO2. Recombinant adenoviral vectors expressing green fluorescent protein (Ad-GFP) and activated mutants of MKK7 and MKK3 (Ad-MKK7D and Ad-MKK3bE) were constructed as previously described (17). Cells were infected with adenoviruses at a multiplicity of infection of 50 plaque-forming units/cell for 5 h and then incubated for another 30 h to allow expression of the protein of interest as described (17).

Immunoblot Analysis—Cells in log-phase growth were treated or not treated with CDDP at 20 µM (2008 and 2008C13 cells) or 100 µM (wild-type c-jun and c-jun/ 3T3 cells) for 1 h, after which they were washed, and fresh medium was added. At various times after CDDP exposure (1 min and 1, 3, 5, 8, and 12 h), the cells were collected and lysed in lysis buffer (25 mM HEPES, pH 7.7, 400 mM NaCl, 0.5% Triton X-100, 1.5 mM MgCl2, 2 mM EDTA, 2 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, protease inhibitors (10 µg/ml leupeptin, 2 µg/ml pepstatin, 50 µg/ml antipain, 2 µg/ml aprotinin, 20 µg/ml chymostatin, and 2 µg/ml benzamidine), and phosphatase inhibitors (50 mM NaF, 0.1 mM Na3VO4, and 20 mM {beta}-glycerophosphate)). For PARP and caspase immunoblotting, cell lysates were prepared using radioimmune precipitation assay lysis buffer (50 mM Tris, pH 7.5, 1% Nonidet P-40, 0.25% sodium deoxycholate, 150 mM NaCl, 1 mM EDTA, 1 mM phenylmethylsulfonyl fluoride, 2 µg/ml aprotinin, 1 µg/ml pepstatin, and 2 µg/ml leupeptin). Aliquots of cell lysates (70 µg of protein) were resolved by 10–12% SDS-PAGE, transferred onto polyvinylidene difluoride membrane (Immobilon, Millipore Corp., Bedford, MA) or Hy-bond-P membrane (Amersham Biosciences), and probed with the appropriate primary antibodies. Reactions were visualized with a suitable secondary antibody conjugated with horseradish peroxidase (Bio-Rad) using enhanced chemiluminescence reagents (Amersham Biosciences).

Drug Uptake and Adduct Level Assays—Cells were treated with 20 µM CDDP, after which cells were washed with phosphate-buffered saline (PBS), and fresh medium was added. Cell pellets were made immediately after 1 min and 1 h of CDDP exposure. After 1 h of drug exposure, cells were washed and cultured in drug-free medium for an additional 4 h. For protein analysis, cells were digested overnight in 0.2 N NaOH at 55–60 °C. Intracellular platinum levels were determined by solubilizing the cell pellet in Hyamine hydroxide and analyzed by flameless atomic absorption spectrophotometry using conditions previously described (detection limit = 100 pg of platinum) (18, 19). For platinum adduct levels, cell pellets were lysed in extraction buffer (10 mM Tris, pH 8.0, 100 mM EDTA, 20 µg/ml RNase, and 0.5% SDS) overnight at 37 °C and then treated with proteinase K (100 µg/ml) for 3 h at 50 °C, and the DNA was extracted in phenol/chloroform. The amount of platinum bound to DNA was determined by flameless atomic absorption spectrophotometry.

Immunocomplex Kinase Assays—Cells were serum-starved in 0.1% serum for 12–16 h before CDDP treatment. Whole cell extracts were prepared and treated as previously described (20). Briefly, endogenous JNK1 was immunoprecipitated from 300 µg of cell lysate with the anti-JNK1 monoclonal antibody (clone 333.8) and protein A-agarose beads for 4 h at 4 °C. The precipitates were washed twice with lysis buffer and twice with kinase buffer (25 mM Hepes, pH 7.6, 20 mM MgCl2,20mM {beta}-glycerophosphate, 0.1 mM sodium orthovanadate, 2 mM DTT). JNK kinase activity was measured using 2 µg of glutathione S-transferase (GST)-c-Jun-(1–79) as the substrate, and the reaction was initiated by the addition of 10 µM ATP and 10 µCi of [{gamma}-32P]ATP (5000 Ci/mmol; ICN Biomedicals, Aurora, OH). After the cells were incubated for 30 min at 30 °C, the reactions were stopped with Laemmli sample buffer. The proteins were resolved by 12% SDS-PAGE and visualized by autoradiography.

Cell Proliferation Assay—Cell proliferation was assessed in 96-well plates after cells had been treated with CDDP for 1 h, washed to remove the drug, and left to proliferate for the indicated times after the addition of fresh medium. The number of surviving cells was measured by nucleic acid staining with the CyQUANT cell proliferation assay kit (Molecular Probes, Inc., Eugene, OR) 4–5 days after seeding. The assay was conducted according to the manufacturer's instructions. The samples were analyzed on a Fluoroskan Ascent CF microplate fluorometer (ThermoLabSystems, Helsinki, Finland). All experiments were carried out in quadruplicate, and the proliferation rate was expressed as the ratio of the number of proliferating cells treated with CDDP to the number of proliferating cells not treated with CDDP.

Flow Cytometry Analysis—To measure DNA content (apoptotic nuclei), cells were harvested; washed with PBS; fixed in 1% paraformaldehyde; stained with a solution containing 15 µg/ml propidium iodide, 0.5% Tween 20, and 0.1% RNase A; and incubated for 30 min at 24 °C. Cells were sorted using a FACScan (BD Biosciences) and analyzed with CELLQuest Version 3.3 software. Data were plotted on a logarithmic scale.

Detection of Fas and FasL mRNA Expression by Reverse Transcriptase-PCR—Total RNA was isolated from the 2008 and 2008C13 cell lines using the RNeasy minikit (QIAGEN Inc., Valencia, CA) according to the instructions of the manufacturer. The reverse transcriptase assay was performed with 2 µg of total RNA using Superscript II reverse transcriptase (Invitrogen) according to the manufacturer's recommendations. A reaction without reverse transcriptase was performed in parallel to ensure the absence of genomic DNA contamination. PCR amplification was carried out in a final volume of 50 µl containing 5 µl of cDNA, 5 µl of 10x PCR buffer (10 mM Tris-HCl, pH 9, 50 mM KCl, and 0.1% Triton X-100), 0.5 µl of dNTP (10 µM), 3 µl of MgCl2 (25 mM), and 2.5 units of AmpliTaq Gold (PerkinElmer Life Sciences). PCR conditions were as follows: an initial denaturation step at 94 °C for 5 min, followed by 35 cycles at 94 °C for 30 s, either 57 °C (for Fas) or 61 °C (for FasL and {beta}-actin) for 30 s, and 72 °C for 30 s. After a final extension at 72 °C for 5 min, PCR products were resolved on 1.2% agarose gels and visualized by ethidium bromide transillumination under UV light. Primer sequences were as follows: Fas, 5'-ATT TCT GCC ACT GCA GCC CTC AGG-3' (forward) and 5'-TCC AGT TCG CTG GGC AGA CTT CTC-3' (reverse); and FasL, 5'-ATG TTT CAG CTC TTC CAC CTA CAG A-3' (forward) and 5'-CCA GAG AGA GCT CAG ATA CGT TGA C-3' (reverse). These sequences span nucleotides 76–706 of Fas cDNA and nucleotides 365–856 of FasL cDNA and yield PCR products of 630 and 492 bp, respectively (21). Each reverse-transcribed mRNA product was internally controlled by {beta}-actin PCR using primers 5'-TGA CGG GGT CAC CCA CAC TGT GCC CAT CTA-3' (forward) and 5'-CTA GAA TTT GCG GTC GAC GAT GGA GGG-3' (reverse), covering region 2199–3065 of {beta}-actin cDNA and yielding a 867-bp PCR product (21). The FasL and Fas reverse transcriptase-PCR products were subsequently confirmed by direct sequencing.

Propidium Iodide and 4,6-Diamidino-2-phenylindole (DAPI) Staining—To detect apoptosis, nuclear staining was performed using 5 µg/ml DAPI, and cells were analyzed with a fluorescence microscope (magnification x400 for nuclear analysis and x100 for morphologic analysis). Apoptotic cells were identified by morphology and by condensation and fragmentation of their nuclei. The percentage of apoptotic cells was calculated as the ratio of apoptotic cells to total cells counted, multiplied by 100. Three separate experiments were conducted, and at least 300 cells were counted for each experiment.

