ology 215 (2006) 198–207www.elsevier.com/locate/ytaap

Toxicology and Applied Pharmac

Activation of peroxisome proliferator-activated receptor-γ (PPARγ) inducescell death through MAPK-dependent mechanism in osteoblastic cells

Sung Hun Kim a, Chong Il Yoo a,c, Hui Taek Kim a,c, Ji Yeon Park b,Chae Hwa Kwon b, Yong Keun Kim b,c,d,⁎

a Department of Orthopedic Surgery, College of Medicine, Pusan National University, Pusan, 602-739, Koreab Department of Physiology, College of Medicine, Pusan National University, Pusan, 602-739, Koreac Medical Research Institute, College of Medicine, Pusan National University, Pusan, 602-739, Korea

d MRC for Ischemic Tissue Regeneration, College of Medicine, Pusan National University, Pusan, 602-739, Korea

Received 12 December 2005; revised 21 February 2006; accepted 2 March 2006Available online 17 April 2006

Abstract

The present study was undertaken to determine the role of the mitogen-activated protein kinase (MAPK) subfamilies in cell death induced byPPARγ agonists in osteoblastic cells. Ciglitazone and troglitazone, PPARγ agonists, resulted in a concentration- and time-dependent cell death,which was largely attributed to apoptosis. But a PPARα agonist ciprofibrate did not affect the cell death. Ciglitazone caused reactive oxygenspecies (ROS) generation and ciglitazone-induced cell death was prevented by antioxidants, suggesting an important role of ROS generation in theciglitazone-induced cell death. ROS generation and cell death induced by ciglitazone were inhibited by the PPARγ antagonist GW9662.Ciglitazone treatment caused activation of extracellular signal-regulated kinase (ERK) and p38. Activation of ERK was dependent on epidermalgrowth factor receptor (EGFR) and that of p38 was independent. Ciglitazone-induced cell death was significantly prevented by PD98059, aninhibitor of ERK upstream kinase MEK1/2, and SB203580, a p38 inhibitor. Ciglitazone treatment increased Bax expression and caused a loss ofmitochondrial membrane potential, and its effect was prevented by N-acetylcysteine, PD98059, and SB203580. Ciglitazone induced caspaseactivation, which was prevented by PD98059 and SB203580. The general caspase inhibitor z-DEVD-FMK and the specific inhibitor of caspases-3DEVD-CHO exerted the protective effect against the ciglitazone-induced cell death. The EGFR inhibitors AG1478 and suramin protected againstthe ciglitazone-induced cell death. Taken together, these findings suggest that the MAPK signaling pathways play an active role in mediating theciglitazone-induced cell death of osteoblasts and function upstream of a mitochondria-dependent mechanism. These data may provide a novelinsight into potential therapeutic strategies for treatment of osteoporosis.© 2006 Elsevier Inc. All rights reserved.

Keywords: PPARγ; Ciglitazone; Cell death; MAPK; Mitochondrial dysfunction; Osteoblastic cells

Introduction

Osteoporosis is characterized by low bone mineral densityand by microarchitectural deterioration of bone tissue with aconsequent increase in bone fragility and susceptibility tofracture (Kanis et al., 1994). Maintenance of bone homeostasisthroughout life relies on the bone remodeling process, whichcontinually replaces old and damaged bone with new bone(Riggs and Parfitt, 2005). Two types of cells are involved in

⁎ Corresponding author. Department of Physiology, College of Medicine,Pusan National University, Pusan, 602-739, Korea. Fax: +51 246 6001.

E-mail address: kim430@pusan.ac.kr (Y.K. Kim).

0041-008X/$ - see front matter © 2006 Elsevier Inc. All rights reserved.doi:10.1016/j.taap.2006.03.001

bone remodeling: osteoclasts originating from hematopoieticcells are responsible for bone resorption, and osteoblastsoriginating from mesenchymal cells are responsible forformation of new bone. Thus, an overproduction of osteoclastsrelative to the need for remodeling and an undersupply ofosteoblasts relative to the need for cavity repair represent thefundamental pathophysiologic changes in type I and type IIosteoporosis (Manolagas, 1998).

