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Originally published In Press as doi:10.1074/jbc.M502956200 on June 29, 2005

J. Biol. Chem., Vol. 280, Issue 35, 31059-31067, September 2, 2005
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Involvement of Intracellular Ca2+ Levels in Nonsteroidal Anti-inflammatory Drug-induced Apoptosis*

Ken-ichiro Tanaka{ddagger}, Wataru Tomisato{ddagger}, Tatsuya Hoshino{ddagger}, Tomoaki Ishihara{ddagger}, Takushi Namba{ddagger}, Mayuko Aburaya{ddagger}, Takashi Katsu§, Keitarou Suzuki{ddagger}, Shinji Tsutsumi{ddagger}, and Tohru Mizushima{ddagger}

From the {ddagger}Graduate School of Medical and Pharmaceutical Sciences, Kumamoto University, Kumamoto 862-0973, and the §Faculty of Pharmaceutical Sciences, Okayama University, Okayama 700-8530, Japan

Received for publication, March 17, 2005 , and in revised form, May 25, 2005.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
We recently reported that nonsteroidal anti-inflammatory drug (NSAID)-induced gastric lesions involve NSAID-induced apoptosis of gastric mucosal cells, which in turn involves the endoplasmic reticulum stress response, in particular the up-regulation of CCAAT/enhancer-binding protein homologous transcription factor (CHOP). In this study, we have examined the molecular mechanism governing this NSAID-induced apoptosis in primary cultures of gastric mucosal cells. Various NSAIDs showed membrane permeabilization activity that correlated with their apoptosis-inducing activity. Various NSAIDs, particularly celecoxib, also increased intracellular Ca2+ levels. This increase was accompanied by K+ efflux from cells and was virtually absent when extracellular Ca2+ had been depleted. These data indicate that the increase in intracellular Ca2+ levels that is observed in the presence of NSAIDs is due to the stimulation of Ca2+ influx across the cytoplasmic membrane, which results from their membrane permeabilization activity. An intracellular Ca2+ chelator partially inhibited celecoxib-induced release of cytochrome c from mitochondria, reduced the magnitude of the celecoxib-induced decrease in mitochondrial membrane potential and inhibited celecoxib-induced apoptotic cell death. It is therefore likely that an increase in intracellular Ca2+ levels is involved in celecoxib-induced mitochondrial dysfunction and the resulting apoptosis. An inhibitor of calpain, a Ca2+-dependent cysteine protease, partially suppressed mitochondrial dysfunction and apoptosis in the presence of celecoxib. Celecoxib-dependent CHOP-induction was partially inhibited by the intracellular Ca2+ chelator but not by the calpain inhibitor. These results suggest that Ca2+-stimulated calpain activity and CHOP expression play important roles in celecoxib-induced apoptosis in gastric mucosal cells.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Nonsteroidal anti-inflammatory drugs (NSAIDs)1 account for nearly 5% of all prescribed medications (1). The anti-inflammatory action of NSAIDs is mediated through their inhibitory effect on cyclooxygenase (COX) activity. COX is an enzyme essential for the synthesis of prostaglandins (PGs), which have a strong capacity to induce inflammation. On the other hand, NSAID use is associated with gastrointestinal complications (2), with about 15-30% of chronic users of NSAIDs suffering from gastrointestinal ulcers and bleeding (3-6).

The inhibition of COX by NSAIDs at gastrointestinal mucosa was previously thought to be the sole explanation for the gastrointestinal side effects of NSAIDs because PGs have a strong protective effect on gastrointestinal mucosa (7, 8). However, now it is believed that the induction of gastrointestinal lesions by NSAIDs involves additional mechanisms, since the increased incidence of gastrointestinal lesions and the decrease in PG levels induced by NSAIDs do not always occur in parallel (9, 10). We have previously demonstrated that NSAIDs induce cell death (apoptosis and necrosis) in primary cultures of gastric mucosal cells in a manner independent of COX inhibition (11, 12). Furthermore, we recently suggested that both COX inhibition and NSAID-induced cell death at gastric mucosa are required for NSAID-induced gastric lesions in vivo (12). Therefore, elucidation of the mechanisms governing NSAID-induced cell death is important for understanding the mechanisms by which NSAIDs cause gastric lesions and may allow the development of new safer NSAIDs. We recently showed that all of the NSAIDs tested have membrane permeabilization activity, which is implicated in NSAID-induced apoptosis and necrosis (13). However, the mechanism by which the membrane permeabilization activity of NSAIDs causes cell death and in particular apoptosis in gastric mucosal cells remains unknown.

Accumulation of unfolded protein in the endoplasmic reticulum (ER) induces the ER stress response, otherwise known as the unfolded protein response. Cells initially adapt to the accumulation of unfolded proteins by inducing the expression of ER-resident molecular chaperones such as glucose-regulated protein 78 (14-17). However, if this adaptation does not prove sufficient, the apoptotic response is initiated, which is primarily the induction of the CCAAT/enhancer-binding protein homologous transcription factor (CHOP) (18). We recently reported that various NSAIDs induce not only glucose-regulated protein 78 but also CHOP expression. In addition to this, we showed, using CHOP-deficient mice and a dominant negative form of CHOP, that this CHOP induction is essential for NSAID-induced apoptosis (19). Therefore, NSAIDs seem to induce apoptosis by acting as ER stressors. However, the mechanism by which NSAIDs induce the ER stress response has remained unknown. Other groups have pointed out the involvement of mitochondrial dysfunction in NSAID-induced apoptosis; some NSAIDs (such as celecoxib) stimulate the release of cytochrome c from mitochondria and decrease the mitochondrial membrane potential (20, 21). Various mechanisms have been proposed for NSAID-induced mitochondrial dysfunction, such as inactivation of phosphatidylinositol 3-kinase/3-phosphoinositide-dependent kinase-1/Akt or mitogen-activated protein kinase/extracellular signal-regulated kinase (20-22). However, data in these studies have some contradictions (may be due to differences in cell species), and the mechanisms by which NSAIDs cause mitochondrial dysfunction have also not been fully characterized.

Increases in intracellular Ca2+ levels, due to cellular Ca2+ overload or perturbation of intracellular Ca2+ compartmentalization, trigger apoptosis. For example, many apoptotic stimuli, such as activation of surface antigen receptors, increase the intracellular Ca2+ level, and compounds that directly increase the intracellular Ca2+ level (Ca2+ ionophores and inhibitors for sarcoendoplasmic reticulum Ca2+ ATPase (SERCA)) have been shown to induce apoptosis (23-25). Various mechanisms have been proposed for the Ca2+-induced apoptosis pathway, such as activation of protein kinase C, opening of permeability transition pores in mitochondria, and stimulation of reactive oxygen species synthesis (26-29). In addition, calpain, a Ca2+-dependent cysteine protease, plays an important role in the Ca2+-induced apoptosis pathway. Calpain activates or inhibits Bax and Bid or Bcl-2 and Bcl-XL, respectively, by their cleavage, resulting in stimulation of the release of cytochrome c from mitochondria and a decrease in the mitochondrial membrane potential. Then cytochrome c activates caspase-9, which in turn activates caspase-3 and induces apoptosis. It is also suggested that calpain directly activates some caspases (26, 29-32). On the other hand, recent reports show that Ca2+ ionophores induce the ER stress response and CHOP expression, suggesting that increases in the intracellular Ca2+ level can induce the ER stress response and CHOP expression (26, 33).