Transfection and Immunofluorescence Staining—Expression vectors for hemagglutinin (HA) epitope-tagged wild-type c-Jun (pSR{alpha}-HA-c-Jun) and dominant-negative c-Jun (pSR{alpha}-HA-c-Jun(S63A/S73A)) were a gift from Dr. M. Karin. Liposome-mediated transfection was performed using LipofectAMINE Plus (Invitrogen). Briefly, the 2008 ovarian carcinoma cells were grown on chamber slides and transfected with vectors containing the HA epitope tag. After transfection, the cells were washed with PBS and fixed in methanol for 10 min at –20 °C, after which they were air-dried, washed three times with PBS, blocked in 1.5% bovine serum albumin in PBS (PBS/bovine serum albumin) for 1 h at room temperature, and then immunostained with a monoclonal antibody to HA (1:50 dilution in PBS/bovine serum albumin) for 1 h at room temperature. After three washes with PBS, transfected cells were visualized by incubation with a fluorescein isothiocyanate-conjugated rabbit anti-mouse antibody (1:40 dilution in PBS/bovine serum albumin; Dako, Carpinteria, CA) for 45 min at 37 °C. To visualize the nuclei of transfected cells, we included DAPI (5 µg/ml) in the wash after the incubation with the secondary antibody. Cells were examined and photographed with an Olympus microscope equipped for epifluorescence with the appropriate filters. Transfected cells were scored blindly for apoptosis.

Detection of Cytochrome c Release—Cytosol extracts were prepared from the 2008 and 2008C13 cells essentially as previously described (22). Briefly, after cisplatin treatment and incubation for 6, 12, 18, and 24 h, the cells were collected by centrifugation. The cell pellet was washed twice with cold PBS and resuspended in ice-cold buffer A (20 mM HEPES, pH 7.5, 1.5 mM MgCl2, 10 mM KCl, 1 mM EDTA, 1 mM EGTA, 1 mM dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 10 µg/ml pepstatin A) containing 250 mM sucrose. The cells were homogenized with 25 strokes of a Dounce homogenizer with a type B pestle. Nuclei and intact cells were cleared by centrifugation at 1000 x g for 10 min at 4 °C. The supernatant was centrifuged at 14,000 x g for 20 min at 4 °C to pellet the mitochondrial fraction. An aliquot of the resulting supernatant was used as the soluble cytosolic fraction. The mitochondrial pellet was washed once and then suspended in buffer A. Protein extracts (equal amounts in the mitochondrial and cytosolic fractions) were subjected to Western blot analysis with a monoclonal antibody to cytochrome c.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
CDDP-induced Apoptosis in Ovarian Carcinoma Cells— CDDP-sensitive 2008 cells and CDDP-resistant 2008C13 cells were exposed to CDDP for 1 h, after which the drug was washed out to mimic in vivo chemotherapy. The time course for the induction of apoptosis was determined by microscopic examination of DAPI-stained cells (Fig. 1).



View larger version (52K):
[in this window]
[in a new window]
 
FIG. 1.
CDDP induces massive apoptosis in chemosensitive 2008 cells, but only moderate apoptosis in chemoresistant 2008C13 ovarian carcinoma cells. A, shown are representative micrographs illustrating morphologic evidence of apoptosis as assessed by nuclear staining at 24 h after CDDP treatment. 2008 and 2008C13 cells were treated with 20 µM CDDP for 1 h and then cultured in CDDP-free fresh medium for the remaining time, after which the cells were fixed, and nuclear condensation was analyzed with DAPI. Typical morphologic changes associated with apoptosis were visualized by fluorescence microscopy (magnification x400). B, CDDP induced apoptosis in ovarian carcinoma cells in a time-dependent manner. 2008 and 2008C13 cells were treated for 1 h with 20 µM CDDP and collected at the indicated times. Cells were then subjected to DAPI staining and fluorescence microscopy counting as described for A. The results shown are the means of five experiments. C, shown are the results from immunoblot analysis illustrating differential effects of CDDP on activation of caspase-3 and cleavage of the caspase substrate PARP in the two cell lines. Cells were treated as described for A and then harvested, and their protein extracts were analyzed by SDS-PAGE using anti-caspase-3 (Casp-3) and anti-PARP antibodies. An anti-{beta}-actin antibody was used as a loading control.

 

In the chemosensitive 2008 cells, exposure to 20 µM CDDP resulted in morphologic alterations characteristic of apoptosis, including membrane blebbing, nuclear condensation and fragmentation (Fig. 1A), and DNA laddering (data not shown). The number of apoptotic cells increased with time and accounted for 50–70% of the total cell population by 18–24 h. The CDDP-resistant 2008C13 cells, in contrast, had a markedly different apoptotic response to this "pulsed" exposure to CDDP (Fig. 1B). Immunoblot analysis revealed cleavage of the pro form of caspase-3 (32 kDa) to its active form (17 kDa), compatible with the induction of apoptosis, from 12 to 48 h after CDDP treatment in 2008 cells, but not in 2008C13 cells (Fig. 1C). PARP cleavage products also persisted from 12 to 48 h in CDDP-treated 2008 cells as detected by immunoblot analysis, whereas in the 2008C13 cell extracts, no PARP cleavage fragments were detected, which correlated with caspase-3 (Fig. 1C). Immunoblotting with an anti-{beta}-actin antibody was used as a loading control. This finding reflected the resistance of 2008C13 cells to CDDP-induced apoptosis.

To study the mechanism behind this CDDP resistance, we first studied drug uptake and DNA adduct formation in both the sensitive and resistant 2008 cells (Table I). As shown, drug uptake and DNA adduct formation were similar in the two cell lines at 1 min after CDDP treatment, whereas at 1 and 5 h after treatment, the differences between the CDDP-sensitive and CDDP-resistant cells were <2-fold, with the resistant cells showing a lower value of DNA adduct formation and drug uptake. Resistance to CDDP in 2008C13 cells is >2-fold (15, 16), which prompted us to investigate additional mechanisms of resistance.


View this table:
[in this window]
[in a new window]
 
TABLE I
CDDP uptake and CDDP-DNA adduct formation in 2008 and 2008C13 cells

 

Differential Activation of MAPK Pathways by CDDP in Sensitive Versus Resistant Cell Lines—Because activation of MAPKs and phosphorylation of c-Jun have been reported after treatment with chemotherapeutic drugs in other cell types (23, 24, 25, 26, 27), we compared the activation of JNK between CDDP-sensitive and CDDP-resistant ovarian cancer cells after treatment with cisplatin. The activity of immunoprecipitated JNK was assayed using a GST-c-Jun fusion protein as substrate. CDDP treatment of the sensitive 2008 cells induced an increase in the ability of the JNKs to phosphorylate the GST-c-Jun substrate, beginning 1 h after treatment and persisting through the next 3–5 h (Fig. 2A). On the other hand, extracts from CDDP-treated resistant 2008C13 cells showed only transient JNK activity after 1 h of treatment, and this activity declined rapidly over the next 3–5 h (Fig. 2A).



View larger version (44K):
[in this window]
[in a new window]
 
FIG. 2.
Activation of the MAPK signal transduction pathway in response to CDDP in ovarian cancer cell lines 2008 and 2008C13. A, effect of CDDP on JNK activity. Cells were kept in 0.1% serum for 12–16 h and either exposed to 20 µM CDDP for 1 h or left untreated (NT). At 1, 3, and 5 h after completion of CDDP treatment, cells were collected, lysed, and subjected to JNK1 immunocomplex kinase assays using the GST-c-Jun-(1–79) fusion protein as a substrate. An equal number of cells were harvested at the indicated times. B, time course of CDDP-induced p38, ATF-2, JNK, c-Jun, and ERK phosphorylation. CDDP-sensitive (2008) and CDDP-resistant (2008C13) human ovarian carcinoma cells were exposed for 1 h to 20 µM CDDP or left untreated (0). At the indicated times after completion of CDDP treatment, whole cell extracts were prepared, and protein extracts were resolved by SDS-PAGE and immunoblotted with anti-phospho-p38 and anti-phospho-ATF-2, anti-phospho-JNK and anti-phospho-c-Jun, and anti-phospho-ERK1/2 antibodies, which recognize the activated forms of p38, JNK, and ERK, respectively. The total amount of p38, ATF-2, JNK1, and c-Jun proteins was assessed using antibodies that recognize these proteins independent of their phosphorylation status.

 

Next, we investigated the effect of CDDP on the phosphorylation of JNK and p38 as well as that of their respective target substrates, the c-Jun and ATF-2 transcription factors, over time. CDDP treatment of the 2008 cells, which resulted in significant apoptosis, led to sustained activation (from 1 min to 12 h after treatment) of JNK and p38, as assessed by their phosphorylation states using specific antibodies that recognize the phosphorylated (activated) forms of theses enzymes (Fig. 2B). p38 MAPK activation occurred over the same period as JNK activation. Although phosphorylation was detected very early (at 1 min), maximal phosphorylation of both kinases became apparent at ~1 h and was sustained over the following 12-h period (Fig. 2B, left panels). Consequently, phosphorylation of c-Jun at Ser73 and of ATF-2 at Thr71 occurred over the same extended period in 2008 cells (Fig. 2B, left panels). This sustained phosphorylation did not result from increased expression of either JNK or p38, as protein levels were unaltered relative to untreated cells (Fig. 2B, left panels). In contrast, the CDDP-resistant variant 2008C13 showed only transient (1–3 h after treatment) JNK and p38 phosphorylation, a pattern that was reproduced upon phosphorylation of the c-Jun and ATF-2 target substrates (Fig. 2B, right panels). Similar results were found with regard to the differences in duration of JNK activation and c-Jun phosphorylation in studies of another ovarian carcinoma cell with CDDP-sensitive and CDDP-resistant variants (A2780 and A2780CP) (data not shown).

