ToxSci Advance Access originally published online on May 3, 2006
Toxicological Sciences 2006 92(2):433-444; doi:10.1093/toxsci/kfl003
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Activation of Multiple Mitogen-Activated Protein Kinases in Pro/Pre–B Cells by GW7845, a Peroxisome Proliferator–Activated Receptor 
Agonist, and Their Contribution to GW7845-Induced Apoptosis
* Department of Environmental Health, Boston University School of Public Health, Boston, Massachusetts 02118; and Department of Microbiology, Boston University School of Medicine, Boston, Massachusetts 02118
1 To whom correspondence should be addressed at Department of Environmental Health, Boston University School of Public Health, 715 Albany Street, R-405, Boston, MA 2118. Fax: (617) 638-6463. E-mail: jschlezi{at}bu.edu.
Received January 17, 2006; accepted April 26, 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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There is growing interest in using peroxisome proliferator–activated receptor (PPAR)
agonists as chemotherapeutic agents in hematologic malignancies. PPAR agonists of diverse chemical structure induce apoptosis in several malignant B cell lines. However, PPAR agonists also induce apoptosis in normal B cells. One such agonist, GW7845, rapidly induces apoptosis in early B cells. Understanding the mechanisms of PPAR agonist–induced death is essential to minimizing loss of normal cells during chemotherapy. PPAR agonists influence mitogen-activated protein kinase (MAPK) cascades in other systems, and MAPKs can be associated with apoptosis. Therefore, we investigated the activation of MAPKs in primary pro–B cells and cultured pro/pre–B cells and their role in GW7845-induced apoptosis. Treatment of a nontransformed murine pro/pre–B-cell line with GW7845 transiently induced the phosphorylation of extracellular signal–related protein kinase (ERK) 1/2, but strongly and persistently induced the activation of p38 MAPK and c-Jun NH2-terminal kinase (JNK). In primary pro–B-cells, p38 MAPK and JNK were activated following treatment with GW7845. Phosphorylation of activating transcription factor-2 (ATF-2) was induced strongly in both B-cell types. In pro/pre–B cells, pretreatment with the p38 MAPK/JNK inhibitor PD169316 potently suppressed multiple facets of GW7845-induced apoptosis signaling. However, when a series of p38 MAPK and JNK inhibitors were used, only SB202190, also a dual inhibitor, completely suppressed GW7845-induced apoptosis. Inhibitors specific for p38 MAPK and JNK were only partially effective, suggesting that suppression of a single MAPK is not sufficient to inhibit death. The results support the hypothesis that GW7845 initiates an apoptotic pathway in early B cells through the activation of a kinase cascade that includes at least p38 MAPK and JNK.
Key Words: bone marrow; B cells; PPAR; apoptosis; MAPK.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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There is growing interest in the use of peroxisome proliferator–activated receptor (PPAR)
agonists as chemotherapeutic agents in hematologic malignancies (Konopleva and Andreeff, 2002
). PPAR agonists of diverse chemical structure induce apoptosis in several leukemia, lymphoma, and myeloma cell lines (Contractor et al., 2005; Konopleva et al., 2004; Piva et al., 2005; Ray et al., 2004). However, PPAR agonists also induce apoptosis in normal B cells (Padilla et al., 2000; Schlezinger et al., 2002, 2004), a result which could limit the use of PPAR agonists as chemotherapeutics. Furthermore, some ubiquitous environmental pollutants similarly activate PPAR and induce death in primary pro– and cultured pro/pre–B cells (Schlezinger et al., 2004), suggesting an inappropriate disruption of the normal development of the B cell repertoire by PPAR agonists.
Apoptosis is a critical event during the early stages of B lymphopoiesis and occurs at the highest rate during the pro/pre–B cell transition and during clonal deletion (Lu and Osmond, 2000), potentially sensitizing B cells at these developmental stages to apoptotic agents. Indeed, treatment of murine primary bone marrow pro–B cells or a nontransformed murine pro/pre–B-cell line with PPAR agonists including the endogenous prostaglandin, 15-deoxy-
12,14-prostaglandin J2 (15d-PGJ2) (Schlezinger et al., 2004), the synthetic antidiabetic drugs ciglitazone and GW7845 (Schlezinger et al., 2002), or the environmental contaminant mono-2-ethylhexyl phthalate (Schlezinger et al., 2004) results in apoptosis. However, studies have only begun to define the mechanisms through which PPAR agonists effect apoptosis in malignant or normal B cells.