Permeabilization of cytoplasmic membranes causes an increase in intracellular Ca2+ levels by stimulating Ca2+ influx across the cytoplasmic membrane. Some NSAIDs were shown to increase intracellular Ca2+ levels (13, 34). Therefore, it is reasonable to speculate that increases in intracellular Ca2+ levels are involved in NSAID-induced apoptosis (i.e. that the increased intracellular Ca2+ level connects the membrane permeabilization and CHOP induction activities of NSAIDs). Furthermore, it is also possible that activation of calpain connects the increases in intracellular Ca2+ levels to mitochondrial dysfunction in the presence of NSAIDs. In this study, we showed, in primary cultures of guinea pig gastric mucosal cells, that all NSAIDs tested increased intracellular Ca2+ levels, which accompanied the induction of apoptosis. An intracellular Ca2+ chelator partially inhibited CHOP induction, release of cytochrome c from mitochondria, the decrease in mitochondrial membrane potential, and apoptotic cell death in the presence of celecoxib (the most potent NSAID for apoptosis induction). Furthermore, an inhibitor of calpain partially suppressed the mitochondrial dysfunction and apoptotic cell death but did not affect CHOP induction in the presence of celecoxib. These results suggest that the celecoxib-dependent increase in intracellular Ca2+ levels and the resulting calpain activation and CHOP induction are involved in celecoxib-induced apoptosis in gastric mucosal cells.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Chemicals, Media, and Animals—Fetal bovine serum was from Invitrogen. RPMI 1640 and Hanks' solution were obtained from Nissui Pharmaceutical Co. Pronase E and type I collagenase were purchased from Kaken Pharmaceutical Co. and Nitta Gelatin Co., respectively. Pluronic F127, fluo-3/AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA-AM), and BAPTA were from Dojindo Co. Nimesulide and flurbiprofen were from Cayman Chemical Co. Indomethacin was from Wako Co. Hoechst 33342 (Ho 342), ibuprofen, diclofenac, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), mefenamic acid, ketoprofen, and flufenamic acid were from Sigma. 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidaozolylcarbocyanine iodide (DePsipher) was from Trevigen. Celecoxib was from LKT Laboratories Inc. Etodolac was a gift kindly provided by Nippon Shinyaku Co. Peptides for the assay of caspase-3 or calpain and carbobenzoxy-L-leucyl-L-leucinal (Z-Leu-Leu-H) were from Peptide Institute, Inc. Egg phosphatidylcholine was from Kanto Chemicals Co. Antibodies against CHOP and actin were from Santa Cruz Biotechnology Inc. Antibody against cytochrome c was from PharMingen. Male guinea pigs weighing 200-300 g were purchased from Kyudo Co. The experiments and procedures described here were carried out in accordance with the Guide for the Care and Use of Laboratory Animals as adopted and promulgated by the National Institutes of Health and were approved by the Animal Care Committee of Kumamoto University.

In Vitro Assay of Cytotoxicity—Gastric mucosal cells were isolated from guinea pig fundic glands as described previously (35). Isolated gastric mucosal cells were cultured for 12 h in RPMI 1640 containing 0.3% (v/v) fetal bovine serum, 100 units/ml penicillin, and 100 µg/ml streptomycin in type I collagen-coated plastic culture plates under conditions of 5% CO2, 95% air and 37 °C. Nonadherent cells were removed, and the cells attached to the plate were used in the experiments. Guinea pig gastric mucosal cells prepared under these conditions have previously been characterized, with the majority (about 90%) of cells identified as pit cells (35).

NSAIDs were dissolved in Me2SO or Na2CO3 (for indomethacin only) and control experiments (without NSAIDs) were performed in the presence of the same concentrations of Me2SO or Na2CO3. Cells were exposed to NSAIDs by changing the medium. Cell viability was determined by the MTT method.

Apoptotic chromatin condensation was monitored as described previously (36). Cells were washed with PBS, stained with 10 µg/ml Ho 342, and observed under a fluorescence microscope.

Immunoblotting Analysis—Whole cell extracts were prepared as described previously (36). The protein concentration of the samples was determined by the Bradford method. The samples were electrophoresed on polyacrylamide gels containing SDS, and the proteins were then transferred to membranes and detected using antibodies.

Mitochondrial Membrane Potential Assay—Mitochondrial membrane potential was assayed using a fluorometric mitochondrial permeability assay kit (Trevigen) (36). Briefly, after treatment with NSAIDs, cells were treated with DePsipher (5 µg/ml) for 20 min at 37 °C and observed under a fluorescence microscope using 590 nm for red emission and 530 nm for green emission.

Measurement of the Intracellular Ca2+ Level—Intracellular Ca2+ levels were monitored as described (37). The cells were washed with assay buffer (115 mM NaCl, 5.4 mM KCl, 1.8 mM CaCl2, 0.8 mM MgCl2, 20 mM HEPES, and 13.8 mM glucose). For Ca2+-free conditions, the modified assay buffer (115 mM NaCl, 5.4 mM KCl, 5 mM EGTA, 20 mM HEPES, and 13.8 mM glucose) was used instead of the normal assay buffer. The cells were then incubated with 4 µM fluo-3/AM in assay buffer supplemented with 0.1% bovine serum albumin, 0.04% pluronic F127, and 2 mM probenecid, for 40 min at 37 °C. After washing twice with assay buffer, cells were suspended in assay buffer supplemented with 2 mM probenecid. Cells were transferred to a water-jacketed cuvette (1.6 x 106 cells/cuvette), and the fluo-3 fluorescence was then measured with a Hitachi F-4500 spectrofluorophotometer by recording excitation signals at 490 nm and the emission signal at 530 nm at 1-s intervals. Maximum and minimum fluorescence values (Fmax and Fmin) were obtained by adding 10 µM ionomycin and 10 µM ionomycin under Ca2+-free conditions, respectively. The intracellular Ca2+ level was calculated according to the equation, [Ca2+]i = Kd(F - Fmin)/(Fmax - F), where Kd is the apparent dissociation constant (400 nM) of the fluorescent dye-Ca2+ complex.

Membrane Permeability Assay—Permeabilization of calcein-loaded liposomes was assayed as described previously (13). Liposomes were prepared using the reversed-phase evaporation method. Egg phosphatidylcholine (10 µmol, 7.7 mg) was dissolved in chloroform/methanol (1:2, v/v), dried, dissolved in 1.5 ml of diethyl ether, and added to 1 ml of 100 mM calcein-NaOH (pH 7.4). The mixture was sonicated to obtain a homogenous emulsion. The diethyl ether solvent was removed, and the resulting suspension of liposomes was centrifuged and washed twice with fresh buffer A (10 mM potassium buffer, containing 150 mM NaCl) to remove untrapped calcein. The final liposome precipitate was resuspended in 5 ml of buffer A. A 0.3-ml aliquot of this suspension was diluted with 19.7 ml of buffer A, and 400 µl of this diluted suspension was then incubated at 30 °C for 10 min in the presence of the NSAID under investigation. The release of calcein from liposomes was determined by measuring fluorescence intensity at 520 nm (excitation at 490 nm).

Assay for K+ Efflux from Cells—K+ efflux from cells was monitored as described previously (38) with some modifications. Cells were washed twice with buffer A and then suspended in fresh buffer A (6 x 106 cells/ml). After incubation with NSAIDs for 30 min at 37 °C, K+ efflux from cells was measured with a K+ ion-selective electrode.

Assay for Caspase Activity—The activity of caspase-3 was determined as described previously (36). Briefly, cells were collected by centrifugation and suspended in extraction buffer (50 mM PIPES (pH 7.0), 50 mM KCl, 5 mM EGTA, 2 mM MgCl2, and 1 mM dithiothreitol). Suspensions were sonicated and centrifuged, after which the supernatants were incubated with fluorogenic peptide substrates (acetyl-DEVD-methylcoumarin amide) in reaction buffer (100 mM HEPES-KOH (pH 7.5), 10% sucrose, 0.1% CHAPS, and 1 mg/ml bovine serum albumin) for 15 min at 37 °C. The release of aminomethylcoumarin (AMC) was determined using a fluorescence spectrophotometer. One unit of protease activity was defined as the amount of enzyme required to release 1 pmol of AMC/min.