Next, we assessed whether cisplatin might activate the ERK pathway in ovarian cells, and we found that CDDP induced transient ERK phosphorylation in both the 2008 and 2008C13 cell lines. Phosphorylation of ERK was detected at a 1-h time point in the 2008 cells, but an early, somewhat persistent phosphorylation (1 min to 1 h) was observed in the 2008C13 cells (Fig. 2B). This ERK phosphorylation pattern was not sustained and was the opposite of the phosphorylation pattern seen for the JNK and p38 MAPKs. In the resistant cells, ERK phosphorylation also seemed to be biphasic, with a second increase occurring at 12–16 h. These data agree with previous findings that CDDP-induced apoptosis correlates with an increase in JNK and c-Jun phosphorylation in other cell types (23, 26, 27). Our findings also suggest that the difference in duration of the activation of the JNK and p38 MAPK pathways in CDDP-sensitive and CDDP-resistant cells could contribute to directing the outcome to survival or apoptosis.

Absence of c-Jun or Inhibition of JNK Pathways Confers Resistance to Apoptosis and Increases Cell Survival after Treatment with CDDP—The delay between JNK (or p38) activation and the onset of apoptosis, together with the requirement for prolonged JNK (or p38) phosphorylation, suggests that expression of new genes may be required to activate apoptosis. Because the c-Jun transcription factor is an important and specific target for JNK (10) and is possibly involved in the apoptosis triggered by DNA-damaging agents (26, 27), we hypothesized that prolonged activation of the JNK and p38 pathways induced by CDDP is required to induce apoptosis in a manner dependent on c-Jun transcription. Conversely, we also hypothesized that a transient activation or immediate inactivation of these kinase pathways is too brief to transactivate AP-1-responsive genes and thus would lead to resistance to CDDP-induced apoptosis.

To further investigate the involvement of c-Jun activation in CDDP-induced apoptosis, we compared immortalized 3T3 fibroblast cell lines that have a targeted disruption of the c-jun gene (28) with their parental 3T3 cells that express the wild-type c-jun gene. We first analyzed whether the absence of the c-jun gene affected cell survival after CDDP exposure. As shown in Fig. 3A, as indicated by DAPI staining, the CDDP-treated c-jun/ cells were significantly more resistant to apoptosis compared with the parental cells. About 80% of the c-jun/ 3T3 cells survived CDDP treatment, but only 25% of the wild-type c-jun 3T3 cells were still viable after 4–5 days, suggesting that activation of a downstream set of target genes through c-jun leads to CDDP-induced apoptosis. To determine the possible role of the specific members of the MAPK family in mediating this process, we tested the parental and c-jun/ 3T3 cells with the pyridinylimidazole compound SB202190, a strong inhibitor of JNK and the p38/HOG kinase (29, 30, 31). Survival measurements revealed that pretreatment with SB202190 led to a significant increase in cell survival in both the absence and presence of c-jun (Fig. 3A). Inhibition of JNK and p38 decreased the effect of CDDP in wild-type c-jun 3T3 cells (72% survived with SB202190 pretreatment versus 26% without). The results of an immunocomplex kinase assay, which showed a decrease in phosphorylation of GST-c-Jun by immunoprecipitated JNK (Fig. 3B), confirmed that SB202190 treatment inhibited CDDP-induced c-Jun N-terminal phosphorylation and that this inhibition of c-Jun phosphorylation correlated with the inhibition of apoptosis (Fig. 3A).



View larger version (45K):
[in this window]
[in a new window]
 
FIG. 3.
CDDP-induced apoptosis depends on c-Jun. A, wild-type c-jun or c-jun/ 3T3 cells were left in 0.25% serum for 12 h and treated with the JNK activation inhibitor SB202190 at 30 µM for 1 h. The cells were then treated with 100 µM CDDP for 1 h and washed four times to remove the drug, and fresh medium containing 0.25% serum was added. Cell viability was measured 4 days later using a fluorescence-based nucleic acid method (the CyQUANT cell proliferation assay kit). B, SB202190 inhibited CDDP-induced JNK activity in 3T3 cells. Wild-type c-jun or c-jun/ 3T3 cells were treated for 1 h with or without SB202190 (SB) as described for A before exposure to CDDP for 1 h. Cell lysates were prepared 1 h after CDDP treatment, and JNK activity was analyzed in an immunocomplex kinase assay using GST-c-Jun-(1–79) as a substrate. Cont., control.

 

These data confirm that c-Jun and its activation by JNK were required for efficient induction of apoptosis by the alkylating agent CDDP. Blockade of JNK activation, whether by a small drug inhibitor or by a lack of c-jun, protected cells from cisplatin-induced apoptosis.

A JNK and p38 Inhibitor Prevents CDDP-induced Apoptosis—To further investigate whether the JNK pathway is required for CDDP-induced apoptosis, we pretreated 2008 cells with SB202190 under conditions in which activation of both JNK and p38 was inhibited (data not shown) before CDDP exposure. Incubation with SB202190 resulted in a marked reduction in cell death (Fig. 4A). However, cells treated with CDDP alone displayed the typical features of apoptosis: condensation of the nuclei (Fig. 4A, left panels), shrinkage of the cytoplasm, and membrane blebbing (as seen by phase-contrast microscopy) (data not shown). Interestingly, pretreatment of the cells with SB202190 markedly suppressed the morphologic changes induced by CDDP (Fig. 4A, left panels). To confirm these findings with an independent assay, we measured apoptosis by propidium iodide staining and flow cytometry. At 24 h after treatment with CDDP, 51% of the 2008 cells showed a hypodiploid (sub-G1) DNA content, reflecting apoptosis (Fig. 4B). However, incubation with SB202190 before CDDP treatment reduced the extent of cell death considerably, from 51 to 9% (Fig. 4B), and inhibited CDDP-induced cell death in a dose-dependent manner (data not shown).



View larger version (60K):
[in this window]
[in a new window]
 
FIG. 4.
Effect of the JNK/p38 MAPK inhibitor SB202190 on CDDP-induced apoptosis in CDDP-sensitive cells. A, shown are the morphologic changes in chemosensitive 2008 cells at 24 h after a 1-h treatment with CDDP either followed or preceded by treatment with SB202190 (SB). Control cells were untreated. Apoptosis was visualized after DAPI staining by fluorescence microscopy (magnification x400) (left panels). The results from quantitative analysis are shown (right panel). B, shown are the results from flow cytometry of propidium iodine-stained untreated 2008 cells and cells treated with CDDP and SB202190 as described for A. Cells were stained with propidium iodine, and the number of apoptotic cells was counted with a FACSCalibur flow cytometer. The percentages of apoptotic cells are indicated as the proportion of cells that contained sub-G1 DNA. Representative results from two independent experiments are shown. C, a phosphorylation-defective c-Jun mutant inhibited CDDP-induced cell death in 2008 cells. Cells cultured on glass coverslips were transfected with HA-tagged wild-type (wt) c-Jun or c-Jun(S63A/S73A) expression vectors. After 24 h, the cells were washed with PBS and incubated for 1 h with 20 µM CDDP. The cells were subsequently fixed and permeabilized, and expression of fluorescein isothiocyanate (FITC)-labeled HA-c-Jun proteins was detected by indirect immunofluorescence using an-HA monoclonal antibody. Nuclear morphology was visualized by staining with DAPI. The white arrowheads indicate cells transfected with either wild-type HA-c-Jun (cells have apoptotic morphology) or HA-c-Jun(S63A/S73A) (cells have non-apoptotic morphology). The gray arrowheads indicate surrounding cells with apoptotic morphology. The results are quantified in the lower panel, where at least 300 cells were counted for each experiment.

 

To determine whether phosphorylation of the c-Jun transactivation domain is necessary for the induction of apoptosis, we transfected 2008 cells with either wild-type c-Jun or HA-c-Jun(S63A/S73A), in which Ser63 and Ser73, which are normally phosphorylated by JNK, have been replaced with alanine, resulting in an inactive protein (20). Transient expression of HA-c-Jun(S63A/S73A) was associated with a marked decrease in CDDP-induced apoptosis compared with the level seen in cells transfected with wild-type c-Jun (Fig. 4C). This protective effect was restricted to the c-Jun(S63A/S73A)-expressing cells, whereas in the surrounding cells that did not express this protein, the induction of apoptosis was not inhibited. As shown in Fig. 4C, ~30% of the 2008 cells expressing wild-type c-Jun were undergoing apoptosis compared with only 7–8% of the 2008 cells expressing HA-c-Jun(S63A/S73A). These data confirm the hypothesis that either c-Jun phosphorylation or JNK activation is required for CDDP-induced apoptosis and that inhibition of this pathway could exert a protective effect.