PPAR is highly expressed in tissues of the immune system (Braissant et al., 1996) and participates in control of B cell proliferation and antigen-specific responses (Setoguchi et al., 2001). PPAR is a member of a nuclear hormone receptor superfamily comprised of three subtypes, 
, 
, and . Following ligand binding, PPAR forms a heterodimeric complex with the retinoid X receptor (RXR), initiating a conformational change that results in the dissociation of corepressors, the association of coactivators, and receptor complex binding to PPAR response elements in PPAR-responsive genes (Willson et al., 2000). Transcriptional activity may be enhanced in the presence of ligands for both PPAR and RXR (Kliewer et al., 1992). A contribution of PPAR to apoptosis is supported by the facts that PPAR antagonists reduce the magnitude of apoptosis in multiple cell types induced by ring-substituted diindolylmethanes, 2-cyano-3,12-dioxoolean-1,9-dien-28-oic acid (CDDO), 15d-PGJ2, glitazone drugs, and mono-2-ethylhexyl phthalate (Chen et al., 2005; Contractor et al., 2005; Kim et al., 2003; Shan et al., 2004; Yokoyama et al., 2003) and that cotreatment of PPAR agonists with 9-cis-retinoic acid, an RXR agonist, synergistically increases the incidence of apoptosis (Contractor et al., 2005; Konopleva et al., 2004; Ray et al., 2004; Schlezinger et al., 2002, 2004).
In addition to activating PPAR and inducing apoptosis, PPAR agonists also appear to share the ability to activate mitogen-activated protein kinase (MAPK) cascades (Bae and Song, 2003; Gardner et al., 2005; Kim et al., 2003; Lennon et al., 2002; Rokos and Ledwith, 1997; Shan et al., 2004; Takeda et al., 2001; Teruel et al., 2003). There are three major groups of MAPKs in mammalian cells, the extracellular signal–related protein kinases (ERKs), the c-Jun NH2-terminal kinases (JNKs), and p38 MAPK, that are activated by phosphorylation within a three-tiered module of a MAP kinase kinase kinase, a MAP kinase kinase, and a MAPK. MAPK cascades transduce signals from the cell surface into changes in gene expression that control diverse functions such as proliferation, differentiation, transformation, and apoptosis. In particular, p38 MAPK and JNK activation have been associated with apoptosis (as reviewed in Wada and Penninger, 2004).
The mechanism of PPAR agonist–induced apoptosis in nontransformed, bone marrow B cells remains unknown; therefore, we sought to investigate the role of MAPKs in PPAR agonist–induced apoptosis using GW7845, the PPAR agonist that induces the strongest apoptotic signal in primary pro– and cultured pro/pre–B cells (Schlezinger et al., 2002, 2004). GW7845 was optimized based on its activity toward PPAR, resulting in transactivation activities in the nanomolar range and a 1000-fold selectivity for PPAR over PPAR (Willson et al., 2000). The data show that GW7845 induces the activation of stress kinases in primary pro– and cultured pro/pre–B cells and that p38 MAPK and JNK contribute to PPAR agonist–induced apoptosis in cultured pro/pre–B cells. Defining the mechanism of PPAR agonist–induced apoptosis may aid in determining the therapeutic window in which normal B cell apoptosis is minimized while malignant B cell apoptosis is maximized and may explain how some environmental chemicals suppress immune cell development.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Materials.
SB202190, SB203580, SP600125, and U0216 were from Biomol (Plymouth Meeting, PA). PD169316 was from Calbiochem (San Diego, CA). Antibodies specific for cleaved caspase-3 and specific for kinases and in vitro kinase assay kits were from Cell Signaling Technology (Beverly, MA). The cytochrome c–specific antibody was from BD Biosciences Clontech (Palo Alto, CA). GW7845 was the generous gift of Dr T. Willson (GlaxoSmithKline, Research Triangle Park, NC). Plasmocin was from Invivogen (San Diego, CA). JC-1 was from Molecular Probes (Eugene, OR). Murine rIL-7 was from Research Diagnostics (Flanders, NJ). Propidium iodide (PI), protease inhibitor cocktail for mammalian cells, and the ß-actin antibody were from Sigma Chemical Co (St Louis, MO). All other reagents were from Fisher Scientific (Suanee, GA).
Cell cultures.
The stromal cell–dependent, C57BL/6-derived BU-11 cell line has been characterized previously (Ryu et al., 2005; Schlezinger et al., 2002, 2004). BU-11 cells represent B cells at the transition between the pro– and early pre–B cell stages as they are CD43+/B220+/IgM– with rearranged Ig heavy-chain genes. BMS2 is a culture dish–adherent, cloned bone marrow stromal cell line that supports BU-11 cell growth. Stocks of BU-11 cells were maintained on BMS2 cell monolayers in an equal mixture of DMEM and RPMI 1640 medium with 5% fetal bovine serum (FBS), plasmocin, L-glutamine, and 2-mercaptoethanol. All cultures were maintained at 37°C in a humidified, 7.5% CO2 atmosphere. Cell cultures were determined to be mycoplasma negative (MycoAlert Mycoplasma Detection Kit; Cambrex, East Rutherford, NJ).
Primary bone marrow pro–B cell cultures were prepared from wild-type C57BL/6 mice (Jackson Laboratories, Bar Harbor, ME) as described previously (Ryu et al., 2005). Bone marrow was flushed from the femurs of 4- to 6-week-old male mice. Red blood cells were lysed by incubation in 0.17M NH4Cl, 10mM KHCO3, and 1mM EDTA at 37°C for 5 min. The remaining cells were cultured for 5–7 days in primary B cell medium (RPMI containing 10% FBS, penicillin/streptomycin/amphotericin B, L-glutamine, 2-mercaptoethanol, and 16 ng/ml murine rIL-7 [RDI, Flanders, NJ]). A pro–B cell phenotype was confirmed by staining with anti-B220–specific and anti-CD43–specific antibodies. At least 95% of the cells express CD43 and B220.