The peroxisome proliferator-activated receptors (PPARs) area family of ligand activated transcription factors belonging to thenuclear receptor superfamily (Willson et al., 2001). PPARsmodulate target gene expression in response to ligand activationafter heterodimerization with retinoid X receptor α and binding

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to peroxisome proliferator-responsive elements of target genes.Three different isoforms have been identified, PPARα, PPARβ,and PPARγ, each with distinct physiological functions (Dreyeret al., 1992; Issemann and Green, 1990). PPARs are the primarytargets of numerous natural and synthetic compounds includingphthalate plasticizers and long-chain fatty acids. Among them,PPARγ is of particular interest because it has been implicated inmany human diseases including type II diabetes, atherosclerosis,hypertension, inflammation, and cancer. PPARs also play acritical physiological role as regulators of lipid metabolism,glucose homeostasis, and cell proliferation (Guan and Breyer,2001).

Although several studies have reported that activation ofPPARγ promote cell survival (Lim et al., 2004; Wang et al.,2002), activation of PPARγ is almost always associated with celldeath (Chattopadhyay et al., 2000; Chen et al., 2003; Chinetti etal., 1998; Kim et al., 2003; Yamakawa-Karakida et al., 2002).

PPARγ has been shown to be expressed in osteoblast cells(Jackson and Demer, 2000; Ogawa et al., 1999). Several studieshave demonstrated that PPARγ plays an essential role inmaintaining bone homeostasis. In vivo treatment of rosiglita-zone, a high affinity ligand of PPARγ, in mice causes reductionin bone formation rate (Ali et al., 2005; Werner and Travaglini,2001). Moreover, treatment of rosiglitazone decreased thenumber of osteoblasts while simultaneously increasing thenumber of marrow adipocytes (Rzonca et al., 2004). PPARγinsufficiency increased bone mass by stimulating osteoblasto-genesis from bone marrow progenitors (Akune et al., 2004). Invitro studies have shown similar results (Jackson and Demer,2000; Rzonca et al., 2004). These findings suggest thatactivation of PPARγ is a negative regulator of bone mass, andthat the increased production of oxidized fatty acids with agemay indeed be an important mechanism for age-relatedosteoporosis in humans. Such effects of PPARγ agonists maybe attributed to their abilities to induce apoptosis of osteoblasts.However, the molecular mechanism by which PPARγ inducesapoptosis in osteoblastic cells has not been established.

PPARγ has been shown to activate members of the mitogen-activated protein kinase (MAPK) family (Gardner et al., 2003;Lennon et al., 2002; Shan et al., 2004). MAPK signalingpathways play a key role in a variety of cellular responses, suchas cell proliferation, differentiation, and cell death (Cobb, 1999;Davis, 2000; Xia et al., 1995). This study was undertaken todetermine whether activation of PPARγ induces cell death and,if so, to examine the role of MAPK pathways in cell death ofMC3T3-E1 cells, an established osteoblast cell line.

Materials and methods

Reagents. Ciglitazone, troglitazone, ciprofibrate, superoxide dismutase(SOD), catalase, N-acetylcysteine (NAC), 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT), carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP), propidium iodide, and suramin were purchased fromSigma-Aldrich Chemical (St. Louis, MO, USA). Tween 20, PD98059,SB203580, GW9662, z-DEVD-FMK, DEVD-CHO, and AG1478 werepurchased from Calbiochem (California, USA). 3,3′-Dihexyloxacarbocyamide[DiOC6(3)] and 2′,7′-dichlorofluorescein diacetate (DCFH-DA) were obtainedfrom Molecular Probes (Eugene, OR, USA). Antibodies of MAPK subfamilies,

Bax, and α-actin were obtained from Cell Signaling Technology Inc. (Beverly,MA, USA). All other chemicals were of the highest commercial grade available.

Cell culture. MC3T3-E1 mouse osteoblastic cells were obtained from theAmerican Type Culture Collection (Rockville, MD, USA) and maintained byserial passages in 75-cm2 culture flasks (Costar, Cambridge, MA,USA). Thecells were grown in Minimum Essential Medium Alpha Medium (αMEM,Gibco, Invitrogen co., California, USA) containing 10% heat inactivated fetalbovine serum (Hyclone, UT, USA) at 37 °C in humidified 95% air/5% CO2

incubator. When the cells reached confluence, subcultures were prepared using0.02% EDTA–0.05% trypsin solution. The cells were cultured on 24- or 6-welltissue culture plates in αMEM medium containing 10% fetal bovine serum.Serum was removed from culture media at the time of agent addition.