Expression of FasL in Response to CDDP Is Impaired in Resistant 2008C13 Variants, but Not in CDDP-sensitive Ovarian Carcinoma Cells—Because the ability of CDDP to induce apoptosis seems to depend on its ability to activate JNK- and c-Jun-dependent transcriptional events, we next sought to identify JNK and c-Jun targets that might mediate CDDP-induced apoptosis. To determine whether the FasL gene is a target of CDDP-dependent c-Jun activation, we examined whether exposure of ovarian carcinoma cells to CDDP affected the expression of FasL. Treatment of the CDDP-sensitive 2008 cells with CDDP led to an up-regulation of FasL (Fig. 5A) that began at 6 h and peaked at 18–24 h, a time span that corresponds to the kinetics of apoptotic cell death (Fig. 1B). In contrast, no FasL mRNA was detected in the CDDP-resistant 2008C13 cells (Fig. 5A, lanes 5–8), even at higher concentrations of CDDP (40 µM) (lanes 14–18). Fas receptor mRNA expression levels did not change after CDDP treatment and were comparable in both cell lines (Fig. 5A). These data identify the FasL gene as a c-jun-regulated target gene for CDDP and suggest that lack of FasL induction may contribute to the CDDP-induced apoptosis defect in the resistant 2008C13 cells. To explore this possibility, we examined whether blocking Fas/FasL interaction after CDDP treatment protected cells from undergoing apoptosis. We used the NOK-2 antibody, which recognizes and neutralizes both membrane-bound and membrane-soluble forms of human FasL, thereby preventing the interaction of either form with Fas. Incubation of the 2008 cells with this neutralizing anti-FasL IgG antibody after CDDP treatment protected the cells from CDDP-induced apoptosis by 30 and 40% at 24 and 48 h, respectively, compared with the level of apoptosis in cells that had been preincubated with an isotype-matched IgG control (Fig. 5B).



View larger version (44K):
[in this window]
[in a new window]
 
FIG. 5.
CDDP leads to up-regulation of FasL in chemosensitive 2008 cells, but not in chemoresistant 2008C13 cells. A, FasL mRNA was induced in 2008 cells and abrogated in 2008C13 cells after stimulation with CDDP. Cells were treated with CDDP at either 20 µM (left panels) or 20 and 40 µM (right panels). Total RNA was purified at the indicated times, and the expression of FasL, Fas receptor, and {beta}-actin mRNAs was examined by reverse transcriptase-PCR using specific primers. B, a neutralizing anti-FasL antibody (NOK-2) abrogated CDDP-induced apoptosis. 2008 ovarian carcinoma cells were treated with CDDP for 1 h, and then the cells were cultured in the presence of either the NOK-2 antibody (50 µg/ml) or an isotype-matched control Ig (50 µg/ml). At 0, 6, 12, 24, and 48 h after treatment, apoptosis was evaluated by DAPI staining and fluorescence microscopy. The percentage of apoptotic cells was determined as described under "Experimental Procedures." C, Fas-mediated apoptosis showed that Fas/FasL was functional in 2008 and 2008C13 cells. For ligation of Fas, cells were treated with a purified anti-Fas monoclonal antibody (Ab; 50 µg/ml) for 24 or 48 h. The percentage of Fas-mediated apoptosis was determined as described for B. D, 2008 cells were treated as described in the legend to Fig. 4A; total RNA was extracted; and expression of FasL and {beta}-actin mRNAs was determined by reverse transcriptase-PCR. E, the conditions were the same as described in the legend to Fig. 4A, except that cells were treated with an anti-Fas monoclonal antibody after SB202190 (SB) treatment as indicated.

 

Because interfering with the Fas/FasL system is known to reduce sensitivity to drug-mediated apoptosis in some cell systems (32, 33, 34), we next determined whether Fas/FasL function influenced apoptosis in the resistant 2008C13 cells by treating the cells with an anti-Fas antibody that binds to the Fas antigen, a process that mimics the role of FasL in the induction of apoptosis. The extent of Fas-mediated apoptosis was similar in the sensitive and resistant cells at 24 and 48 h (Fig. 5C). These results suggest that the Fas receptor that was expressed on the surface of both the resistant and sensitive 2008 cells was functional and could trigger apoptosis. Taken together, these data show that both FasL up-regulation and Fas/FasL interactions were important for the induction of apoptosis in ovarian carcinoma cells after their exposure to CDDP.

To determine whether the JNK/p38 MAPK pathway in ovarian cancer cells was involved in the FasL-mediated apoptosis induced by CDDP, we tested whether inhibiting JNK and p38 with the small drug inhibitor SB202190 blocked FasL expression. We found that pretreatment of the 2008 cells with SB202190 significantly inhibited FasL mRNA induction in response to CDDP (Fig. 5D), but had no significant effect on Fas-mediated apoptosis (Fig. 5E). The effect of this inhibition on FasL transcriptional activity might have been due to the inhibition of AP-1 activity because SB202190 completely inhibited JNK activation and therefore c-Jun phosphorylation (29, 30, 31). Taken together, these findings strongly suggest that FasL induction is required for CDDP-induced ovarian carcinoma apoptosis and that this process depends on JNK and the phosphorylation of c-Jun at Ser63 and Ser73. However, incubation of cells with SB202190 did not prevent Fas/FasL from triggering apoptosis (Fig. 5E), suggesting that the JNK pathway might not be downstream or might not be required for Fas-mediated apoptosis.

Caspase Inhibition Does Not Block CDDP-induced FasL Expression—Initiation of apoptotic cell death and caspase activation in response to various stimuli, including CDDP, requires the release of cytochrome c from the mitochondrial intermembrane space into the cytosol (35). In the cytoplasm, cytochrome c promotes the assembly of a protein complex called the apoptosome, which includes caspase-9 bound to the CED-4 homolog Apaf-1 (36, 37). Upon activation, caspase-9 instigates a proteolytic cascade involving multiple caspases, a process culminating in the cleavage of numerous substrate proteins and, ultimately, cell death (38). Activation of caspase-3 and cleavage of PARP were seen from 12 to 48 h after CDDP treatment in the CDDP-sensitive cells undergoing apoptosis (Fig. 1C). Caspases may function both upstream and downstream of Fas/FasL in the apoptotic signaling pathways. To examine further the effect of caspase activation on CDDP-induced apoptosis in the 2008 cells, we used the pan-caspase inhibitor Z-VAD-fmk to determine whether activation of FasL was linked to the caspase activation leading to apoptosis after CDDP treatment. We treated the 2008 cells with CDDP in the absence or presence of Z-VAD-fmk (50 µM) and examined them at 18 and 24 h after treatment. As shown in Fig. 6, Z-VAD-fmk substantially attenuated CDDP-induced PARP cleavage and apoptosis, as determined by visualization of DAPI-stained cells (data not shown); however, Z-VAD-fmk had no effect on CDDP-induced FasL induction (Fig. 6, A–C). These findings indicate that, in ovarian carcinoma cells, CDDP-induced FasL up-regulation is upstream of caspase activation.



View larger version (27K):
[in this window]
[in a new window]
 
FIG. 6.
Induction of caspase activity and cytochrome c release by CDDP. A, the caspase inhibitor Z-VAD-fmk prevented CDDP-mediated cell death without affecting CDDP-induced FasL transcription. CDDP-induced apoptosis was caspase-dependent in chemosensitive 2008 cells. Cells were treated with 20 µM CDDP in the presence or absence of 50 µM Z-VAD-fmk as indicated. Cells were then harvested at the indicated times; cell lysates were prepared; and PARP cleavage was examined by immunoblotting with an anti-PARP monoclonal antibody. B, CDDP-induced apoptosis was inhibited by Z-VAD-fmk. Cells were treated as described for A, and the percentage of cells exhibiting apoptotic features was determined. C, CDDP-related FasL mRNA induction was insensitive to Z-VAD-fmk. Cells were treated as described for A, and FasL and {beta}-actin mRNAs were examined by reverse transcriptase-PCR. D, shown are the time course and release of cytochrome c in the CDDP-sensitive cell line 2008 and its resistant variant 2008C13. Cells were collected at the indicated time points after a 1-h exposure to CDDP, and cytosolic (C) and mitochondrial (M) fractions were subjected to Western blot analysis with a monoclonal antibody to cytochrome c (CYT-C; upper panels). The blot was reprobed with an anti-actin antibody to evaluate the loading of the extracts (lower panels). Cont., control.