For experiments, pro/pre–B cells were cultured in 24-well plates (4 x 105 cells in 1 ml of medium) or in T25 flasks (6 x 106 cells in 10 ml medium) overnight in RPMI with 5% FBS and treated with Vh (ethanol:DMSO, 50:50, 0.1% final concentration) or GW7845 (40µM) for 15 min to 2 h. Cells were pretreated with Vh (DMSO, 0.1% final concentration), PD169316 (1–5µM), SB202190 (20–40µM), SB230580 (20–40µM), SP600125 (5–10µM), and/or U0126 (20µM) for 30 min. Primary pro–B cells (107) were cultured in T25 flasks in 10 ml of primary B cell medium with 7.5% FBS overnight and treated with Vh (ethanol:DMSO, 50:50, 0.1%) or GW7845 (80µM) for 30 min to 3 h.
Analysis of apoptosis.
For PI staining, cells were harvested into cold PBS containing 5% FBS and 10µM azide. Cells were resuspended in 0.25 ml of hypotonic buffer containing 50 µg/ml PI, 1% sodium citrate, and 0.1% Triton X-100 and analyzed with FL-2 in the log mode on a Becton Dickinson FACScan flow cytometer. The percentage of cells undergoing apoptosis was determined to be those having a weaker PI fluorescence than cells in the G0/G1 phase of the cell cycle.
For determination of DNA fragmentation, cells were harvested into ice-cold PBS. DNA was isolated and processed as previously described (Schlezinger et al., 2002). DNA (3 µg) was treated with 10 µg/ml RNase A at 37°C for 10 min prior to separation on a 1.5% agarose gel. DNA was visualized using ethidium bromide staining.
Kinase activation and substrate phosphorylation.
Cells were harvested into ice-cold PBS. Cytoplasmic extracts were prepared as described previously (Schlezinger et al., 2002). Whole-cell extracts were prepared using lysis buffer from Cell Signaling Technology. Protein concentrations were determined by the Bradford method. Proteins (40–80 µg) were resolved on 10 or 12% gels, transferred to a 0.2-µm nitrocellulose membrane, and incubated with phospho-ERK1/2 (Thr202/Tyr204)–specific rabbit monoclonal antibody (4377), phospho-p38 MAPK (Thr180/Tyr182)–specific rabbit monoclonal antibody (9215), phospho-stress activated protein kinase (SAPK)/JNK (Thr183/Tyr185)–specific mouse monoclonal antibody (9255), phospho-ATF-2 (Thr71)–specific rabbit monoclonal antibody (5112), or MAPKAP-K2–specific rabbit polyclonal antibody (3042). The secondary antibodies were horseradish peroxidase–linked goat anti-rabbit or goat anti-mouse IgG (BioRad, Hercules, CA). Immunoreactive bands were visualized with enhanced chemiluminescence. To control for equal protein loading, blots were reprobed with antibodies to total ERK1/2 (9102), p38 MAPK (9212), SAPK/JNK (9252), ATF-2 (9226), and/or ß-actin (A5441) and analyzed as above. To quantify changes in protein expression, band densities were determined using the UVP Bioimaging System and the Labworks 4 program (UVP, Inc., Upland, CA). The band density of the phosphorylated protein was divided by the band density of the total protein. To normalize for differences between experiments, the phosphorylated/total ratio for experimental samples then was divided by the phospho/total ratio in the naive sample from that experiment.
In vitro kinase assay.
Kinase activity was determined by in vitro kinase assay using the nonradioactive ERK1/2 (9800), p38 MAPK (9820), and SAPK/JNK (9810) kinase activity kits. Briefly, cell extracts were prepared, and protein concentrations were determined by the Bradford assay. Extracted protein (200 µg) was incubated in a final volume of 250 µl of lysis buffer in the presence of immobilized phospho-ERK1/2 (Thr202/Tyr204) or phospho-p38 MAPK (Thr180/Tyr184)–specific antibody or immobilized c-jun fusion protein at 4°C overnight. The beads were washed four times and incubated in kinase buffer in the presence of substrate (2 µg) (Elk-1, ATF-2, or c-jun) and 200µM ATP at 30°C for 30 min. The reaction was terminated with excess SDS sample buffer. Following boiling for 5 min, the reaction mixtures were resolved on 12% gels, transferred to a 0.2-µm nitrocellulose membrane, and incubated with phospho-Elk-1 (Ser383)–, phospho-ATF-2 (Thr71)–, or phospho-c-jun (Ser63)–specific rabbit polyclonal antibody. Blots were developed and analyzed as described above. Analysis of total ERK1/2, p38 MAPK, or JNK was performed to determine that the expression of the proteins did not change over time.
Analysis of mitochondrial membrane potential (
m).
JC-1 (1.4µM) was added to each well at the time of dosing. At the time of harvest, B cells were transferred to FACS tubes without washing and analyzed immediately for green and red fluorescence by flow cytometry. Only cells in the live gate were analyzed. Cells with low mitochondrial membrane potential (
) were determined to be those having an increased green fluorescence with or without a loss of red fluorescence.