Measurement of cell viability. Cell viability was evaluated using a MTTassay (Denizot and Lang, 1986). Tetrazolium salts such as MTTare metabolizedby mitochondrial dehydrogenases to form a blue formazan dye and are thereforeuseful for the measurement of cell viability. After washing the cells, culturemedium containing 0.5 mg/ml of MTT was added to each well. The cells wereincubated for 2 h at 37 °C, the supernatant was removed, and the formedformazan crystals in viable cells were solubilized with 0.11 ml of dimethylsulf-oxide. A 0.1 ml aliquot of each sample was then translated to 96-well plates, andthe absorbance of each well was measured at 550 nm with ELISA Reader (Bio-Tek instrument, EL,311). Data were expressed as a percentage of controlmeasured in the absence of PPARγ agonists. Unless otherwise stated, the cellswere treated with 10 μM ciglitazone for 24 h. Test reagents were added to themedium 30 min before ciglitazone exposure.

Measurement of apoptosis

1) Annexin V staining: phosphatidylserine exposure on the outer layer of thecell membrane was measured using the binding of annexin V–fluoresceinisothiocyanate (FITC). Cells were harvested and washed with cold PBS.And then the cells were incubated for 15 min with annexin V–FITC andpropidium iodide and were analyzed by flow cytometry (Becton Dickinson,Franklin Lakes, NJ).

2) Flow cytometric analysis: cells were grown in 6-well plates and were treatedas indicated. Then attached and floating cells were pooled, pelleted bycentrifugation, washed in PBS, and fixed with cold 70% ethanol containing0.5% Tween 20 at 4 °C overnight. Cells were washed and resuspended in1.0 ml of propidium iodide solution containing 100 μg of RNase A/ml and50 μg propidium iodide/ml and incubated for 30 min at 37 °C. Apoptoticcells were assayed using FACSort Becton Dickinson Flow Cytometer at488 nm, and data were analyzed with CELLQuest Software. Cells with sub-G1 propidium iodide incorporation were considered as apoptotic. Thepercentage of apoptotic cells was calculated as the ratio of events on sub-G1

to events from the whole population.

Western blot analysis. Cells were harvest at various times after ciglitazonetreatment and disrupted in lysis buffer (1% Triton X-100, 1 mM EGTA, 1 mMEDTA, 10 mM Tris–HCl, pH 7.4). Cell debris was removed by centrifugation at10,000 g for 10 min at 4 °C. The resulting supernatants were resolved on a 10%SDS-PAGE under denatured reducing conditions and transferred to nitrocellu-lose membranes. The membranes were blocked with 5% nonfat dried milk atroom temperature for 30 min and incubated with different primary antibodies.The membranes were washed and incubated with horseradish peroxidase-conjugated secondary antibodies. The signal was visualized using an enhancedchemiluminescence (Amersham, Buckinghamshire, UK).

Measurement of reactive oxygen species (ROS). The intracellular generationof ROS was measured using DCFH-DA. The nonfluorescent ester penetratesinto the cells and is hydrolyzed to DCFH by the cellular esterases. The probe(DCFH) is rapidly oxidized to the highly fluorescent compound 2′,7′-dichlorofluorescein (DCF) in the presence of cellular peroxidase and ROSsuch as hydrogen peroxide or fatty acid peroxides. Cells cultured in 24-wellplate were preincubated in αMEM with 30 μM DCFH-DA for 1 h at 37 °C.After the preincubation, the cells were exposed to 10 μM ciglitazone for variousdurations. Changes in DCF fluorescence was assayed using FACSort Becton

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Dickinson Flow Cytometer (Becton-Dickinson Bioscience, San Jose, CA,USA), and data were analyzed with CELLQuest Software. In other experiments,the fluorescent intensity was monitored using a fluorescence microscope (LeicaDMIRB, Wetzlar, Germany) at an excitation of 475 nm and emission of 525 nm.