 

Recent findings have indicated that activation of the JNK pathway influences cytochrome c release and that apoptotic stimuli fail to release cytochrome c in JNK-null cells (39). To determine whether cytochrome c is released in response to treatment with CDDP and participates in the JNK pathway, 2008 cells were treated with CDDP, after which cytosolic fractions and mitochondrial proteins were extracted and tested for cytochrome c release by immunoblotting. Cytochrome c levels decreased in the mitochondria and increased in the cytoplasm of the 2008 cells, but not the 2008C13 cells, at 18–24 h after CDDP treatment (Fig. 6D). This observation agreed with our other findings on the extent and time course of PARP cleavage and apoptosis in these cells (Figs. 1 and 6, A and B). Whereas JNK activation in the 2008 cells was detected earlier (1 min to 5 h) after CDDP treatment (Fig. 2), cytochrome c release was markedly delayed, starting only after 12 h (Fig. 6D). These findings are compatible with JNK acting upstream of cytochrome c release. Notably, cytochrome c release was abrogated in the resistant 2008C13 cells, even at 24 h after CDDP treatment (Fig. 6D). Thus, it seems that the cells selected for resistance to CDDP have defects in the release of cytochrome c and in the resulting activation of downstream caspases.

Constitutive Activation of JNK/p38 MAPKs through MKK7/3 Sensitizes Chemoresistant 2008C13 Cells to CDDP-induced Apoptosis by Inducing FasL Expression—That the chemoresistant subclone 2008C13 showed only a transient phosphorylation of JNK and p38 in response to CDDP compared with the sustained JNK and p38 phosphorylation seen in the lysates of the chemosensitive 2008 cell line (Figs. 1 and 2) suggests that the duration of JNK and p38 phosphorylation seems to play an important role in the regulation of apoptosis in ovarian cancer cells because inhibition of the JNK and p38 kinases by SB202190 prevented CDDP-induced FasL expression (Figs. 4 and 5D). Possibly, the failure of the JNK and p38 kinases to be activated because of their inactivation by specific MAPK phosphatases that are induced in response to genotoxic stress (40) could underlie chemoresistance. We therefore examined whether selective reactivation of endogenous JNK and p38 induces or sensitizes the resistant subclone 2008C13 to apoptosis. To test this hypothesis, we used recombinant adenoviruses encoding constitutively active MKK7 (MKK7D) to selectively induce JNK activity (41, 42) or MKK3 (MKK3bE) to selectively induce p38 activity (43). The efficiency of the adenoviral gene delivery system in these ovarian cancer cells reached 90%, and the results showed increased phosphorylation of JNK and p38 after infection with Ad-MKK7D and Ad-MKK3bE, respectively, compared with infection with Ad-GFP (Fig. 7A). These data indicate that activated MKK7D and MKK3bE functioned as JNK- and p38-specific activators in ovarian tumor cells in the same fashion as previously demonstrated in other cell types (43). Moreover, FasL was up-regulated when MKK7D and MKK3bE were transduced, and FasL up-regulation correlated with the rate of apoptosis (Fig. 7, B and C). Control virus expressing GFP had no effect (Fig. 7, B and C). Taken together, these results suggest that activation of the JNK and p38 pathways through MKK7D and MKK3bE, respectively, is sufficient to induce expression of FasL and to promote apoptosis in resistant ovarian carcinoma cells. These data are consistent with our previous observations that failure to activate JNK and p38 led to a survival/chemoresistant phenotype and that persistent activation of JNK and p38 promoted apoptosis through FasL gene expression in the CDDP-sensitive 2008 ovarian carcinoma cell line (Figs. 2, 4, and 5).



View larger version (35K):
[in this window]
[in a new window]
 
FIG. 7.
Specific activation of JNK and p38 by MKK7 and MKK3 in ovarian carcinoma cells leads to FasL induction and induces 2008C13 apoptosis. CDDP-resistant 2008C13 ovarian carcinoma cells were infected with recombinant adenoviral vectors encoding activated MKK7 (Ad-MKK7D), MKK3 (Ad-MKK3bE), or GFP (Ad-GFP) as indicated. A, cell lysates were assayed for JNK or p38 activation by Western blot analysis. B, shown is nuclear fragmentation in 2008C13 cells after infection. Cells were fixed, and nuclear condensation was analyzed with DAPI. C, chemoresistant 2008C13 cells were treated as described for B; total RNA was extracted; and expression of FasL and {beta}-actin mRNAs was determined by semiquantitative reverse transcriptase-PCR.

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Primary and secondary resistance to chemotherapy is a central problem in cancer treatment (1, 4, 9). Resistance to chemotherapy may result from failure of the apoptotic pathways that are activated in response to drug treatment. Recent evidence indicates that the MAPK family protein kinases JNK and p38 are important mediators of apoptosis induced by stressful stimuli (25, 31, 39, 44, 45). The JNKs and p38 MAPKs are collectively termed stress-activated protein kinases because they are activated by a variety of stress-related stimuli (for review, see Refs. 10 and 11). The stress kinases are also activated by chemotherapy drugs, including paclitaxel, doxorubicin, vinblastine, and etoposide (46), and by certain DNA-damaging agents such as 1-D-arabinofuranosylcytosine, CDDP, and mitomycin C (24, 46, 47).

JNK activity parallels c-Jun phosphorylation in intact cells, suggesting that this protein kinase plays an important role in regulating c-Jun transcriptional activity. JNK activation results in the phosphorylation of transcription factors such as c-Jun and ATF-2, which then bind to AP-1-binding sites in the promoters of multiple target genes. JNK may contribute to death receptor transcription-dependent apoptotic signaling via c-Jun/AP-1, leading to transcriptional activation of FasL. JNK was initially thought to be a mediator of apoptosis in neuronal cells (44), and phosphorylation of c-Jun was shown to be essential for neuronal cell death induced by withdrawal of survival signals (31, 48, 49, 50).

The results described in this study map the early signaling events by which activation of JNK and p38 after CDDP treatment can lead to apoptosis in ovarian cancer cells. Previous studies have described the link between induction of JNK activation and apoptosis in response to cisplatin treatment in ovarian cells and other cell types (25, 26, 27). Those studies also suggested that the transcriptional activity of the c-Jun protein, which is increased by phosphorylation of c-Jun at Ser63 and Ser73 by JNK, is closely associated with apoptosis. However, none of those studies indicated a potential mechanism by which JNK activation, c-Jun phosphorylation, or both could trigger apoptosis.

Our findings demonstrate that prolonged phosphorylation (1 min to 12 h) of JNK and p38 MAPK, accompanied by c-Jun/ATF-2 phosphorylation, is an important step in the apoptotic signaling cascade induced by CDDP. More importantly, this sustained JNK activation and c-Jun phosphorylation paralleled the phosphorylation of p38 and ATF-2 (Fig. 2). These effects preceded and triggered up-regulation of FasL, which in turn contributed to the apoptotic response (Figs. 1, 2, 4, and 5). Thus, the duration of activated JNK and p38 signaling pathway is a critical factor in determining cell survival or apoptosis; transient activation was insufficient to induce death in CDDP-resistant cells, and prolonged JNK and p38 activation triggered cell death in CDDP-sensitive cells. Our findings indicate that resistance to CDDP in ovarian carcinoma cells is due in part to lack of prolonged activation of stress kinases and phosphorylation of c-Jun and ATF-2, a c-Jun dimerization partner. The transient activation observed in resistant cells seems to be insufficient to induce gene expression of a major initiator of apoptosis, FasL (Fig. 5). This differential response to CDDP between 2008 and 2008C13 cells may be due to differences in cellular uptake, and induced DNA damage (Table I) is unlikely because expression of FasL was uninduced in 2008C13 cells even after a 2-fold increase in the drug concentration (Fig. 5A).

We have also shown that inhibition of c-Jun activity, either by using a mutant defective in the JNK phosphoacceptor sites (c-Jun(S63A/S73A)) (Fig. 4C) or by inhibiting JNK activation with the small drug inhibitor SB202190 (Fig. 4, A and B), could block CDDP-induced apoptosis. Most importantly, this blockade correlated with the inability of c-Jun to activate FasL gene expression (Fig. 5D). In line with this evidence, others have reported that activation of JNK is required for an apoptotic response to alkylating agents (26, 27, 39, 51). Several studies have suggested that c-Jun is involved in genotoxin-induced apoptosis (52). One study demonstrated that a dominant-negative c-Jun mutant reduced apoptosis in human monoblastic leukemia cells after exposure to various DNA-damaging agents (53). Both c-jun/ fibroblasts and jnk1/jnk2/ double-knockout murine embryonic fibroblasts were found to be resistant to apoptosis induced by UV irradiation, anisomycin, and alkylating agents (39, 51, 52, 54), all of which may be mediated by the induction of FasL (52). Our data are also consistent with previous demonstrations that long-lasting activation of JNK and p38 kinase after withdrawal of survival factors or induction of MEKK1{Delta} results in enhanced c-Jun phosphorylation and in induction of FasL, leading to neuronal cell death (31, 50). Furthermore, the direct inhibition of JNK or c-Jun can block neuronal apoptosis induced by survival factor withdrawal (44, 55, 56). Most importantly, mice harboring a mutant allele of c-jun with Ser63 and Ser73 mutated to alanines are resistant to neuronal apoptosis induced by kainate (48). However, the normal physiological function of c-Jun or JNK, even in the context of a stress response, does not necessarily include induction of apoptosis (10, 11).