Cytochrome c release.
Cells were harvested into ice-cold PBS. Cell pellets were resuspended immediately in permeabilization buffer (10mM HEPES, pH 7.4, 210mM mannitol, 70mM sucrose, 5mM succinate, 0.2mM EGTA) containing 1.4 µl/ml of a 10% digitonin solution in DMSO. Following a 5-min incubation on ice, the same volume of permeabilization buffer without digitonin was added. The mixture was vortexed briefly and then centrifuged at 14,000 x g for 30 min. The supernatant was used to determine cytochrome c release. Protein concentrations were determined by the Bradford method. Proteins (5 µg) were resolved on 15% gels, transferred to a 0.2-µm nitrocellulose membrane, and incubated with a cytochrome c–specific rabbit polyclonal antibody (S2050). Blots were developed and analyzed as described above. Blots were reprobed with a ß-actin–specific mouse monoclonal antibody to determine equal protein loading.
Caspase activation.
Cells were harvested into ice-cold PBS. Cytoplasmic extracts were prepared as described previously (Schlezinger et al., 2002). Protein concentrations were determined by the Bradford method. Proteins (40 µg) were resolved on 15% gels, transferred to a 0.2-µm nitrocellulose membrane, and incubated with cleaved caspase-3 (Asp175)–specific rabbit polyclonal antibody (9661). Blots were developed and analyzed as described above. Blots were reprobed with a ß-actin–specific mouse monoclonal antibody to determine equal protein loading.
Statistics.
Statistical analyses were performed with Statview (SAS Institute, Cary, NC). Data are presented as means ± SEs. At least three experiments were performed in each pro/pre–B cell protocol. Experiments with primary pro–B cells were performed with a minimum of three pools of bone marrow cells, and each pool of bone marrow cells was prepared and maintained separately. One-factor ANOVAs were used in combination with the Dunnett or Tukey-Kramer multiple comparisons tests to analyze the data and determine significant differences.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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MAPK Activation by GW7845, a PPAR
AgonistGW7845 (40µM) induced a time-dependent increase in the percentage of sub-G0/G1 cells indicative of apoptosis in cultured pro/pre–B cells (Fig. 1). Significant death occurred within 60 min of treatment, indicating rapid activation and propagation of a cell death–signaling pathway. Similarly, GW7845 has been shown to activate rapid apoptosis in primary pro–B cells (Schlezinger et al., 2002
). MAPK cascades can be activated within minutes of exposure to the appropriate stimuli and are associated with apoptosis (Wada and Penninger, 2004). Therefore, we investigated the activation of MAPKs in primary pro– and cultured pro/pre–B cells following treatment with GW7845.
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BU-11, a nontransformed murine pro/pre–B-cell line, was treated with Vh (DMSO:ethanol, 50:50, 0.1% final concentration) or GW7845 (40µM) for 15–90 min and analyzed for MAPK phosphorylation and activation by Western blotting and in vitro kinase assay. A low basal level of ERK1/2 phosphorylation was evident in untreated and Vh-treated cells (Fig. 2A). GW7845 induced ERK1/2 phosphorylation within 15 min of treatment, and this was reflected in a small increase in activity measured by an in vitro kinase assay (Fig. 2B). However, the ERK1/2 phosphorylation level returned to the basal level by 30 min and decreased to below the basal level at subsequent time points. Primary pro–B cells did not exhibit a detectable basal level of ERK1/2 phosphorylation, and ERK1/2 phosphorylation was not induced by treatment with GW7845 (Fig. 2C).
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In order to determine if the activation of ERK1/2 has a survival effect, cultured pro/pre–B cells were pretreated with the MEK1/2 inhibitor U0126 (20µM). This concentration of U0126 was sufficient to completely suppress ERK1/2 phosphorylation, as well as the phosphorylation of its substrate p90RSK (data not shown). Death was reduced slightly (26 ± 6% [n = 3, p < 0.05, ANOVA, Tukey-Kramer]), rather than exacerbated, suggesting that the small increase in ERK1/2 activity does not have a survival function. The low-level, transient activation of ERK1/2 suggests that it does not play a major role in supporting or suppressing GW7845-induced pro/pre–B cell apoptosis.
Naive and Vh-treated cultured pro/pre–B cells had a low basal level of p38 MAPK phosphorylation (Fig. 3A). A significant increase in the phosphorylation of p38 MAPK was evident 15 min following treatment with GW7845 and was sustained over time. To determine if the p38 MAPK indeed was active, an in vitro kinase assay was performed using whole-cell extracts from cells treated with either Vh or GW7845 for 60 min. There was a significant increase in the phosphorylation of ATF-2 by p38 MAPK immunoprecipitated from GW7845-treated cells (Fig. 3B). Like the cultured pro/pre–B cells, primary pro–B cells responded to treatment with GW7845 with a sustained increase in p38 MAPK phosphorylation (Fig. 3C).