Measurement of mitochondrial membrane potential. The mitochondrialtransmembrane potential was measured with DiOC6(3), a fluorochrome that isincorporated into cells depending upon the mitochondrial membrane potential(Pastorino et al., 1998). Loss in DiOC6(3) staining indicates disruption of themitochondrial inner transmembrane potential. Cells were stained with DiOC6(3)at a final concentration of 50 nM for 20 min at 37 °C in the dark. Cells werewashed and resuspended in Hank's balanced salts solution containing Ca2+ andMg2+. The fluorescence intensity was analyzed with a FACScan flow cytometerusing the fluorescence signal 1 channel.

Statistical analysis. The data are expressed as means ± SEM, and the diffe-rence between two groups was evaluated using Student's t test. Multiple groupcomparison was done using one-way analysis of variance followed by the Tur-key post hoc test. A probability level of 0.05 was used to establish significance.

Results

Effect of PPARγ agonists on cell viability

Cell viability was measured in cells exposed to variousconcentrations of PPARγ agonists ciglitazone and troglitazoneand a PPARα agonist ciprofibrate. Ciglitazone and troglita-zone caused loss of cell viability in dose- and time-dependentmanner with similar potency, whereas ciprofibrate did notinduce loss of cell viability (Figs. 1A and B). Ciprofibrate didnot induce a loss of cell viability even when the concentrationwas increased up to 100 μM (data not shown).

To determine whether the cell death was attributed toapoptosis, the cells were subjected to annexin V staining.Cytometric analyses of annexin V-positive cells demonstrated a42.70% increase in cells treated with ciglitazone as shown inright lower and upper quadrants (Fig. 1C). We further confirmedthe onset of apoptotic cell death by flow cytometric analysis. Asshown in Fig. 1D, treatment of ciglitazone induced an increasein the sub-G0/G1 peak from 1.16% to 45.75%, an indicator ofapoptosis cells. These results suggest that ciglitazone-inducedcell death is largely due to apoptosis.

Role of ROS in ciglitazone cytotoxicity

To determine whether ciglitazone induces ROS generation inosteoblasts, cells were exposed to ciglitazone and changes inDCF fluorescence were measured using flow cytometry. After3 h of treatment, ciglitazone caused an increase in theproportion of cells with higher fluorescence intensity (Fig.2A), indicating that ciglitazone induces ROS generation.Further evidence of ROS generation by ciglitazone was detectedby fluorescence microscope. In control cells, little or nofluorescence was observed. However, there was a significantincrease (green color) in DCF fluorescence in cells exposed tociglitazone (Fig. 2B). Such changes were inhibited by thePPARγ antagonist GW9662 (Fig. 2C), suggesting thatciglitazone induces ROS generation through PPARγ receptor.

To examine whether ROS generation is responsible for theciglitazone-induced cell death, effects of various ROS scaven-

gers on the cell death were evaluated. Cell death induced byciglitazone was prevented by the superoxide scavenger, SOD,the hydrogen peroxide scavenger, catalase, and the antioxidant,NAC (Fig. 3). These results suggest that ROS generation isinvolved in the ciglitazone-induced cell death in osteoblasticcells.

Role of PPARγ receptor activation in ciglitazone cytotoxicity

The role of PPARγ receptor in ciglitazone cytotoxicitywas evaluated using receptor antagonist. As shown in Fig.3, the PPARγ antagonist GW9662 significantly preventedcell death induced by ciglitazone, indicating that ciglitazoneinduces cell death through PPARγ receptor.

Roles of MAPK signaling in ciglitazone-induced cell death

To investigate possible pathways of cell death induced byciglitazone, MAPK activities were evaluated by detectingphosphorylation of MAPK subfamilies. Cells were exposed to10 μM ciglitazone for various times and activation of ERK, p38,and JNK was determined by Western blot analysis usingantibodies specific to the respective phosphorylated form.Activation of ERK and p38 was observed after addition ofciglitazone to osteoblasts, and the activation was detectable after30 min and was sustained for 12 h (Figs. 4A and B). Activationof ERK and p38 was inhibited by PD98059, a specific inhibitorof ERK upstream kinase MEK1/2, and SB203580, a p38inhibitor, respectively (Fig. 4C).

To evaluate if these kinases are involved in ciglitazone-induced cell death, the effects of MAPK inhibitors on the celldeath were examined. Cells were preincubated for 30 min withPD98059 and. Thereafter, the cells were exposed to 10 μMciglitazone for 24 h. These inhibitors were effective inpreventing the cell death (Fig. 4D), suggesting that activationof the ERK and p38 plays an important role in the ciglitazone-induced osteoblastic cell death.