Tumor cells can inactivate pro-apoptotic cytokines such as Fas/FasL in a way that confers resistance to chemotherapy (8). Furthermore, it is most likely that upstream MAPKs are involved in this pathway because selective reactivation of JNK or p38 kinase by MKK7 or MKK3 induced apoptosis in the chemoresistant cells through transcriptional up-regulation of FasL expression (Fig. 7). Interestingly, activation of both MKK6 and p38 is required for {gamma}-irradiation-induced G2 arrest, and the expression of dominant-negative alleles of MKK6 or p38 allows cells to escape the DNA damage-induced G2 delay (57). Zanke et al. (25) reported, for instance, that cell lines defective in JNK activation are resistant to the lethal effects of CDDP. Moreover, either the expression of a dominant-negative SEK1/MKK4 mutant that blocks JNK activation or the transient expression of a dominant-negative JNK1 mutant is sufficient to confer resistance to apoptosis induced by several different stressful stimuli, including heat shock, UV irradiation, and a 2-h exposure to CDDP (25). These findings support the concept that the JNK pathway plays an important role in c-Jun-induced apoptosis. They also suggest that activation of the JNK pathway by diverse cell stress agents plays a critical part in mediating the toxicity of these treatments, including cell death. JNK activation in this context could broadly influence the cellular response of tumor cells to cytotoxic therapies.

In this study, we demonstrated that expression of FasL, the AP-1 target gene whose product can promote apoptosis, was highly induced by CDDP in 2008 cells and that this induction correlated with persistent activation of JNK and p38 and c-Jun phosphorylation. On the other hand, FasL expression was impaired in the CDDP-resistant variant 2008C13, even at a higher dose of CDDP (40 µM), and this impairment correlated with transient JNK phosphorylation (Figs. 2 and 5). The addition of a purified anti-Fas antibody resulted in significant and comparable apoptosis, indicating that the Fas-induced cell death pathway is functional in both cell types (Fig. 5C). Fas and FasL are a cognate receptor-ligand pair and play central roles in regulating programmed cell death. Interaction with FasL induces trimerization of the Fas receptor, leading to the recruitment of adaptor molecules such as the Fas-associated death domain protein, which directly binds and activates caspase-8, resulting in the induction of apoptosis. Activation of the Fas/FasL system is known to occur in a range of tumor cell lines after exposure to various types of anticancer drugs (32, 34). However, no clear explanation exists as to why some activators of AP-1 lead to FasL induction, whereas others do not. Kasibhatla et al. (33) initially reported that death ligands are subject to transcriptional regulation and that the FasL promoter is therefore directly activated by c-Jun through an AP-1-binding site in transient transfection experiments. Other studies have shown that withdrawal of survival factor or activation of the JNK pathway leads to an induction of apoptosis that is preceded by up-regulation of an immediate downstream target (FasL) in cerebellar granule neurons and PC12 cells (31, 33, 58). In addition, inhibition of the interaction of FasL with the Fas receptor leads to a reduction in apoptosis in response to genotoxic stress and growth factor withdrawal (31, 33, 52, 58). Consistent with the idea that FasL is a target in the JNK/c-Jun signaling pathway that induces apoptosis is the presence of AP-1-binding sites in the human FasL promoter region, which presumably contribute to the dependence of Fas/FasL interactions on c-Jun phosphorylation (31, 33, 58, 59). Indeed, several reports have identified AP-1 sites in the FasL promoter that are recognized by Jun/Fos or c-Jun/ATF-2 heterodimers. The presence of these sites is required for optimal responsiveness to such cellular stresses as exposure to UV and {gamma}-irradiation and alkylating agents (33, 52, 59, 60).

In ovarian tumors, FasL may be a pro-apoptotic target of JNK/AP-1 signaling because inhibition of JNK and p38 with SB202190 led to inhibition of the induction of FasL mRNA (Fig. 5D); and, conversely, reactivation of FasL induction by activated MKK7 or MKK3 (which induced persistent activation of JNK or p38, respectively) triggered cell death through FasL expression (Fig. 7). Interestingly, sustained suppression of Fas/FasL has been reported in CDDP-resistant cells (8), suggesting that the inability of these cells to up-regulate these receptors and ligands may be an important determinant of the ability of the cells to undergo apoptosis in response to chemotherapeutic agents. Clearly, c-Jun is required for FasL up-regulation because c-Jun-deficient fibroblasts, in contrast to wild-type cells, exhibited a defect in CDDP-induced apoptosis and because the inhibition of JNK by SB202190 significantly prevented CDDP-induced cell death in wild-type cells (Fig. 3A). In support of our findings is the fact that c-Jun-dependent FasL induction has been demonstrated in several systems in response to DNA-damaging agents, including the topoisomerase II inhibitors, UV irradiation, and the alkylating agent methyl methanesulfonate (21, 33, 52). Kolbus et al. (52) also showed that the resistance of c-jun/ fibroblasts to apoptosis is accompanied by impaired expression of FasL, providing evidence that c-Jun-dependent expression of FasL represents a rate-limiting step in the apoptosis induced by methyl methanesulfonate. Thus, lack of c-Jun activity increases survival against genotoxic stresses (52, 54), and reintroduction of the c-jun gene into c-jun/ fibroblast cells sensitizes cells to UV-induced apoptosis (54). The loss-of-function approach in fibroblasts allowed the identification and dissection of c-Jun-dependent and c-Jun-independent processes upstream and downstream of Fas activation. Once activated, Fas-induced death signaling is not affected by the loss of c-Jun or JNK activation, demonstrating that only the initiation and not the execution of stress-induced apoptosis depends on c-Jun (Fig. 5E) (52). These data strongly suggest that one mechanism underlying chemoresistance might be the inability to sustain activation of stress kinases and therefore to enable up-regulation of FasL, the downstream target death gene.

Downstream consequences of Fas/FasL interaction are complex and depend in part on the cell type being studied. Some studies implicate JNK in apoptosis, and others describe a lack of correlation between JNK and cell death or even interference of JNK activation with apoptosis (for reviews, see Refs. 10 and 11). The present study demonstrated that JNK and p38 activation is not due to a downstream FasL signaling event because JNK and p38 activation occurred during the first hour (i.e. before FasL was expressed 6–18 h later) (Figs. 2 and 5). Moreover, the caspase inhibitor Z-VAD-fmk did not prevent CDDP-induced FasL (Fig. 6) or JNK and p38 activation (data not shown), indicating that FasL expression is an event down-stream of the JNK/p38 pathway. Also in support of our findings are reports that treatment with recombinant FasL in wild-type and c-jun/ 3T3 cells does not result in any detectable JNK activity (data not shown) (52). However, in the studies of Kolbus et al. (52), both cell lines responded to methyl methanesulfonate by showing activation of kinase activities within the first hour, and no additional increase in JNK activity was observed at later times, when FasL induction reached maximal levels and apoptosis became detectable. Clearly, neither c-Jun nor JNK is required for the expression and activity of cellular components located downstream of Fas because c-jun/ cells and jnk1/jnk2/ mouse embryonic fibroblasts are sensitive to FasL-induced apoptosis (39, 52).

The molecular pathways triggered by anticancer drugs that lead to the activation of stress pathways are not well understood. Exactly how cisplatin triggers stress kinase pathways is not yet known, nor are the sequential events between CDDP-induced oxidative stress, DNA damage, JNK/p38 activation, and apoptosis. We have also reported that cisplatin resistance in ovarian carcinoma is associated with a defect in apoptosis through XIAP (X chromosome-linked inhibitor of apoptotic) regulation (61). Based on our data and literature reports, several hypotheses can be proposed. It is clear that powerful pro-oxidants that cause generation of reactive oxygen species and free radicals are generated in response to non-redox-active initiators of apoptosis such as cisplatin. In addition, studies have shown that pretreatment of cells with the antioxidants glutathione and N-acetylcysteine effectively blocks CDDP-induced apoptosis and CDDP-induced activation of JNK and p38 (57, 62, 63). Finally, another possibility in the control of stress kinase activities in response to CDDP may relate to interference with phosphatases (64). MAPK phosphatases play an important role in selectively regulating the duration of JNK and p38 phosphorylation and dephosphorylation (65) and are activated by various stresses that activate JNK. All known phosphotyrosine and threonine phosphatases, including the dual-specificity phosphatases, contain an essential catalytic cysteinyl residue (66) that is sensitive to thio(SH)-reactive agents such as CDDP. Therefore, oxidative stress generated by CDDP may not only deplete reduced glutathione and other antioxidant molecules, but may also cause the oxidation of the sulfhydryl groups on these phosphatases, leading to their inactivation (51, 67). Our study shows that the sensitization of carcinoma cells to genotoxic stress is largely due to potentiation of the JNK and p38 pathways. Indeed, the CDDP-resistant 2008C13 cells exhibited a defect in the activation of JNK that may contribute to the resistance of advanced tumors to cancer therapy. This suppression of stress kinase activation supports the possible role of an alteration of phosphatase activities in CDDP-resistant cells. Thus, the specific induction of MAPK phosphatases by these agents seems to be responsible for protecting cells from apoptosis by preventing prolonged activation of JNK/p38 kinases (51, 67). The duration of JNK/p38 activation could thus be regulated by MAPK phosphatases through a feedback mechanism. Whether CDDP inactivates JNK in resistant cell lines via stimulation of MAPK phosphatases is under investigation by our group.