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Very low levels of phosphorylated JNK were detected in untreated cultured pro/pre–B cells (Fig. 4A). However, like p38 MAPK, JNK was phosphorylated significantly within 15 min of treatment with GW7845, and the phosphorylation was sustained (Fig. 4A). An in vitro kinase assay showed that there was a significant increase in the phosphorylation of c-jun by JNK pulled down from GW7845-treated cells (Fig. 4B). Similarly, primary pro–B cells treated with GW7845 showed a substantial increase in JNK phosphorylation that was sustained through 3 h (Fig. 4C).
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In order to determine if phosphorylation of MAPKs led to activation of these enzymes in vivo, phosphorylation of an endogenous substrate of all three MAPKs, ATF-2, was examined. In cultured pro/pre–B cells and primary pro–B cells, very little ATF-2 is constitutively phosphorylated (Fig. 5). In cultured pro/pre–B cells, ATF-2 phosphorylation was increased significantly following 15 min of treatment with GW7845 and remained high through 30 min (Fig. 5A). After 30 min, the phosphorylation of ATF-2 decreased. In primary pro–B cells, phosphorylation of ATF-2 was sustained through 3 h (Fig. 5B). Results presented below (see Fig. 9 and the "Discussion" section) suggest that, in fact, JNK was responsible for the majority of ATF-2 phosphorylation in these cells. We also investigated the phosphorylation of more restricted substrates, MAPKAP-K2 for p38 MAPK and c-jun for JNK. We present evidence below (see Fig. 9) that suggests that MAPKAP-K2 was phosphorylated following treatment with GW7845. Surprisingly, while we saw a small, but consistent, increase in c-jun phosphorylation, it did not reach statistical significance (data not shown). These results not only show that GW7845 induces the phosphorylation of MAPKs but also show that p38 MAPK and JNK are catalytically active in cultured pro/pre– and primary pro–B cells.
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A p38 MAPK/JNK Dual Inhibitor Blocks Multiple Facets of GW7845-Induced Pro/Pre–B-Cell Apoptosis Signaling
The rapid, strong, and persistent activation of p38 MAPK and JNK prior to overt apoptosis suggests that these kinases are participating in GW7845-induced pro/pre–B cell apoptosis. In addition, inhibition and reduced expression of JNK suppress PPAR
agonist–induced apoptosis in malignant cells of various types (Bae and Song, 2003; Shan et al., 2004). Therefore, we examined the effect of p38 MAPK and JNK inhibitors on GW7845-induced apoptosis in cultured pro/pre–B cells.
Cultured pro/pre–B cells were pretreated for 30 min with Vh (DMSO, 0.1% final concentration) or PD169316 (1–5µM), which acts as a p38 MAPK inhibitor at less than 3µM and as a dual p38 MAPK/JNK inhibitor at more than 5µM (Ortiz et al., 2001). Cells then were treated with Vh (DMSO:ethanol, 50:50, 0.1% final concentration) or GW7845 (40µM) for 2 h and analyzed for apoptosis by PI staining and flow cytometry. GW7845 induced significant apoptosis (
50%) in the cultured pro/pre–B cells after 2 h of treatment as indicated by a shift in the forward- and side-scatter profiles and by the appearance of a sub-G0/G1 peak in the PI-stained cells (Fig. 6). GW7845-induced apoptosis was reduced significantly by 2.5µM PD169316 and completely suppressed by 5µM PD169316 (Fig. 6), suggesting that p38 MAPK and/or JNK are involved in GW7845 apoptosis signaling.
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Apoptosis can be induced by a variety of stresses including cytotoxic agents and irradiation. Stress-induced apoptosis typically involves caspase-8–independent mitochondrial membrane potential depolarization (
m) and/or permeabilization, the formation of an "apoptosome," a death complex composed of cytochrome c, Apaf-1, and caspase-9, and the activation of executioner caspases (reviewed in Jin and El-Deiry, 2005). Therefore, we examined alterations in mitochondrial membrane integrity following treatment with GW7845 and downstream evidence of caspase activation.
Treatment with glitazone drugs can impair the metabolic function of mitochondria (Scatena et al., 2004) and induces the loss of mitochondrial membrane potential in other cell types (Perez-Ortiz et al., 2004; Ray et al., 2004), suggesting that mitochondria are a likely target for GW7845. Cultured pro/pre–B cells were treated with Vh or GW7845 (40µM) for 30–45 min and analyzed either for mitochondrial membrane potential loss by JC-1 staining or for cytochrome c release by Western blotting. Within 30 min of treatment, GW7845 induced a nearly complete loss of mitochondrial membrane potential (Fig. 7A) and the release of cytochrome c (Fig. 7B). Next, we examined the effect of PD169316 on GW7845-induced changes in mitochondrial membrane integrity. Cultured pro/pre–B cells were pretreated with Vh or PD169316 (5µM) for 30 min, treated with Vh or GW7845 (40µM) for 30–45 min, and analyzed as above. PD169316 had no effect on the baseline level or GW7845-induced loss of mitochondrial membrane potential (Fig. 7A). In contrast, PD169316 completely blocked GW7845-induced cytochrome c release (Fig. 7B). Since 5µM PD169316 completely inhibited GW7845-induced apoptosis (Fig. 6), these data suggest that the loss of mitochondrial membrane potential is not sufficient for GW7845-induced apoptosis, but that cytochrome c release is required, and that activation of MAPKs contributes to cytochrome c release but not to changes in mitochondrial membrane potential.