Assay for Calpain Activity—The activity of calpain was determined as described previously (30). Briefly, cells were collected by centrifugation, washed by Hanks' solution, and suspended with the same solution at 2.5 x 105 cells/ml. Suspensions were incubated with fluorogenic peptide substrates succinyl-L-leucyl-L-leucyl-L-valyl-L-tyrosine 4-methylcoumaryl-7-amide for 3 min at 37 °C. The release of AMC was determined using a fluorescence spectrophotometer.

Statistical Analyses—All values are expressed as the means ± S.E. One-way analysis of variance, followed by Scheffe's multiple comparison, was used for evaluation of differences between the groups. Student's t test for unpaired results was performed to evaluate differences between two groups. Differences were considered to be significant for values of p < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Close Relationship between NSAID-induced Apoptosis and Membrane Permeabilization—We recently reported that some NSAIDs (celecoxib, mefenamic acid, flufenamic acid, nimesulide, and flurbiprofen) cause not only apoptosis in primary cultures of guinea pig gastric mucosal cells but also membrane permeabilization in calcein-loaded liposomes (13). To examine the relationship between the apoptosis-inducing and membrane permeabilization activities of NSAIDs, in this study, we have examined these activities of other NSAIDs. As shown in Fig. 1A, treatment of primary cultures of guinea pig gastric mucosal cells with celecoxib for 16 h decreased cell viability in a dose-dependent manner, and this is consistent with our previous results (13). Each of the NSAIDs tested here (indomethacin, diclofenac, etodolac, ibuprofen, and ketoprofen) also decreased cell viability in a dose-dependent manner. Because cell death under these conditions was accompanied by apoptotic DNA fragmentation and chromatin condensation (data not shown), it is most likely mediated by apoptosis.

Two subtypes of COX, COX-1 and COX-2, are responsible for the majority of COX activity in gastric mucosal and inflammatory tissues, respectively, and recently a number of COX-2-selective NSAIDs have been developed (39). Among the NSAIDs tested in Fig. 1A, etodolac and celecoxib have selectivity for COX-2. No relationship was evident between NSAID-induced apoptosis and selectivity for COX-2 (Fig. 1A). We also confirmed that exogenously added PGE2 (either native PGE2 or 16,16-dimethyl-PGE2) did not affect NSAID-induced apoptosis even at a higher concentration of PGE2 in the culture medium than is present endogenously (10-9 M) (data not shown). These data show that NSAID-induced apoptosis is independent of COX-inhibition by NSAIDs and are consistent with our previous conclusion (12, 13).



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  		FIG. 1.
Apoptosis induced by NSAIDs. Cultured guinea pig gastric mucosal cells were incubated with the indicated concentrations of each NSAID for 16 h (A) or with 100 µM celecoxib for the indicated periods (B). Cell viability was determined by the MTT method. Values are mean ± S.E. (n = 3).

 
Calcein fluoresces very weakly at high concentrations due to self-quenching, so the addition of membrane-permeabilizing drugs to a medium containing calcein-loaded liposomes should cause an increase in fluorescence by diluting out the calcein (13). As shown in Fig. 2A, not only celecoxib but also other NSAIDs increased the calcein fluorescence, suggesting that they have membrane permeabilization activity.

Combining the results from the previous (13) and present studies, we obtained dose-response curves of 10 different NSAIDs for both induction of apoptosis and membrane permeabilization. To examine the relationship between NSAID-induced apoptosis and membrane permeabilization, we determined ED50 values of the 10 NSAIDs for apoptosis (concentrations of NSAIDs required for 50% inhibition of cell viability by apoptosis) and ED20 values for membrane permeabilization (concentration of NSAIDs required for 20% release of calcein) (Table I). We used ED20 values instead of ED50 values for estimating the activity of each NSAID for membrane permeabilization, because ED50 values of etodolac for calcein release could not to be determined (Fig. 2A). Plotting ED50 values for apoptosis versus ED20 values for membrane permeabilization (calcein release) yielded an r2 value of 0.929 (Fig. 3A), which suggests that NSAID-induced apoptosis is mediated by their ability to permeabilize membranes.


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  		TABLE I
NSAID concentrations required for apoptosis, membrane permeabilization, and Ca2+ increase ED50 values of NSAIDs for apoptosis (concentrations of NSAIDs required for 50% inhibition of cell viability by apoptosis) were calculated based on results provided in Fig. 1 in the present study and Fig. 2B in our previous study (13). ED20 values of NSAIDs for calcein release (concentrations of NSAIDs required for 20% release of calcein) were calculated based on results provided in Fig. 2 in the present study and Fig. 4 in our previous study (13). EDdouble values of NSAIDs for Ca2+ increase (concentrations of NSAIDs required for 2-fold increase in the intracellular Ca2+ level) were from Fig. 4 in the present study. ND, not determined.

 
Mechanism whereby the Intracellular Ca2+ Level Is Increased by NSAIDs—Fig. 4A shows the effect of various NSAIDs on the intracellular Ca2+ level. Not only celecoxib but also all of the other NSAIDs tested increased the intracellular Ca2+ level in a dose-dependent manner. Since indomethacin absorbs fluo-3 fluorescence (530 nm), we could not measure the intracellular Ca2+ level in the presence of indomethacin by this assay system. In order to estimate the activity of each NSAID for Ca2+ increase, we determined the concentration of each NSAID required for 2-fold increase in the intracellular Ca2+ level based on results in Fig. 4A (EDdouble value for Ca2+ increase in Table I). Compared with ED20 values for membrane permeabilization (calcein release), etodolac showed a very high EDdouble value for Ca2+ increase (Table I). Plotting the EDdouble value for Ca2+ increase versus ED20 values for membrane permeabilization (calcein release) of NSAIDs other than etodolac and indomethacin yielded an r2 value of 0.738 (Fig. 3B), which suggests that increase in the intracellular Ca2+ level by most of NSAIDs is mediated through their ability to permeabilize membranes. The membrane permeabilizing activity of etodolac may cause apoptosis independently of the Ca2+ increase; however, the mechanism is unknown at present. As well as showing the greatest potency for induction of apoptosis and membrane permeabilization (Figs. 1 and 2) (13), celecoxib increased intracellular Ca2+ levels more potently than the other NSAIDs (Fig. 4A). We therefore selected celecoxib for use in subsequent experiments.



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  		FIG. 2.
Membrane permeabilization by NSAIDs. Calcein-loaded liposomes were incubated for 10 min with the indicated concentrations of each NSAID (A) or with 100 µM celecoxib for the indicated periods (B). The release of calcein from liposomes was determined by measuring fluorescence intensity. Melittin (10 µM) was used to determine the 100% level of membrane permeabilization (38).

 



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  		FIG. 3.
Relationship between apoptosis-inducing, membrane permeabilization, and Ca2+-increasing activities of NSAIDs. ED20 values for membrane permeabilization (calcein release), ED50 values for apoptosis and EDdouble values for Ca2+ increase are from Table I. All 10 NSAIDs and NSAIDs except etodolac and indomethacin are plotted in A and B, respectively.