In conclusion, we have demonstrated that an early key determinant of CDDP-induced apoptosis is the duration of JNK and p38 phosphorylation. Prolonged JNK and p38 activation results in phosphorylation of the AP-1 target transcription factors c-Jun and ATF-2, thus promoting the expression of FasL and the binding of FasL to the Fas receptor, which leads to cell death (Fig. 8). Because stress kinase pathways remain potentiated in CDDP-sensitive cells, the cells remain sensitive to stress signals such as genotoxic agents. These findings raise the possibility that defects in this cascade may contribute to failure of chemotherapy-induced apoptosis. Therefore, modulation of apoptotic pathways through the MAPK signaling cascade may become a therapeutic goal for the prevention and treatment of cancer.



View larger version (24K):
[in this window]
[in a new window]
 
FIG. 8.
Proposed model of CDDP-induced signaling pathways leading to JNK activation and apoptosis. Exposure of ovarian carcinoma cells to CDDP caused activation and potent phosphorylation of JNK/p38, resulting in phosphorylation of c-Jun and ATF-2. In the CDDP-sensitive cell line 2008, the persistence of this activation led to the activation of FasL, the AP-1 target gene. Release of FasL triggered apoptosis through the Fas receptor, which activates specific caspases, and key substrates were cleaved. Protection against cell death was conferred either by overexpression of a c-Jun mutant lacking the JNK phosphoacceptor sites or by a chemical inhibitor (SB202190) of p38 and JNK that inhibits FasL induction. Inhibition of Fas/FasL interaction through a neutralizing anti-FasL antibody (NOK-2) protected the chemosensitive 2008 cells from apoptosis. In the CDDP-resistant 2008C13 cells, the protection against CDDP-induced apoptosis was presumably due to rapid inactivation of the JNK/p38 activation of MAPK phosphatases (MKPs).

 


    FOOTNOTES
 
* This work was supported in part the University of Texas M. D. Anderson Cancer Center, National Institutes of Health Grants P30CA16672-24 and 5P50CA83639 and Core Grant CA16672, and the Ovarian Cancer Research Program of the United States Department of Defense (to F. X. C.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ These authors contributed equally to this work. Back

Recipient of fellowships from the Association pour la Recherche sur le Cancer and the Fondation pour la Recherche Medicale. Back

{ddagger}{ddagger} To whom correspondence should be addressed: Dept. of Molecular Therapeutics, University of Texas M. D. Anderson Cancer Center, 1515 Holcombe Blvd., P. O. Box 317, Houston, TX 77030. Tel.: 713-792-6306; Fax: 713-792-4005; E-mail: fxclaret{at}mdanderson.org.

1 The abbreviations used are: CDDP, cisplatin; MAPK, mitogen-activated protein kinase; ERK, extracellular signal-regulated kinase; JNK, c-Jun N-terminal kinase; MEK, MAPK/ERK kinase; FasL, Fas ligand; ATF-2, activating transcription factor-2; PARP, poly(ADP-ribose) polymerase; Z-VAD-fmk, benzyloxycarbonyl-Val-Ala-Asp-(O-methyl)fluoromethyl ketone; Ad, adenovirus; GFP, green fluorescent protein; MKK, mitogen-activated protein kinase kinase; PBS, phosphate-buffered saline; GST, glutathione S-transferase; DAPI, 4,6-diamidino-2-phenylindole; HA, hemagglutinin; MEKK, MEK kinase. Back


    ACKNOWLEDGMENTS
 
We are grateful to S. B. Howell, S. G. Chaney, and Z. H. Siddik for kindly providing the 2008 and 2008C13 cells and for sharing data; to M. Karin and E. F. Wagner for the gift of c-jun/ and c-jun/ fibroblasts and plasmids; and to N. Hollbrook and O. Potapova for sharing unpublished data.