Role of Bax expression and mitochondria inciglitazone-induced cell death

Since numerous death signals converge on mitochondriathrough the activation of pro-apoptotic members of the Bcl-2family such as Bax (Birbes et al., 2002), ciglitazone may inducecell death through a Bax-dependent pathway. To test thispossibility, we examined the effect of ciglitazone on Baxexpression. Ciglitazone treatment caused an increase in Baxexpression after 30 min of ciglitazone addition (Fig. 5A).

The increase in Bax expression may cause mitochondrialpermeability transition (MPT) to induce cell death. Cells wereexposed to 10 μM ciglitazone and mitochondrial membranepotential was measured using the fluorescence dye. After 6 hof ciglitazone treatment, disruption of mitochondrial mem-brane potential was observed as evidenced by an increase inthe proportion of cells with lower fluorescence intensity,which was prevented by the antioxidant NAC (Fig. 5B).Ciglitazone-induced changes in mitochondrial membrane

Fig. 1. Dose- and time-dependent effect of PPARα and PPARγ agonists on cell viability. Cells were exposed to various concentrations of ciprofibrate, ciglitazone, andtroglitazone for 24 h (A) or to each 10 μM of ciglitazone and troglitazone for various times (B). Cell viability was estimated by MTT assay. Data are mean ± SEM offour independent experiments performed in duplicate. *P < 0.05 compared with control without reagents. (C) Effect of ciglitazone on apoptotic cell death. Cellsexposed to 10 μM of ciglitazone for 24 h were stained with FITC-conjugated annexin and quantified by flow cytometric analysis. Numbers indicate the percentage ofcells in each quadrant. Early apoptotic and late apoptotic cells were shown in right lower and right upper quadrants, respectively. (D) Cells were stained with propidiumiodide and analyzed by flow cytometry. Numbers indicate the percentage of cells with sub-G1 peak.

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potential were compared with those of FCCP which serves asa positive control. Exposure of cells to 10 μM ciglitazone for6 h showed changes comparable to cells exposed to 20 μMFCCP for 30 min. To ascertain whether activation of MAPKsubfamilies mediates MPT induced by ciglitazone, cells wereexposed to 10 μM ciglitazone for 6 h in the presence ofMAPK inhibitors. Changes in mitochondrial membranepotential was estimated by the extent of M2 region thatrepresents a population of cells with lower fluorescenceintensity. As shown in Fig. 5C, inhibitors of ERK and p38prevented the ciglitazone-induced MPT as evidenced by adecrease in the extent of M2 region.

Role of caspase in ciglitazone-induced cell death

Caspases play a key role during the execution phase invarious forms of apoptosis (Cohen, 1997). Therefore, in

order to investigate whether ciglitazone induces apoptosisthrough a caspase-dependent mechanism, the effect ofcaspase inhibitors on ciglitazone-induced cell death wasexamined. As shown in Fig. 6A, ciglitazone increasedcaspase activity, and its effect was prevented by inhibitors ofERK and p38. The general caspase inhibitor z-DEVD-FMKand the caspase-3 inhibitor DEVD-CHO also prevented theciglitazone-induced cell death (Fig. 6B), indicating that theciglitazone-induced cell death is associated with caspaseactivation.

Role of epidermal growth factor receptor (EGFR) in ERKactivation and cell death induced by ciglitazone

To determine if ciglitazone induced phosphorylation of ERKand p38 through activation of EGFR, the effect of AG1478, aselective inhibitor of EGFR, on the phosphorylation of ERK

Fig. 3. Effects of antioxidants and PPARγ antagonist on ciglitazone-induced celldeath. Cells were exposed to 10 μM ciglitazone for 24 h in the presence orabsence of 500 units/ml superoxide dismutase (SOD), 500 units/ml catalase(Cat), 2 mM N-acetylcysteine (NAC), and 10 μM GW9662. Data aremean ± SEM of three independent experiments performed in duplicate.*P < 0.05 compared with ciglitazone alone (None).