 



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  		FIG. 4.
Changes in the intracellular Ca2+ level induced by NSAIDs. The intracellular Ca2+ level was monitored using a fluo-3/AM assay system. Indicated concentrations of each NSAID were added to fluo-3/AM-loaded cells, and the time course of fluo-3 fluorescence change was monitored. The maximum values for the intracellular Ca2+ level ([Ca2+]i) are shown (A). The time course of fluo-3 fluorescence change in the presence of 100 µM celecoxib is shown (B).

 
In order to reveal the mechanism by which intracellular Ca2+ levels are increased by celecoxib, we monitored intracellular Ca2+ levels under Ca2+-free conditions (see "Experimental Procedures"). As shown in Fig. 5A, the increase in the intracellular Ca2+ level induced by 50-100 µM celecoxib (apoptosis-inducing conditions) was drastically inhibited under Ca2+-free conditions, suggesting that most of the increase is derived from extracellular Ca2+ and not from intracellular compartments (such as the ER and mitochondria). In other words, stimulation of Ca2+ influx across the cytoplasmic membrane due to membrane permeabilization by celecoxib seems to be responsible for the increase in the intracellular Ca2+ level in the presence of 50-100 µM celecoxib. Since permeabilization of cytoplasmic membranes should stimulate K+ efflux from cells, we examined the effect of celecoxib on K+ concentrations in the medium. As shown in Fig. 5C, K+ concentration in the medium was increased, depending on the dose of celecoxib, suggesting that celecoxib stimulated K+ efflux across the cytoplasmic membranes. These data suggest that celecoxib (50-100 µM) makes the cytoplasmic membranes permeable and stimulates Ca2+ influx, resulting in the observed increase in the intracellular Ca2+ level.



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  		FIG. 5.
Mechanism for celecoxib-induced increases in the intracellular Ca2+ level. Indicated concentrations of celecoxib were added to fluo-3/AM-loaded cells under Ca2+-containing (+) or Ca2+-free (-) conditions (A and B). Fluo-3/AM-loaded cells were pretreated with 1 µM thapsigargin for 10 min before the celecoxib treatment (B). The maximum value for the intracellular Ca2+ level ([Ca2+]i) was determined as described in the legend to Fig. 4 (A and B). Cultured guinea pig gastric mucosal cells were incubated with the indicated concentrations of celecoxib for 30 min (C and D). Cells were pretreated with BAPTA-AM for 10 min before the celecoxib treatment (D). The level of K+ efflux was measured using a K+ ion-selective electrode. Melittin (10 µM) was used to determine the 100% level of K+ efflux (38). Values are mean ± S.E. (n = 3). ***, p < 0.001; n.s., not significant (C and D).

 
Since the increase in intracellular Ca2+ levels by 10-20 µM celecoxib was not clearly inhibited under Ca2+-free conditions (Fig. 5, A and B), a mechanism other than stimulation of Ca2+ influx across the cytoplasmic membranes seems to be involved. We examined the contribution of the ER Ca2+ pool to the celecoxib-induced increase in the intracellular Ca2+ levels. Cells were pretreated with thapsigargin, an inhibitor of SERCA, which pumps Ca2+ from the cytoplasm to the ER, and then treated with celecoxib under either Ca2+-containing or Ca2+-free conditions (Fig. 5B). This pretreatment with thapsigargin depleted the ER Ca2+ pool (data not shown). The increase in the intracellular Ca2+ level induced by 10-20 µM celecoxib was inhibited more drastically by this thapsigargin pretreatment than by the depletion of extracellular Ca2+ (Fig. 5B). This contrasts with the results for 50-100 µM celecoxib (Fig. 5B). These results suggest that the ER Ca2+ pool (and perhaps permeabilization of ER membrane) plays an important role in the increase in the intracellular Ca2+ level induced by 10-20 µM celecoxib. In other words, the results indicate that most of the increase in intracellular Ca2+ levels that is induced by 10-20 µM celecoxib is derived from the ER and not from extracellular sources. A previous report showed that celecoxib inhibits SERCA (34). Therefore, the inhibitory action of celecoxib on SERCA may also contribute to the increase in the intracellular Ca2+ level induced by 10-20 µM celecoxib. In the presence of various concentrations of celecoxib, pretreatment with thapsigargin under Ca2+-free conditions caused almost complete inhibition of the celecoxib-induced increase in intracellular Ca2+ levels (Fig. 5B). Therefore, both stimulation of Ca2+ influx across cytoplasmic membranes and depletion of the ER Ca2+ pool are responsible for the celecoxib-dependent increase in intracellular Ca2+ levels, which is consistent with previous results (34, 40).

Contribution of the Increase in the Intracellular Ca2+ Level to Celecoxib-induced Apoptosis—In order to examine the contribution of the membrane permeabilization and the resulting increase in intracellular Ca2+ level to celecoxib-induced apoptosis, at first, we examined the kinetics of celecoxib-induced apoptosis, membrane permeabilization, and increase in intracellular Ca2+ level (Figs. 1B, 2B, and 4B). Comparing to induction of apoptosis by celecoxib, the membrane permeabilization (calcein release) and increase in intracellular Ca2+ level by celecoxib occurred very rapidly, suggesting that the membrane permeabilization and the resulting increase in intracellular Ca2+ level are located upstream of the induction of apoptosis.

We then examined induction of apoptosis under Ca2+-free conditions that is achieved by the addition of BAPTA, Ca2+ chelator, to the culture medium. As shown in Fig. 6A, the activation of caspase-3-like activity in cells treated with celecoxib for 6 h was partially suppressed by the addition of BAPTA, suggesting that the increase in intracellular Ca2+ level by celecoxib is important for celecoxib-induced apoptosis. However, depletion of Ca2+ in medium was toxic to cells; treatment of cells with 1 mM BAPTA for 16 h or incubation of cells with Ca2+-free medium decreased the cell viability even in the absence of celecoxib (data not shown). Furthermore, since BAPTA is not permeable for cytoplasmic membranes, BAPTA cannot chelate intracellular Ca2+. In order to address these issues, we used BAPTA-AM, an intracellular Ca2+ chelator that is permeable for cytoplasmic membranes. As shown in Fig. 6B, BAPTA-AM partially inhibited celecoxib-induced cell death. At the concentrations used, BAPTA-AM did not affect cell viability in the absence of celecoxib (data not shown). Furthermore, treatment of cells with celecoxib caused apoptotic chromatin condensation, which was also partially inhibited by the addition of BAPTA-AM in culture medium (Fig. 6, D and E). Therefore, the increase in intracellular Ca2+ levels caused by celecoxib seems to be involved in celecoxib-induced apoptosis. BAPTA-AM did not affect the celecoxib-dependent stimulation of K+ efflux (Fig. 5D), suggesting that the membrane permeabilization by celecoxib is located upstream of increase in intracellular Ca2+ level and induction of apoptosis by celecoxib.

We also examined mitochondrial dysfunction in the presence of celecoxib. As shown in Fig. 7A, treatment of cells with celecoxib increased the amount of cytochrome c in the cytosolic fractions, suggesting that celecoxib stimulated the release of cytochrome c from mitochondria. The addition of BAPTA-AM partially suppressed this increase in the amount of cytochrome c (Figs. 7, A and B), suggesting that the celecoxib-dependent release of cytochrome c from mitochondria is partly due to the increase in the intracellular Ca2+ level.