    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Reed, J. C. (1999) Curr. Opin. Oncol. 11, 68–75[CrossRef][Medline] [Order article via Infotrieve]
  2. Kelland, L. R., and Farreell, N. P. (2000) Platimium-based Drugs in Cancer Therapy, Humana Press Inc., Totowa, NJ
  3. Lippert, B. (1999) Cisplatin: Chemistry and Biochemistry of a Leading Anticancer Drug, Wiley-VCH, New York
  4. Andrews, P. A., and Howell, S. B. (1990) Cancer Cells 2, 35–43[Medline] [Order article via Infotrieve]
  5. Kelley, S. L., Basu, A., Teicher, B. A., Hacker, M. P., Hamer, D. H., and Lazo, J. S. (1988) Science 241, 1813–1815[Medline] [Order article via Infotrieve]
  6. Cohen, S. M., and Lippard, S. J. (2001) Prog. Nucleic Acids Res. Mol. Biol. 67, 93–130[Medline] [Order article via Infotrieve]
  7. Evan, G., and Littlewood, T. (1998) Science 281, 1317–1322[Abstract/Free Full Text]
  8. Herr, I., and Debatin, K. M. (2001) Blood 98, 2603–2614[Abstract/Free Full Text]
  9. Niedner, H., Christen, R., Lin, X., Kondo, A., and Howell, S. B. (2001) Mol. Pharmacol. 60, 1153–1160[Abstract/Free Full Text]
  10. Chang, L., and Karin, M. (2001) Nature 410, 37–40[CrossRef][Medline] [Order article via Infotrieve]
  11. Davis, R. J. (2000) Cell 103, 239–252[Medline] [Order article via Infotrieve]
  12. Cobb, M. H. (1999) Prog. Biophys. Mol. Biol. 71, 479–500[CrossRef][Medline] [Order article via Infotrieve]
  13. Treisman, R. (1996) Curr. Opin. Cell Biol. 8, 205–215[CrossRef][Medline] [Order article via Infotrieve]
  14. Holmstrom, T. H., Schmitz, I., Soderstrom, T. S., Poukkula, M., Johnson, V. L., Chow, S. C., Krammer, P. H., and Eriksson, J. E. (2000) EMBO J. 19, 5418–5428[Abstract/Free Full Text]
  15. Andrews, P. A., Velury, S., Mann, S. C., and Howell, S. B. (1988) Cancer Res. 48, 68–73[Abstract]
  16. Delmastro, D. A., Li, J., Vaisman, A., Solle, M., and Chaney, S. G. (1997) Cancer Chemother. Pharmacol. 39, 245–253[CrossRef][Medline] [Order article via Infotrieve]
  17. Wang, Y., Su, B., Sah, V. P., Brown, J. H., Han, J., and Chien, K. R. (1998) J. Biol. Chem. 273, 5423–5426[Abstract/Free Full Text]
  18. Siddik, Z. H., Boxall, F. E., and Harrap, K. R. (1987) Anal. Biochem. 163, 21–26[Medline] [Order article via Infotrieve]
  19. Yoshida, M., Khokhar, A. R., and Siddik, Z. H. (1994) Cancer Res. 54, 3468–3473[Abstract]
  20. Hibi, M., Lin, A., Smeal, T., Minden, A., and Karin, M. (1993) Genes Dev. 7, 2135–2148[Abstract]
  21. Eichhorst, S. T., Muller, M., Li-Weber, M., Schulze-Bergkamen, H., Angel, P., and Krammer, P. H. (2000) Mol. Cell. Biol. 20, 7826–7837[Abstract/Free Full Text]
  22. Bossy-Wetzel, E., Newmeyer, D. D., and Green, D. R. (1998) EMBO J. 17, 37–49[Abstract/Free Full Text]
  23. Pandey, P., Raingeaud, J., Kaneki, M., Weichselbaum, R., Davis, R. J., Kufe, D., and Kharbanda, S. (1996) J. Biol. Chem. 271, 23775–23779[Abstract/Free Full Text]
  24. Liu, Z. G., Baskaran, R., Lea-Chou, E. T., Wood, L. D., Chen, Y., Karin, M., and Wang, J. Y. (1996) Nature 384, 273–276[CrossRef][Medline] [Order article via Infotrieve]
  25. Zanke, B. W., Boudreau, K., Rubie, E., Winnett, E., Tibbles, L. A., Zon, L., Kyriakis, J., Liu, F. F., and Woodgett, J. R. (1996) Curr. Biol. 6, 606–613[Medline] [Order article via Infotrieve]
  26. Sanchez-Perez, I., Murguia, J. R., and Perona, R. (1998) Oncogene 16, 533–540[CrossRef][Medline] [Order article via Infotrieve]
  27. Potapova, O., Haghighi, A., Bost, F., Liu, C., Birrer, M. J., Gjerset, R., and Mercola, D. (1997) J. Biol. Chem. 272, 14041–14044[Abstract/Free Full Text]
  28. Hilberg, F., Aguzzi, A., Howells, N., and Wagner, E. F. (1993) Nature 365, 179–181[CrossRef][Medline] [Order article via Infotrieve]
  29. Jacinto, E., Werlen, G., and Karin, M. (1998) Immunity 8, 31–41[Medline] [Order article via Infotrieve]
  30. Whitmarsh, A. J., Yang, S. H., Su, M. S., Sharrocks, A. D., and Davis, R. J. (1997) Mol. Cell. Biol. 17, 2360–2371[Abstract]
  31. Le-Niculescu, H., Bonfoco, E., Kasuya, Y., Claret, F. X., Green, D. R., and Karin, M. (1999) Mol. Cell. Biol. 19, 751–763[Abstract/Free Full Text]
  32. Friesen, C., Herr, I., Krammer, P. H., and Debatin, K. M. (1996) Nat. Med. 2, 574–577[Medline] [Order article via Infotrieve]
  33. Kasibhatla, S., Brunner, T., Genestier, L., Echeverri, F., Mahboubi, A., and Green, D. R. (1998) Mol. Cell 1, 543–551[Medline] [Order article via Infotrieve]
  34. Fulda, S., Strauss, G., Meyer, E., and Debatin, K. M. (2000) Blood 95, 301–308[Abstract/Free Full Text]
  35. Liu, X., Kim, C. N., Yang, J., Jemmerson, R., and Wang, X. (1996) Cell 86, 147–157[Medline] [Order article via Infotrieve]
  36. Li, P., Nijhawan, D., Budihardjo, I., Srinivasula, S. M., Ahmad, M., Alnemri, E. S., and Wang, X. (1997) Cell 91, 479–489[Medline] [Order article via Infotrieve]
  37. Zou, H., Henzel, W. J., Liu, X., Lutschg, A., and Wang, X. (1997) Cell 90, 405–413[Medline] [Order article via Infotrieve]
  38. Srinivasula, S. M., Ahmad, M., Fernandes-Alnemri, T., and Alnemri, E. S. (1998) Mol. Cell 1, 949–957[Medline] [Order article via Infotrieve]
  39. Tournier, C., Hess, P., Yang, D. D., Xu, J., Turner, T. K., Nimnual, A., Bar-Sagi, D., Jones, S. N., Flavell, R. A., and Davis, R. J. (2000) Science 288, 870–874[Abstract/Free Full Text]
  40. Keyse, S. M. (2000) Curr. Opin. Cell Biol. 12, 186–192[CrossRef][Medline] [Order article via Infotrieve]
  41. Tournier, C., Whitmarsh, A. J., Cavanagh, J., Barrett, T., and Davis, R. J. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 7337–7342[Abstract/Free Full Text]
  42. Wu, Z., Wu, J., Jacinto, E., and Karin, M. (1997) Mol. Cell. Biol. 17, 7407–7416[Abstract]
  43. Raingeaud, J., Whitmarsh, A. J., Barrett, T., Derijard, B., and Davis, R. J. (1996) Mol. Cell. Biol. 16, 1247–1255[Abstract]
  44. Xia, Z., Dickens, M., Raingeaud, J., Davis, R. J., and Greenberg, M. E. (1995) Science 270, 1326–1331[Abstract]
  45. Verheij, M., Bose, R., Lin, X. H., Yao, B., Jarvis, W. D., Grant, S., Birrer, M. J., Szabo, E., Zon, L. I., Kyriakis, J. M., Haimovitz-Friedman, A., Fuks, Z., and Kolesnick, R. N. (1996) Nature 380, 75–79[CrossRef][Medline] [Order article via Infotrieve]
  46. Osborn, M. T., and Chambers, T. C. (1996) J. Biol. Chem. 271, 30950–30955[Abstract/Free Full Text]
  47. Kharbanda, S., Ren, R., Pandey, P., Shafman, T. D., Feller, S. M., Weichselbaum, R. R., and Kufe, D. W. (1995) Nature 376, 785–788[CrossRef][Medline] [Order article via Infotrieve]
  48. Behrens, A., Sibilia, M., and Wagner, E. F. (1999) Nat. Genet. 21, 326–329[CrossRef][Medline] [Order article via Infotrieve]
  49. Watson, A., Eilers, A., Lallemand, D., Kyriakis, J., Rubin, L. L., and Ham, J. (1998) J. Neurosci. 18, 751–762[Abstract/Free Full Text]
  50. Herdegen, T., Claret, F. X., Kallunki, T., Martin-Villalba, A., Winter, C., Hunter, T., and Karin, M. (1998) J. Neurosci. 18, 5124–5135[Abstract/Free Full Text]
  51. Sanchez-Perez, I., and Perona, R. (1999) FEBS Lett. 453, 151–158[CrossRef][Medline] [Order article via Infotrieve]
  52. Kolbus, A., Herr, I., Schreiber, M., Debatin, K. M., Wagner, E. F., and Angel, P. (2000) Mol. Cell. Biol. 20, 575–582[Abstract/Free Full Text]
  53. Stadheim, T. A., and Kucera, G. L. (2002) Leuk. Res. 26, 55–65[CrossRef][Medline] [Order article via Infotrieve]
  54. Shaulian, E., Schreiber, M., Piu, F., Beeche, M., Wagner, E. F., and Karin, M. (2000) Cell 103, 897–907[Medline] [Order article via Infotrieve]
  55. Ham, J., Babij, C., Whitfield, J., Pfarr, C. M., Lallemand, D., Yaniv, M., and Rubin, L. L. (1995) Neuron 14, 927–939[Medline] [Order article via Infotrieve]
  56. Whitfield, J., Neame, S. J., Paquet, L., Bernard, O., and Ham, J. (2001) Neuron 29, 629–643[Medline] [Order article via Infotrieve]
  57. Wang, X., McGowan, C. H., Zhao, M., He, L., Downey, J. S., Fearns, C., Wang, Y., Huang, S., and Han, J. (2000) Mol. Cell. Biol. 20, 4543–4552[Abstract/Free Full Text]
  58. Faris, M., Kokot, N., Latinis, K., Kasibhatla, S., Green, D. R., Koretzky, G. A., and Nel, A. (1998) J. Immunol. 160, 134–144[Abstract/Free Full Text]
  59. Faris, M., Latinis, K. M., Kempiak, S. J., Koretzky, G. A., and Nel, A. (1998) Mol. Cell. Biol. 18, 5414–5424[Abstract/Free Full Text]
  60. Eichhorst, S. T., Muerkoster, S., Weigand, M. A., and Krammer, P. H. (2001) Cancer Res. 61, 243–248[Abstract/Free Full Text]
  61. Mansouri, A., Zhang, Q., Ridgeway, L., Tian, L., and Claret, F. X. (2003) Oncol. Res., in press
  62. Benhar, M., Dalyot, I., Engelberg, D., and Levitzki, A. (2001) Mol. Cell. Biol. 21, 6913–6926[Abstract/Free Full Text]
  63. Adler, V., Yin, Z., Fuchs, S. Y., Benezra, M., Rosario, L., Tew, K. D., Pincus, M. R., Sardana, M., Henderson, C. J., Wolf, C. R., Davis, R. J., and Ronai, Z. (1999) EMBO J. 18, 1321–1334[Abstract/Free Full Text]
  64. Sanchez-Perez, I., Martinez-Gomariz, M., Williams, D., Keyse, S. M., and Perona, R. (2000) Oncogene 19, 5142–5152[CrossRef][Medline] [Order article via Infotrieve]
  65. Camps, M., Nichols, A., and Arkinstall, S. (2000) FASEB J. 14, 6–16[Abstract/Free Full Text]
  66. Neel, B. G., and Tonks, N. K. (1997) Curr. Opin. Cell Biol. 9, 193–204[CrossRef][Medline] [Order article via Infotrieve]
  67. Palacios, C., Collins, M. K., and Perkins, G. R. (2001) Curr. Biol. 11, 1439–1443[CrossRef][Medline] [Order article via Infotrieve]