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Caspase-3 is considered to be the primary apoptosis executioner, and its activation results in cleavage of proteins responsible for the classic nuclear features associated with apoptosis, including nuclear condensation, chromatin margination, and DNA fragmentation. As predicted from studies with PI staining, GW7845 (40µM) induced the formation of the 17-kDa active caspase-3 fragment (Fig. 7C) and DNA fragmentation (Fig. 7D). Both these markers of end-stage apoptosis signaling were inhibited with PD169316. The data support the hypothesis that activation of MAPKs is required for GW7845-induced apoptosis signaling and that MAPKs are activated upstream of the mitochondria.
Contribution of p38 MAPK and JNK to GW7845-Induced Pro/Pre–B-Cell Apoptosis
Because both p38 MAPK and JNK are activated in our B cell models following GW7845 treatment and PD169316 is a dual p38 MAPK/JNK inhibitor, we sought to determine if one or both of these MAPKs is/are necessary for apoptosis signaling.
To this end, we examined the ability of several inhibitors of varying specificities to block GW7845-induced apoptosis. Cultured pro/pre–B cells were pretreated with Vh (DMSO, 0.1%), SB202190 (20–40µM), a dual p38 MAPK/JNK inhibitor, SB203580 (20–40µM), a preferential p38 MAPK inhibitor, and/or SP600125 (5–10µM), a preferential JNK inhibitor, for 30 min. Cells then were treated with Vh (DMSO:ethanol, 0.1%) or GW7845 (40µM) for 2 h and analyzed for apoptosis by PI staining and flow cytometry. GW7845-induced apoptosis was suppressed significantly by 20µM SB202190, and the suppression increased at 40µM (Fig. 8A). SB203580 significantly, but incompletely, inhibited GW7845-induced apoptosis (Fig. 8B). Similarly, SP600125 significantly, but incompletely, inhibited GW7845-induced apoptosis (Fig. 8C). The expectation would be that addition of both SB203580 and SP600125 would at least additively inhibit GW7845-induced apoptosis. Indeed, coadministration of SB203580 and SP600125 inhibited GW7845-induced apoptosis in an additive fashion (Fig. 8D).
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To examine the effect of these inhibitors more specifically on p38 MAPK and JNK activation in cultured pro/pre–B cells, cells were pretreated with Vh, PD169316 (5µM), SB202190 (40µM), SB203580 (40µM), or SP600125 (10µM) for 30 min, treated with Vh or GW7845 (40µM) for 30 min, and analyzed for activation of p38 MAPK and JNK. PD169316 has been shown to block the activation and function of p38 MAPK and JNK (Ortiz et al., 2001
). Therefore, we began by examining the phosphorylation of p38 MAPK. As seen in previous experiments, GW7845 induced significant phosphorylation of p38 MAPK after 30 min of treatment (Fig. 9A). PD169316 reduced the basal level of p38 MAPK phosphorylation (lane 2) and completely suppressed the GW7845-induced p38 MAPK phosphorylation (lane 4). GW7845 induced a shift in the apparent molecular weight of MAPKAP-K2, which we interpreted as an indication of phosphorylation because MAPKAP-K2 is a known substrate of p38 MAPK, the shift was induced by H2O2, a positive control (data not shown), and the shift could be completely inhibited by SB203680 (see below), a well-established property of this compound (Eyers et al., 1999). Like p38 MAPK, GW7845 induced JNK phosphorylation, but PD169316 only slightly reduced this phosphorylation (Fig. 9A). However, PD169316 significantly reduced GW7845-induced ATF-2 phosphorylation, either when ATF-2 phosphorylation levels were normalized to total ATF-2 (Fig. 9A) or to ß-actin levels (data not shown). The data suggest that PD169316 inhibits the activation of p38 MAPK and the activity of JNK in cultured pro/pre–B cells. Despite their similarities in structure to PD169316, SB202190 and SB203580 are only known to inhibit MAPK activity; therefore, we examined the effects of these inhibitors on the GW7845-induced changes in MAPKAP-K2 molecular weight and ATF-2 phosphorylation (Fig. 9B). Both SB202190 and SB203580 reversed the change in the apparent molecular weight of MAPKAP-K2 induced by GW7845. SB203580 did not inhibit GW7845-induced ATF-2 phosphorylation and potentially enhanced it. However, SB202190 also reduced ATF-2 phosphorylation that was induced by GW7845 when the phosphorylated ATF-2 levels were normalized to total ATF-2 (Fig. 9B) or to ß-actin levels (data not shown). Interestingly, we observed that GW7845 appeared not only to increase the level of phosphorylated ATF-2 in cytoplasmic extracts but also the expression of total ATF-2 (fold change from naive: Vh; 1.0 ± 0.1, GW; 1.8 ± 0.2, p < 0.01, Student t test), and that SB202190, along with PD169316 (Fig. 9A) and SP600125 (Fig. 9C), reduced both the level of GW7845-induced phosphorylated ATF-2 and total cytoplasmic ATF-2. Thus, the data show that while SB203580 appeared to specifically inhibit p38 MAPK activity and only partially inhibit GW7845-induced death, SB202190 appeared to have a broader effect on p38 MAPK and JNK that correlated with its greater ability to suppress GW7845-induced death.