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  		FIG. 6.
Effect of Ca2+ chelators or an inhibitor of calpain on celecoxib-induced apoptosis. Cultured guinea pig gastric mucosal cells were preincubated with indicated concentrations of BAPTA-AM (B, D, and E) or Z-Leu-Leu-H (C-E) for 1 h. Cells were further incubated with or without 100 µM celecoxib for 6 h (A) or 16 h (B-E) under the same conditions as the preincubation step (medium containing BAPTA was used for A). Caspase-3-like activity was measured using fluorogenic peptide substrates (acetyl-DEVD-methylcoumarin amide) (A). Cell viability was determined by the MTT method (B and C). Values are mean ± S.E. (n = 3). ***, p < 0.001; **, p < 0.01 (A-C). After staining with Ho 342, cells were observed using a fluorescence microscope. The arrows show condensed chromatin (D). The numbers of apoptotic cells with condensed chromatin were counted from representative photomicrographs and are expressed as a percentage of total cell number (n = 100). Values are mean ± S.E. (n = 3). ***, p < 0.001 (E).

 
We also examined mitochondrial dysfunction by monitoring the mitochondrial membrane potential, using DePsipher, a mitochondrial dye. This mitochondrial dye normally exists in solution as a monomer emitting green fluorescence (530 nm) and forms a dimer emitting red fluorescence (590 nm) in a reaction that is driven by the mitochondrial membrane potential. Thus, a decrease in the mitochondrial membrane potential should reduce the red fluorescence. As shown in Fig. 7E, the red fluorescence was apparent in control cells (without celecoxib) but was markedly decreased in celecoxib-treated cells. However, when cells were treated with celecoxib in the presence of BAPTA-AM, the red fluorescence was partially recovered (Fig. 7, E and F), suggesting that the celecoxib-dependent decrease in mitochondrial membrane potential is partly due to the celecoxib-induced increase in the intracellular Ca2+ level. These results show that the observed increase in the intracellular Ca2+ level is involved in celecoxib-induced mitochondrial dysfunction.

Mechanism for Ca2+-dependent Celecoxib-induced Apoptosis—Then we examined the pathway for celecoxib-induced apoptosis downstream of the increase in the intracellular Ca2+ level. First, we examined the contribution of the intracellular Ca2+ level to celecoxib-induced CHOP expression, which was shown to play an important role in NSAID-induced apoptosis (19). Treatment of cells with celecoxib increased the amount of CHOP, as was previously reported (19) (Fig. 8, A and B). However, when cells were treated with celecoxib in the presence of BAPTA-AM, this CHOP induction was partially inhibited, suggesting that CHOP induction by celecoxib is partly due to the celecoxib-induced increase in the intracellular Ca2+ level.

We also examined the participation of calpain in celecoxib-induced apoptosis. We found that the activity of calpain was increased in celecoxib-treated cells by use of cell-permeable calpain substrate (Fig. 9). This calpain activation was partially inhibited by pretreatment of cells with BAPTA-AM (Fig. 9), suggesting that this activation is partially due to an increase in the intracellular Ca2+ level in the presence of celecoxib. An inhibitor of calpain, Z-Leu-Leu-H, partially suppressed the celecoxib-dependent decrease in cell viability and apoptotic chromatin condensation (Fig. 6, C-E). At the concentrations used, Z-Leu-Leu-H did not affect cell viability in the absence of celecoxib (data not shown). These results suggest that calpain is involved in NSAID-induced apoptosis. As shown in Fig. 7, C-F, both the celecoxib-stimulated release of cytochrome c from mitochondria and the celecoxib-dependent decrease in mitochondrial membrane potential were also partially suppressed by the addition of Z-Leu-Leu-H. From these observations, it is likely that calpain also plays an important role in celecoxib-dependent mitochondrial dysfunction. In other words, activation of calpain by the celecoxib-induced increase in the intracellular Ca2+ level seems to damage the mitochondria. On the other hand, in contrast to BAPTA-AM, addition of Z-Leu-Leu-H did not significantly affect CHOP induction in the presence of celecoxib (Fig. 8, C and D), suggesting that induction of CHOP by celecoxib is independent of calpain.



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  		FIG. 7.
Effect of an intracellular Ca2+ chelator or an inhibitor of calpain on celecoxib-dependent mitochondrial dysfunction. Cultured guinea pig gastric mucosal cells were preincubated with indicated concentrations of BAPTA-AM (A, B, E, and F) or Z-Leu-Leu-H (C-F) for 1 h. Cells were further incubated with or without 100 µM celecoxib for 16 h under the same conditions as the preincubation step. After subcellular fractionation, cytosolic fractions were analyzed by immunoblotting with an antibody against cytochrome c or actin (A and C). After normalization by the use of actin bands, the amounts of cytochrome c relative to control (without celecoxib and BAPTA-AM or Z-Leu-Leu-H) were determined and expressed as intensity. Values are mean ± S.E. (n = 3). *, p < 0.05 (B and D). Mitochondrial membrane potential was analyzed as described under "Experimental Procedures" (E). The numbers of red-stained cells with DePsipher were counted from representative photomicrographs and are expressed as a percentage of total cell number (n = 100). Values are mean ± S.E. (n = 3). ***, p < 0.001 (F).

 



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  		FIG. 8.
Effect of an intracellular Ca2+ chelator or an inhibitor of calpain on celecoxib-dependent CHOP induction. Cultured guinea pig gastric mucosal cells were preincubated with indicated concentrations of BAPTA-AM (A and B) or Z-Leu-Leu-H (C and D) for 1 h. Cells were further incubated with or without 100 µM celecoxib under the same conditions as the preincubation step. Whole cell extracts were analyzed by immunoblotting with an antibody against CHOP or actin (A and C). The intensity of the actin bands was used to correct for differences in sample loading, and the relative amounts of CHOP to control (with celecoxib but without BAPTA-AM or Z-Leu-Leu-H) are expressed as intensity. Values are mean ± S.E. (n = 3). **, p < 0.01; *, p < 0.05; n.s., not significant (B and D).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this study, we aimed to identify the pathway for celecoxib-induced apoptosis using primary cultures of gastric mucosal cells. First, using 10 different NSAIDs, we demonstrated the close relationship between their apoptosis-inducing and membrane permeabilization activities. Therefore, it seems that membrane phospholipids are the primary target of NSAIDs for the induction of apoptosis. To investigate the mechanism by which the membrane permeabilization activity of NSAIDs causes apoptosis, we focused on the intracellular Ca2+ level, because permeabilization of cytoplasmic membranes increases the intracellular Ca2+ level by stimulating Ca2+ influx, and the intracellular Ca2+ level is a key regulator of apoptosis. We have concluded that intracellular Ca2+ levels are involved in celecoxib-induced apoptosis, since each of the NSAIDs that we tested increased the intracellular Ca2+ level, this Ca2+-increasing activity of NSAIDs (except etodolac) correlated with their membrane permeabilization activity, and an intracellular Ca2+ chelator (BAPTA-AM) partially inhibited celecoxib-dependent apoptosis and mitochondrial dysfunction. To elucidate the mechanism by which the increase in the intracellular Ca2+ level causes apoptosis, we focused on calpain, because Ca2+ activates this enzyme, and activation of this enzyme can induce apoptosis. We have demonstrated the involvement of calpain in celecoxib-induced apoptosis by showing that an inhibitor of calpain (Z-Leu-Leu-H) partially inhibited celecoxib-dependent apoptosis and mitochondrial dysfunction. Cleavage by calpain activates and inhibits Bax and Bid and Bcl-2 and Bcl-XL, respectively. Since Bax and Bid cause mitochondrial membrane permeabilization (stimulate release of cytochrome c from the mitochondria and decrease the mitochondrial membrane potential) (25, 30-32), whereas Bcl-2 and Bcl-XL protect mitochondrial membrane from permeabilization, activation of calpain by increasing the intracellular Ca2+ level seems to play an important role in celecoxib-induced mitochondrial dysfunction. Calpain cleaves and thereby activates procaspase-12 and apoptosis-inducing factor and cleaves and thereby inhibits the X-linked inhibitor of apoptosis (26, 27, 41). These events may be involved in celecoxib-induced apoptosis.