Fig. 2. Effect of ciglitazone on reactive oxygen species generation. (A) Cells were exposed to 10 μM ciglitazone for 3 and 12 h and the DCF fluorescence intensity wasmeasured by a flow cytometer. (B) Cells were exposed to 10 μMciglitazone for 12 h, and theDCF fluorescence intensity wasmonitored by fluorescencemicroscope. (C)Cells were exposed to 10 μMciglitazone for 12 h in the presence or absence of 10 μMGW9662, and the DCF fluorescence intensity wasmonitored by a flow cytometer.

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and p38 was examined. As shown in Fig. 4C, the phosphor-ylation of ERK, but not p38, was inhibited by AG1478.However, phosphorylation of both kinases was inhibited byNAC, suggesting involvement of ROS in activation of ERK andp38 induced by ciglitazone.

To determine whether ciglitazone causes cell death throughEGFR activation, the effect of EGFR inhibitors on the cell deathwas examined. The ciglitazone-induced cell death waseffectively prevented by AG1478 and the general inhibitor ofgrowth factor receptor, suramin (Fig. 7), suggesting involve-ment of EGFR/ERK signaling pathway in the ciglitazone-induced cell death.

Discussion

PPAR ligands can increase cell survival or induce cell deathdepending on the experimental system. The molecular mech-anism underlying these conflicting effects on cell death is

Fig. 4. Effect of activation of MAPK subfamilies on ciglitazone-induced cell death. (A) Representative expression of phospho-ERK1/2 (p-ERK) and phospho-p38 (p-p38). Cells were exposed to 10 μM ciglitazone for various times. α-Actin was employed as loading control. (B) Ciglitazone-induced activation of ERK and p38 wasquantified by densitometry. Data are mean ± SEM of four independent experiments performed in duplicate. (C) Effects of inhibitors on phosphorylation of ERK andp38. Cells were exposed to 10 μM ciglitazone for 12 h in the presence or absence of 10 μM of PD98059 (PD), 10 μM SB203580 (SB), 10 μM AG1478 (AG), and2 mM N-acetylcysteine (NAC). (D) Effects of MAPK inhibitors on ciglitazone-induced cell death. Cells were exposed to 10 μM ciglitazone for 24 h in the presence orabsence of each 10 μM of PD98059 (PD) and SB203580 (SB). Data are mean ± SEM of three independent experiments performed in duplicate. *P < 0.05 comparedwith ciglitazone alone (None).

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poorly understood, but different effects of PPAR ligands appearto depend on the ligand used and/or the cell type studied(Nikitakis et al., 2002; Perez-Ortiz et al., 2004; Rosen andSpiegelman, 2001; Zander et al., 2002). The present studydemonstrates that osteoblastic cells are sensitive to cytotoxicityinduced by two PPARγ agonists ciglitazone and troglitazonebut not by the PPARα agonist ciprofibrate. Ciglitazone has beenreported to have a dual effect on the survival of primaryastrocytes because it increased survival at 10 μM and causes adecrease in viability at 20 μM (Perez-Ortiz et al., 2004). Similaropposing biological results depending on the concentrationwere reported in studies with 15-deoxy-Δ12,14-prostaglandin J2(15d-PGJ2), a natural ligand of PPARγ receptor (Clay et al.,

2002; Rovin et al., 2002). In the present study, however, twoglitazones induced apoptotic cell death in a dose- and time-dependent manner without any evidence for survival over theconcentrations examined (Fig. 1).

Cell deaths by a number of stimuli are frequentlyaccompanied by the generation of ROS, which may act asimportant mediators of cell death (Martindale and Holbrook,2002). Previous studies have shown that various PPARγ ligandsinduce ROS production, and this production plays key role inthe cytotoxicity of ciglitazone and troglitazone (Kondo et al.,2001; Lennon et al., 2002; Li et al., 2001; Narayanan et al.,2003; Perez-Ortiz et al., 2004). However, Atarod and Kehrer(2004) reported that ROS production induced by PPARγ