Finally, we examined the effect of SP600125 on GW7845-induced MAPK activity (Fig. 9C). As would be predicted, SP600125 did not affect the GW7845-induced increase in the apparent molecular weight of MAPKAP-K2, but did significantly suppress the phosphorylation of ATF-2, whether phosphorylation levels were normalized to total ATF-2 (Fig. 9C) or to ß-actin levels (data not shown). The data support the hypothesis that activation of both p38 MAPK and JNK contribute to initiating GW7845-induced death; however, activation of either alone is not sufficient to support maximal GW7845-induced apoptosis.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Apoptosis is a critical event during the early stages of B lymphopoiesis and occurs at the highest rate during the pro/pre–B-cell transition and during clonal deletion (Lu and Osmond, 2000
). Studies performed with transformed and primary immature B cells have begun to map out apoptosis pathways invoked during clonal deletion (as reviewed in Ryu et al., 2005). Our studies are focused on determining the events that can lead to apoptosis at an earlier stage of B-cell development when cultures are exposed to immunosuppressive chemicals and determining whether B cells early in development are sensitized to apoptotic agents due to the importance of this process at this developmental stage.
We have shown that the synthetic PPAR agonists GW7845 and ciglitazone induce apoptosis in cultured pro/pre–B cells (Schlezinger et al., 2002). Results presented here show that GW7845-induced death is associated with activation of MAPKs. GW7845 induced the sustained phosphorylation of the MAPKs, p38 MAPK, and JNK in primary pro– and cultured pro/pre–B cells. ERK1/2, p38 MAPK, and JNK activation have been observed following treatment with 15d-PGJ2 (Kim et al., 2003; Lennon et al., 2002; Shan et al., 2004; Takeda et al., 2001), glitazone drugs (Bae and Song, 2003; Gardner et al., 2005; Lennon et al., 2002; Takeda et al., 2001; Teruel et al., 2003), and mono-2-ethylhexyl phthalate (Rokos and Ledwith, 1997). Cell context and the agonist used are likely to play a role in determining which kinases are activated and the duration of their activation. In the case of GW7845 and our B-cell models, the phosphorylation of p38 MAPK and JNK was very rapid (< 15 min) and sustained, while ERK1/2 was phosphorylated only transiently.
Not only were p38 MAPK and JNK phosphorylated in our B-cell models following treatment with GW7845, in vitro kinase assays confirmed their functional activation. In addition, we were interested in determining which substrates were phosphorylated in vivo. ATF-2 is a substrate for all three MAPKs (Morton et al., 2004) and thus a likely target in our system. ATF-2 was strongly phosphorylated following treatment with GW7845 in both the B-cell models. The facts that the p38 MAPK inhibitor SB203580 did not reduce ATF-2 phosphorylation and that the JNK inhibitor SP600125 significantly reduced ATF-2 phosphorylation support the idea that JNK is responsible for the majority of ATF-2 phosphorylation (see Fig. 9). While JNK was activated strongly in these B cells, we observed only a small change in c-jun phosphorylation (data not shown). This may result from the fact that there is a relatively low expression level of c-jun in these cells or that the ratio of ATF-2/c-jun phosphorylation is mediated by the type of activating agent, i.e., ATF-2 is activated to a greater extent than c-jun by genotoxic agents and cellular stresses such as UV irradiation (van Dam et al., 1995). Interestingly, we observed that GW7845 treatment appeared to increase the expression of ATF-2 in the cytoplasm. This is unlikely to be a result of a change in gene expression, as this effect was not observed when total ATF-2 was quantified in whole-cell extracts (see Fig. 5). However, this effect may result from a GW7845-induced change in ATF-2 nuclear/cytoplasmic shuttling, a phenomenon important in regulating ATF-2 activity (Liu et al., 2006). Finally, we show evidence that treatment with GW7845 results in changes in the apparent molecular weight of MAPKAP-K2 that is consistent with its phosphorylation.
The dual p38 MAPK/JNK inhibitor PD169316 potently inhibited GW7845-induced cultured pro/pre–B-cell death. As expected, the results showed that PD169316 blocked not only p38 MAPK activation but also JNK activity. Cytochrome c release, caspase-3 activation, and DNA fragmentation were induced following GW7845 treatment, and PD169316 inhibited all three GW7845-induced effects. These results are not unlike those found with PD169316 and retinoid-related, molecule-induced death in Jurkat T cells (Ortiz et al., 2001). Interestingly, PD169316 did not block the GW7845-induced loss of mitochondrial membrane potential, indicating that loss of mitochondrial membrane potential is not sufficient to induce death in this system and may indeed be irrelevant to apoptosis signaling. Importantly, the results suggest that MAPKs participate in initiating the GW7845-induced death pathway upstream of the mitochondria.