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  		FIG. 9.
Activation of calpain by celecoxib. Cultured guinea pig gastric mucosal cells were preincubated with or without 1 µM BAPTA-AM for 1 h. Cells were further incubated with or without 100 µM celecoxib for 16 h under the same conditions as the preincubation step. Calpain activity was measured using fluorogenic peptide substrates (succinyl-L-leucyl-L-leucyl-L-valyl-L-tyrosine 4-methylcoumaryl-7-amide). Values are mean ± S.E. (n = 3). ***, p < 0.001; *, p < 0.05.

 
Suppression of celecoxib-induced apoptosis by BAPTA-AM was partial, suggesting that a Ca2+-independent pathway is also involved in this process. One candidate is the death receptor signaling pathway. It was recently reported that celecoxib induces expression of death receptors, especially DR5, and small interfering RNA for DR5 partially suppresses celecoxib-induced apoptosis in lung cancer cells (42). Induction of death receptors causes activation of caspase-8, and it has been shown that celecoxib activates caspase-8 and that an inhibitor of caspase-8 partially suppresses celecoxib-dependent apoptosis (42). We previously reported that various NSAIDs activate caspase-8 in cultured gastric mucosal cells (11). This DR5-dependent pathway may be involved in celecoxib-induced apoptosis in cultured gastric mucosal cells. As described in the Introduction, inactivation of phosphatidylinositol 3-kinase/3-phosphoinositide-dependent kinase-1/Akt and mitogen-activated protein kinase/extracellular signal-regulated kinase signal transduction systems may also be involved in NSAID-induced apoptosis (20-22). However, at present, it is not clear whether these pathways are Ca2+-dependent or Ca2+-independent, because upstream signals for these systems have not yet been revealed.

Suppression of celecoxib-induced apoptosis by Z-Leu-Leu-H was also partial, and this implies that a calpain-independent pathway is involved in this process. Of course, logically, the Ca2+-independent pathways discussed above may be responsible for the calpain-independent pathway. However, we have considered the contribution of Ca2+-dependent but calpain-independent pathways to celecoxib-induced apoptosis. One candidate for such a pathway is the ER stress response, in particular the induction of CHOP. We recently reported that NSAID-induced apoptosis is suppressed in cultured guinea pig gastric mucosal cells by expression of the dominant negative form of CHOP and in peritoneal macrophages from CHOP-deficient mice (19). In the current study, we have shown that celecoxib-dependent CHOP induction is suppressed by BAPTA-AM but not by Z-Leu-Leu-H, suggesting that in the celecoxib-induced apoptosis pathway, CHOP induction is located downstream of the increase in the intracellular Ca2+ level but not of calpain activation. Furthermore, various Ca2+-dependent but calpain-independent signal pathways other than CHOP induction may be involved in celecoxib-induced apoptosis. For example, Ca2+-dependent activation of protein kinase C{delta}, a protein kinase C isoform, induces release of cytochrome c from mitochondria (43). Calcineurin, a Ca2+-dependent serine-threonine phosphatase, dephosphorylates BAD (a proapoptotic member of the Bcl-2 family) and stimulates apoptosis (44). Opening of permeability transition pores in mitochondria and stimulation of reactive oxygen species synthesis are also possible Ca2+-dependent but calpain-independent pathways for celecoxib-induced apoptosis (26-29).

We recently proposed that both COX inhibition at gastric mucosa and direct gastric mucosal cell damage (such as induction of apoptosis) by NSAIDs are required for the production of gastric lesions in vivo. Gastric lesions develop in a manner that depends on both an intravenously administered low dose of indomethacin (inhibition of COX activity at the gastric mucosa without direct gastric mucosal cell damage) and an orally administered COX-2-selective NSAID, such as celecoxib (causes direct gastric mucosal cell damage without inhibition of COX at gastric mucosa) (12). Therefore, based on the results in this study, it may be suggested that NSAIDs without membrane-permeabilizing activity and intracellular Ca2+ level-increasing activity may have reduced gastrointestinal side effects compared with other NSAIDs. An issue that was recently raised concerning the use of COX-2-selective NSAIDs, a class of gastrointestinally safe NSAIDs, is their potential risk for promoting cardiovascular thrombotic events (45). Prostacyclin, a potent anti-aggregator of platelets and a vasodilator, is mainly produced by COX-2 in vascular endothelial cells, whereas thromboxane A2, a potent aggregator of platelets and a vasoconstrictor, is mainly produced by COX-1 in platelets (46, 47). Until recently, rofecoxib and celecoxib were leading COX-2-selective NSAIDs in the market. Rofecoxib was withdrawn from the market due to the risk of its promoting cardiovascular thrombotic events, and the United States Food and Drug Administration advised physicians to consider alternatives to celecoxib due to the risk of its causing cardiovascular thrombotic events (48, 49). Based on our findings, NSAIDs that do not exhibit membrane permeabilization activity may be safe for the gastrointestinal tract even if they are not highly selective for COX-2. This type of NSAID may be of clinical benefit, because they are predicted to be safe for both the gastrointestinal tract and cardiovascular system.

On the other hand, much attention has also been given to the direct cytotoxicity of NSAIDs in relation to cancer therapy. Epidemiological studies have shown that prolonged use of aspirin or other NSAIDs reduces the risk of cancer (50, 51). Furthermore, several NSAIDs are presently under clinical development as anti-cancer drugs. A number of in vivo and in vitro studies have revealed that NSAIDs cause apoptosis (or stimulate apoptosis by anti-cancer reagents) in cancer cells and that these activities of NSAIDs are involved in their anti-cancer activity (52). PGs, such as PGE2, inhibit apoptosis (53). Furthermore, overexpression of COX-2 has been reported in various tumor cells and tissues (54, 55). Therefore, the inhibition of COX by NSAIDs was previously thought to be solely responsible for their chemopreventive effect. However, several lines of evidence suggest that chemoprevention by NSAIDs also involves COX-independent mechanisms. Sulindac sulfone, a derivative of the NSAID sulindac, does not inhibit COX activity and has been shown to display anti-tumor activity in vivo as well as induce apoptosis and inhibit cell growth in tumor cells in vitro (56, 57). Moreover, NSAID induction of apoptosis and inhibition of cell growth in COX-null fibroblasts and tumor cells in which COX expression was absent have been reported (58, 59). Therefore, an understanding of COX-independent mechanisms for the induction of apoptosis by NSAIDs is very important in order to develop more effective NSAIDs for cancer therapy. If the results presented here can be applied to the case of cancer cells, NSAIDs with more potent membrane-permeabilizing and Ca2+-increasing activity may have more potent anti-cancer effects than other NSAIDs.


   FOOTNOTES
 
* This work was supported by grants-in-aid for scientific research from the Ministry of Health, Labour, and Welfare of Japan as well as by the Suzuken Memorial Foundation, the Tokyo Biochemical Research Foundation, the Kumamoto Technology and Industry Foundation, and the Japan Research Foundation for Clinical Pharmacology. 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

To whom correspondence should be addressed: Graduate School of Medical and Pharmaceutical Sciences, Kumamoto University, Kumamoto 862-0973, Japan. Tel./Fax: 81-96-371-4323; E-mail: mizu{at}gpo.kumamoto-u.ac.jp.