Fig. 5. (A) Effect of ciglitazone on Bax expression. Cells were exposed to 10 μMciglitazone for various times. (B) Effect of ciglitazone on mitochondrialmembrane potential. Cells were exposed to 10 μM ciglitazone (Cig) for 6 h inthe presence or absence of 2 mM N-acetylcysteine (N). Cells were exposed to20 μM FCCP for 30 min. FCCP serves as a positive control. Mitochondrialmembrane potential was estimated by the uptake of a membrane potential-sensitive fluorescence dye DiCO6(3). The fluorescence intensity was analyzedwith a flow cytometer. Numbers indicate the extent of M2 region that representsa population of cells with lower fluorescence intensity. (C) Effects of MAPKinhibitors on ciglitazone-induced changes in mitochondrial membrane potential.Cells were exposed to 10 μM ciglitazone for 6 h in the presence or absence ofeach 10 μM of PD98059 (PD) and SB203580 (SB). Changes in mitochondrialmembrane potential were estimated by the extent of M2 region that represents apopulation of cells with lower fluorescence intensity. Data are mean ± SEM ofthree independent experiments performed in duplicate. *P < 0.05 compared withcontrol (C); #P < 0.05 compared with ciglitazone alone (None).

Fig. 6. (A) Effects of MAPK inhibitors on ciglitazone-induced caspase-3activation. Cells were exposed to 10 μM ciglitazone for 24 h in the presence orabsence of each 10 μM of PD98059 (PD) and SB203580 (SB). Data aremean ± SEM of four independent experiments performed in duplicate. *P < 0.05compared with control; #P < 0.05 compared with ciglitazone alone. (B) Effectsof caspase inhibitors on ciglitazone-induced cell death. Cells were exposed to10 μM ciglitazone for 24 h in the presence or absence of each 20 μM of z-DEVD-FMK (FMK) and DEVD-CHO. Data are mean ± SEM of fourindependent experiments performed in duplicate. *P < 0.05 compared withciglitazone alone (None).

Fig. 7. Effects of growth factor receptor inhibitors on ciglitazone-induced celldeath. Cells were exposed to 10 μM ciglitazone for 24 h in the presence orabsence of 10 μM AG1478 (AG) and 0.3 mM suramin (Sur). Data aremean ± SEM of four independent experiments performed in duplicate. *P < 0.05compared with ciglitazone alone (None).

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agonists is unrelated to a decrease in cell viability and to thePPARγ receptor. Thus, the role of ROS in PPARγ ligand-induced cell death remains to be controversial. In the presentstudy, ciglitazone increased ROS generation before induction ofcell death, and the generation was inhibited by the antagonistGW9662 (Fig. 2), indicating that ciglitazone induces ROS

production through PPARγ receptor. The facts that radicalscavengers prevented the cell death (Fig. 3) suggest involve-ment of ROS production in the cytotoxicity of ciglitazone.

Although ciglitazone is known as a ligand of PPARγ, itseffect might be mediated by a PPARγ-independent mechanism

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because some reports suggest that 15d-PGJ2 and glitazones areable to act by a nongenomic mechanism (Chen et al., 2003;Lennon et al., 2002; Ward et al., 2002). The results of thepresent study showed that ciglitazone induces cell deaththrough a PPARγ-dependent pathway as evidenced by themarked protection of cell death with GW9662 (Fig. 3).GW9662 is an inhibitor of PPARγ with high affinity andselectivity and can fully abrogate PPARγ-dependent mecha-nism (Leesnitzer et al., 2002). PPARγ-dependent induction ofcell death by ciglitazone and 15d-PGJ2 have been reported inother cell types (Bishop-Bailey and Hla, 1999; Kim et al., 2003;Zander et al., 2002).

To elucidate the precise mechanism involved in theciglitazone-induced osteoblastic cell death, the effects ofciglitazone on MAPK activation were examined. PPARγagonists have shown to activate different MAPK subfamiliesdepending on the cell types (Gardner et al., 2003; Lennon etal., 2002; Shan et al., 2004; Teruel et al., 2003). The presentstudy showed that ciglitazone treatment caused a sustainedactivation of ERK and p38 (Figs. 4A and B). Role of MAPKsignaling pathway in cell death induced by PPARγ agonistsremains controversial. ERK activation induced by PPARγagonists mediates anti-apoptotic signal (Shan et al., 2004).However, other studies have shown that the ERK activationis involved in PPARγ-induced cell death (Kim et al., 2003;Padilla et al., 2000). Although it has been generally acceptedthat ERK activation delivers survival signals (Cobb, 1999;Xia et al., 1995), several studies have shown involvement ofERK activation in cell death induced by various stimuli(Bhat and Zhang, 1999; Choi et al., 2004; Kim et al., 2005;Lee et al., 2005). In the present study, ERK activationappears to play an important role in the ciglitazone-inducedcell death as confirmed by reduction in cell death by ERKinhibitor (Fig. 4D).