The fact that PD169316 inhibits both p38 MAPK and JNK did not allow us to determine if one or both of these MAPKs contribute to GW7845-induced death. Previous studies have stressed the role of JNK in particular in PPAR agonist–induced apoptosis. The JNK inhibitor SP600125 and dominant-negative JNK constructs decreased troglitazone- and 15d-PGJ2–induced death in hepatoma cells (Bae and Song, 2003). In addition, SB202190, a more general MAPK/JNK inhibitor, blocked 15d-PGJ2–induced articular chondrocyte apoptosis (Shan et al., 2004). Here, the data suggest that both p38 MAPK and JNK play a role in GW7845-induced apoptosis in pro/pre–B cells. Compounds that blocked the apparent phosphorylation of only one of the MAPK substrates, SB203580 and SP600125, only partially blocked GW7845-induced apoptosis. However, the compounds that altered the apparent phosphorylation of both MAPKAP-K2 and ATF-2 had the greatest efficacy in blocking GW7845-induced death. Therefore, it appears that at least activation of p38 MAPK and JNK contribute to initiation of GW7845-induced apoptosis.
The results do not rule out the possibility of the contribution of another downstream kinase, as suggested by the failure of a combination of single specificity kinase inhibitors to completely block apoptosis. Although PD169316 was originally shown to inhibit p38 MAPK activity, we and others have shown that it also inhibits phosphorylation of p38 MAPK (see Ortiz et al., 2001). This may reflect PD169316-mediated blocking of an essential phosphorylation site for the upstream MAPKK. Alternatively, PD169316 may inhibit the activity of an upstream kinase. In addition, PD169316 inhibited GW7845-induced death with a greater potency than SB202190, the other compound that had a similar efficacy in suppressing death.
The question remains as to how p38 MAPK and JNK initiate the apoptotic pathway following treatment with GW7845. One pathway by which JNK acts is through the phosphorylation of activator protein-1 (AP-1) family members and the induction of AP-1–DNA binding. Despite the phosphorylation of JNK and its substrate ATF-2, we saw no change in AP-1–DNA binding following treatment with GW7845 (data not shown). This is reminiscent of results seen following cross-linking of Fas in Jurkat T cells and might result from a lack of required heterodimerization partners (Lenczowski et al., 1997). Considering that we have shown that GW7845-induced alteration of mitochondrial membrane integrity consistent with a mitochondrially mediated apoptotic pathway, it is more likely that the MAPKs are exerting their effect on Bcl2 family members. Both p38 MAPK and JNK phosphorylate Bcl2 family members such as Bid and Bax, and MAPK-dependent Bcl2 phosphorylation has been shown to result in apoptosis (Lei et al., 2002; Tourian et al., 2004).
Data presented here have established the role of MAPKs in GW7845-induced pro/pre–B cell apoptosis; however, it remains to be determined how PPAR interfaces with the MAPK cascade. Recent studies have questioned the role that PPAR plays in inhibition of proliferation caused by its agonists. Troglitazone was found to be equally effective at inhibiting proliferation in both PPAR–/– and PPAR+/+ mouse embryonic stem cells (Palakurthi et al., 2001), and there is growing evidence for PPAR-independent effects of PPAR agonists, particularly at higher concentrations (Berry et al., 2005; Chintharlapalli et al., 2005). Indeed, the concentration of GW7845 required to induce MAPK activation and apoptosis (20µM) is substantially higher than that required to activate PPAR (10–1000nM).
Nevertheless, a contribution of PPAR to apoptosis in general is supported by the facts that PPAR antagonists reduce the magnitude of apoptosis induced by several diverse PPAR agonists, including ring-substituted diindolylmethanes, CDDO, 15d-PGJ2, glitazone drugs, and mono-2-ethylhexyl phthalate (Chen et al., 2005; Contractor et al., 2005; Kim et al., 2003; Shan et al., 2004; Yokoyama et al., 2003). PPAR is expressed in these cultured pro/pre–B cells, and GW7845 induces PPAR-DNA binding (Schlezinger et al., 2002, 2004). In addition, cotreatment of PPAR agonists with 9-cis-retinoic acid synergistically increases the incidence of apoptosis in cultured pro/pre–B cells, an effect also seen in other cell types (Contractor et al., 2005; Konopleva et al., 2004; Ray et al., 2004; Schlezinger et al., 2002, 2004). An important role for PPAR and PPAR agonists in B-cell growth and death is supported by the high level of expression of PPAR in the immune system (Braissant et al., 1996) and the alteration of B-cell proliferation and function in the haploinsufficient mouse (Setoguchi et al., 2001). Studies are underway to determine the interface of PPAR with MAPK activation induced by PPAR agonists, to identify the upstream members of the kinase cascade, and to determine the role of MAPK and PPAR in pro/pre–B-cell apoptosis induced by structurally distinct PPAR agonists. By understanding these mechanisms, strategies may be developed to avoid the potentially immunosuppressive effects of PPAR agonists used as chemotherapeutics and present in the human environment as pollutants.
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ACKNOWLEDGMENTS |
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This work was supported by NIH RO1-ES06086, NIH PO1-HL68705, and Superfund Basic Research Grant 2P42ES007381-12. We thank Ms Stephanie Bissonnette for her technical assistance.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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