1 The abbreviations used are: NSAIDs, nonsteroidal anti-inflammatory drugs; COX, cyclooxygenase; PG, prostaglandin; PGE2, prostaglandin E2; ER, endoplasmic reticulum; CHOP, CCAAT/enhancer-binding protein homologous transcription factor; SERCA, sarcoendoplasmic reticulum Ca2+-ATPase; BAPTA-AM, 1,2-bis(2-aminophenoxy)ethane-N,N,N'N'-tetraacetic acid; Ho 342, Hoechst 33342; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; DePsipher, 5,5',6,6'-tetrachloro-1, 1',3,3'-tetraethylbenzimidaozolylcarbocyanine iodide; Z-Leu-Leu-H, carbobenzoxy-L-leucyl-L-leucinal; AMC, aminomethylcoumarin; PIPES, 1,4-piperazinediethanesulfonic acid; CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid. Back



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Smalley, W. E., Ray, W. A., Daugherty, J. R., and Griffin, M. R. (1995) Am. J. Epidemiol. 141, 539-545[Abstract/Free Full Text]
  2. Hawkey, C. J. (2000) Gastroenterology 119, 521-535[CrossRef][Medline] [Order article via Infotrieve]
  3. Barrier, C. H., and Hirschowitz, B. I. (1989) Arthritis Rheum. 32, 926-932[Medline] [Order article via Infotrieve]
  4. Gabriel, S. E., Jaakkimainen, L., and Bombardier, C. (1991) Ann. Intern. Med. 115, 787-796[Abstract/Free Full Text]
  5. Fries, J. F., Miller, S. R., Spitz, P. W., Williams, C. A., Hubert, H. B., and Bloch, D. A. (1989) Gastroenterology 96, 647-655[Medline] [Order article via Infotrieve]
  6. Kurata, J. H., and Abbey, D. E. (1990) J. Clin. Gastroenterol. 12, 260-266[Medline] [Order article via Infotrieve]
  7. Vane, J. R., and Botting, R. M. (1996) Scand. J. Rheumatol. Suppl. 102, 9-21[Medline] [Order article via Infotrieve]
  8. Miller, T. A. (1983) Am. J. Physiol. 245, G601-G623[Medline] [Order article via Infotrieve]
  9. Ligumsky, M., Golanska, E. M., Hansen, D. G., and Kauffman, G. J. (1983) Gastroenterology 84, 756-761[Medline] [Order article via Infotrieve]
  10. Ligumsky, M., Sestieri, M., Karmeli, F., Zimmerman, J., Okon, E., and Rachmilewitz, D. (1990) Gastroenterology 5, 1245-1249
  11. Tomisato, W., Tsutsumi, S., Rokutan, K., Tsuchiya, T., and Mizushima, T. (2001) Am. J. Physiol. 281, G1092-G1100
  12. Tomisato, W., Tsutsumi, S., Hoshino, T., Hwang, H. J., Mio, M., Tsuchiya, T., and Mizushima, T. (2004) Biochem. Pharmacol. 67, 575-585[CrossRef][Medline] [Order article via Infotrieve]
  13. Tomisato, W., Tanaka, K., Katsu, T., Kakuta, H., Sasaki, K., Tsutsumi, S., Hoshino, T., Aburaya, M., Li, D., Tsuchiya, T., Suzuki, K., Yokomizo, K., and Mizushima, T. (2004) Biochem. Biophys. Res. Commun. 323, 1032-1039[CrossRef][Medline] [Order article via Infotrieve]
  14. Kaufman, R. J. (1999) Genes Dev. 13, 1211-1233[Free Full Text]
  15. Mori, K. (2000) Cell 101, 451-454[CrossRef][Medline] [Order article via Infotrieve]
  16. Liu, H., Bowes, R. C., III, van de Water, B., Sillence, C., Nagelkerke, J. F., and Stevens, J. L. (1997) J. Biol. Chem. 272, 21751-21759[Abstract/Free Full Text]
  17. Reddy, R. K., Lu, J., and Lee, A. S. (1999) J. Biol. Chem. 274, 28476-28483[Abstract/Free Full Text]
  18. Zinszner, H., Kuroda, M., Wang, X., Batchvarova, N., Lightfoot, R. T., Remotti, H., Stevens, J. L., and Ron, D. (1998) Genes Dev. 12, 982-995[Abstract/Free Full Text]
  19. Tsutsumi, S., Gotoh, T., Tomisato, W., Mima, S., Hoshino, T., Hwang, H. J., Takenaka, H., Tsuchiya, T., Mori, M., and Mizushima, T. (2004) Cell Death Differ. 11, 1009-1016[CrossRef][Medline] [Order article via Infotrieve]
  20. Ding, H., Han, C., Zhu, J., Chen, C. S., and D'Ambrosio, S. M. (2005) Int. J. Cancer 113, 803-810[CrossRef][Medline] [Order article via Infotrieve]
  21. Zhang, Z., Lai, G. H., and Sirica, A. E. (2004) Hepatology 39, 1028-1037[CrossRef][Medline] [Order article via Infotrieve]
  22. Hsu, A. L., Ching, T. T., Wang, D. S., Song, X., Rangnekar, V. M., and Chen, C. S. (2000) J. Biol. Chem. 275, 11397-11403[Abstract/Free Full Text]
  23. Jiang, S., Chow, S. C., Nicotera, P., and Orrenius, S. (1994) Exp. Cell Res. 212, 84-92[CrossRef][Medline] [Order article via Infotrieve]
  24. Gwag, B. J., Canzoniero, L. M., Sensi, S. L., Demaro, J. A., Koh, J. Y., Goldberg, M. P., Jacquin, M., and Choi, D. W. (1999) Neuroscience 90, 1339-1348[CrossRef][Medline] [Order article via Infotrieve]
  25. Gil-Parrado, S., Fernandez-Montalvan, A., Assfalg-Machleidt, I., Popp, O., Bestvater, F., Holloschi, A., Knoch, T. A., Auerswald, E. A., Welsh, K., Reed, J. C., Fritz, H., Fuentes-Prior, P., Spiess, E., Salvesen, G. S., and Machleidt, W. (2002) J. Biol. Chem. 277, 27217-27226[Abstract/Free Full Text]
  26. Orrenius, S., Zhivotovsky, B., and Nicotera, P. (2003) Nat. Rev. Mol. Cell. Biol. 4, 552-565[CrossRef][Medline] [Order article via Infotrieve]
  27. Rizzuto, R., Pinton, P., Ferrari, D., Chami, M., Szabadkai, G., Magalhaes, P. J., Di Virgilio, F., and Pozzan, T. (2003) Oncogene 22, 8619-8627[CrossRef][Medline] [Order article via Infotrieve]
  28. Szabadkai, G., and Rizzuto, R. (2004) FEBS Lett. 567, 111-115[CrossRef][Medline] [Order article via Infotrieve]
  29. Hajnoczky, G., Davies, E., and Madesh, M. (2003) Biochem. Biophys. Res. Commun. 304, 445-454[CrossRef][Medline] [Order article via Infotrieve]
  30. Mandic, A., Viktorsson, K., Strandberg, L., Heiden, T., Hansson, J., Linder, S., and Shoshan, M. C. (2002) Mol. Cell. Biol. 22, 3003-3013[Abstract/Free Full Text]
  31. Wood, D. E., Thomas, A., Devi, L. A., Berman, Y., Beavis, R. C., Reed, J. C., and Newcomb, E. W. (1998) Oncogene 17, 1069-1078[CrossRef][Medline] [Order article via Infotrieve]
  32. Nakagawa, T., and Yuan, J. (2000) J. Cell Biol. 150, 887-894[Abstract/Free Full Text]
  33. Nozaki, S., Sledge Jr., G. W., and Nakshatri, H. (2001) Oncogene 20, 2178-2185[CrossRef][Medline] [Order article via Infotrieve]
  34. Johnson, A. J., Hsu, A. L., Lin, H. P., Song, X., and Chen, C. S. (2002) Biochem. J. 366, 831-837[Medline] [Order article via Infotrieve]
  35. Tomisato, W., Hoshino, T., Tsutsumi, S., Tsuchiya, T., and Mizushima, T. (2002) Dig. Dis. Sci. 47, 2125-2133[CrossRef][Medline] [Order article via Infotrieve]
  36. Tsutsumi, S., Tomisato, W., Takano, T., Rokutan, K., Tsuchiya, T., and Mizushima, T. (2002) Biochim. Biophys. Acta 1589, 168-180[Medline] [Order article via Infotrieve]
  37. Kao, J. P., Harootunian, A. T., and Tsien, R. Y. (1989) J. Biol. Chem. 264, 8179-8184[Abstract/Free Full Text]
  38. Katsu, T., Kobayashi, H., Hirota, T., Fujita, Y., Sato, K., and Nagai, U. (1987) Biochim. Biophys. Acta 899, 159-170[Medline] [Order article via Infotrieve]
  39. Vane, J. (1994) Nature 367, 215-216[CrossRef][Medline] [Order article via Infotrieve]
  40. Wang, J. L., Lin, K. L., Chen, J. S., Lu, Y. C., Jiann, B. P., Chang, H. T., Hsu, S. S., Chen, W. C., Huang, J. K., Ho, C. M., and Jan, C. R. (2004) Biochem. Pharmacol. 67, 1123-1130[CrossRef][Medline] [Order article via Infotrieve]
  41. Polster, B. M., Basanez, G., Etxebarria, A., Hardwick, J. M., and Nicholls, D. G. (2005) J. Biol. Chem. 280, 6447-6454[Abstract/Free Full Text]
  42. Liu, X., Yue, P., Zhou, Z., Khuri, F. R., and Sun, S. Y. (2004) J. Natl. Cancer Inst. 96, 1769-1780[Abstract/Free Full Text]
  43. Majumder, P. K., Pandey, P., Sun, X., Cheng, K., Datta, R., Saxena, S., Kharbanda, S., and Kufe, D. (2000) J. Biol. Chem. 275, 21793-21796[Abstract/Free Full Text]
  44. Wang, H. G., Pathan, N., Ethell, I. M., Krajewski, S., Yamaguchi, Y., Shibasaki, F., McKeon, F., Bobo, T., Franke, T. F., and Reed, J. C. (1999) Science 284, 339-343[Abstract/Free Full Text]
  45. Mukherjee, D. (2002) Biochem. Pharmacol. 63, 817-821[CrossRef][Medline] [Order article via Infotrieve]
  46. McAdam, B. F., Catella, L. F., Mardini, I. A., Kapoor, S., Lawson, J. A., and FitzGerald, G. A. (1999) Proc. Natl. Acad. Sci. U. S. A. 96, 272-277[Abstract/Free Full Text]
  47. Belton, O., Byrne, D., Kearney, D., Leahy, A., and Fitzgerald, D. J. (2000) Circulation 102, 840-845[Abstract/Free Full Text]
  48. Singh, D. (2004) Br. Med. J. 329, 816[Free Full Text]
  49. Ray, W. A., Griffin, M. R., and Stein, C. M. (2004) N. Engl. J. Med. 351, 2767[Free Full Text]
  50. Giovannucci, E., Rimm, E. B., Stampfer, M. J., Colditz, G. A., Ascherio, A., and Willett, W. C. (1994) Ann. Intern. Med. 121, 241-246[Abstract/Free Full Text]
  51. Baron, J. A. (2003) Prog. Exp. Tumor Res. 37, 1-24[Medline] [Order article via Infotrieve]
  52. Chan, T. A. (2002) Lancet Oncol. 3, 166-174[CrossRef][Medline] [Order article via Infotrieve]
  53. Hoshino, T., Tsutsumi, S., Tomisato, W., Hwang, H. J., Tsuchiya, T., and Mizushima, T. (2003) J. Biol. Chem. 278, 12752-12758[Abstract/Free Full Text]
  54. Eberhart, C. E., Coffey, R. J., Radhika, A., Giardiello, F. M., Ferrenbach, S., and DuBois, R. N. (1994) Gastroenterology 107, 1183-1188[Medline] [Order article via Infotrieve]
  55. Ristimaki, A., Honkanen, N., Jankala, H., Sipponen, P., and Harkonen, M. (1997) Cancer Res. 57, 1276-1280[Abstract/Free Full Text]
  56. Piazza, G. A., Alberts, D. S., Hixson, L. J., Paranka, N. S., Li, H., Finn, T., Bogert, C., Guillen, J. M., Brendel, K., Gross, P. H., Sperl, G., Ritchie, J., Burt, R. W., Ellsworth, L., Ahnen, D. J., and Pamukcu, R. (1997) Cancer Res. 57, 2909-2915[Abstract/Free Full Text]
  57. Reddy, B. S., Kawamori, T., Lubet, R. A., Steele, V. E., Kelloff, G. J., and Rao, C. V. (1999) Cancer Res. 59, 3387-3391[Abstract/Free Full Text]
  58. Zhang, X., Morham, S. G., Langenbach, R., and Young, D. A. (1999) J. Exp. Med. 190, 451-459[Abstract/Free Full Text]
  59. Hanif, R., Pittas, A., Feng, Y., Koutsos, M. I., Qiao, L., Staiano-Coico, L., Shiff, S. I., and Rigas, B. (1996) Biochem. Pharmacol. 52, 237-245[CrossRef][Medline] [Order article via Infotrieve]