Activation of p38 induced by PPARγ agonists has beenreported to be regulated differently in various cell types. 15d-PGJ2 causes activation of this kinase in primary astrocytes(Lennon et al., 2002) but not in human leukemic cell lines(Yamakawa-Karakida et al., 2002). 15d-PGJ2 or ciglitazoneinduces activation of p38 in chondrocytes (Shan et al., 2004),liver epithelial cells (Gardner et al., 2003), and skeletal muscle(Kramer et al., 2005). In this study, ciglitazone caused p38activation and the activation was involved in the cell death (Fig.4). Our data are at variance with observations by Yasuda et al.(2005) in osteoblasts inwhich treatment of ciglitazone alone dosenot induce significant increases in activation of MAPKsubfamilies. The reason for the discrepancy is not clear at present.

Bax is a member of the Bcl-2 family of proteins and has beenassociated with apoptotic cell death both in cell culture(Hassouna et al., 1996) and in intact animals (Yin et al., 1997).During apoptosis, Bax is translocated to the mitochondria whereit promotes the release of proteins from the intermembrane spaceinto cytoplasm, an important step in apoptosis (Antonsson et al.,2000; Green and Reed, 1998). Bax interacts with thepermeability transition pore to facilitate protein movementthrough the mitochondrial membrane (Antonsson et al., 2000;Desagher andMartinou, 2000).Whether PPARγ agonists induce

cell death through a Bax-dependent mechanism remains unclear.Bax expression is not altered by troglitazone, a PPARγ agonist,in different cell types (Okura et al., 2000; Tsuchiya et al., 2003;Yamakawa-Karakida et al., 2002), whereas 15d-PGJ2 causes anincrease in Bax expression (Kim et al., 2003; Liu et al., 2005;Shen et al., 2005). Thus, the effects of PPARγ agonists may bedepend on the ligand used and/or the cell type studied. In thepresent study, ciglitazone increased Bax expression (Fig. 5A).These data are consistent with reports in human glioma cell lines(Strakova et al., 2005). Enhanced Bax expression by ciglitazonemay suggest that MPT is involved in the cell death. Indeed,exposure of cells to ciglitazone caused a loss of mitochondrialmembrane potential, and its effect was prevented by theantioxidant NAC and the MAPK inhibitors (Figs. 5B and C).

The mitochondria-dependent cell death may imply thatcaspase activation is involved in the cell death. The presentstudy showed that ciglitazone induced caspase activation, whichwas inhibited by inhibitors of ERK and p38 (Fig. 6A). Thegeneral caspase inhibitor z-DEVD-FMK and the caspase-3inhibitor DEVD-CHOprevented ciglitazone-mediated cell death(Fig. 6B), suggesting that ciglitazone induces cell death througha caspase-dependent mechanism, supporting previous findingswith PPARγ agonists in various cell types (Bruemmer et al.,2003; Piva et al., 2005; Shan et al., 2004; Strakova et al., 2005).

Ciglitazone induced activation of ERK through an EGFR-dependent mechanism as evidenced by inhibition of ERKphosphorylation by the EGFR inhibitor AG1478 (Fig. 4C).However, AG1478 did not affect the phosphorylation of p38MAPK. These data suggest that ERK and p38 activation inducesthrough two independent signaling mechanisms in osteoblasticcells. These results are consistent with those reported in rat liverepithelial cells (Gardner et al., 2003). However, whetherciglitazone induces cell death through EGFR is not clear. Inthe present study, the ciglitazone-induced cell death wasprevented by AG1478 and suramin (Fig. 7), suggestinginvolvement of EGFR in the ciglitazone-induced cell death.

In conclusion, the present study demonstrated that MAPKsignaling pathways are involved in the ciglitazone-induced celldeath of osteoblasts. Such signaling pathways lead to theactivation of the downstream proapoptotic molecule Bax andcaspases cascades.

Acknowledgment

This work was partly supported by the program of MOST/KOSEF (R13-2005-009).

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