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Home page
 J. Biol. Chem.Home page
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Up-regulation of S100P Expression by Non-steroidal Anti-inflammatory Drugs and Its Role in Anti-tumorigenic Effects
J. Biol. Chem., February 13, 2009; 284(7): 4158 - 4167.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
P. Maity, S. Bindu, S. Dey, M. Goyal, A. Alam, C. Pal, K. Mitra, and U. Bandyopadhyay
Indomethacin, a Non-steroidal Anti-inflammatory Drug, Develops Gastropathy by Inducing Reactive Oxygen Species-mediated Mitochondrial Pathology and Associated Apoptosis in Gastric Mucosa: A NOVEL ROLE OF MITOCHONDRIAL ACONITASE OXIDATION
J. Biol. Chem., January 30, 2009; 284(5): 3058 - 3068.
[Abstract] [Full Text] [PDF]

Home page
 J. Immunol.Home page
Y. Zhang, S. Liu, J. Liu, T. Zhang, Q. Shen, Y. Yu, and X. Cao
Immune Complex/Ig Negatively Regulate TLR4-Triggered Inflammatory Response in Macrophages through Fc{gamma}RIIb-Dependent PGE2 Production
J. Immunol., January 1, 2009; 182(1): 554 - 562.
[Abstract] [Full Text] [PDF]

Home page
 CarcinogenesisHome page
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NSAIDs suppress the expression of claudin-2 to promote invasion activity of cancer cells
Carcinogenesis, October 1, 2008; 29(10): 1994 - 2000.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
P. Maity, S. Bindu, V. Choubey, A. Alam, K. Mitra, M. Goyal, S. Dey, M. Guha, C. Pal, and U. Bandyopadhyay
Lansoprazole Protects and Heals Gastric Mucosa from Non-steroidal Anti-inflammatory Drug (NSAID)-induced Gastropathy by Inhibiting Mitochondrial as Well as Fas-mediated Death Pathways with Concurrent Induction of Mucosal Cell Renewal
J. Biol. Chem., May 23, 2008; 283(21): 14391 - 14401.
[Abstract] [Full Text] [PDF]

Home page
 Cancer Res.Home page
A. Kardosh, E. B. Golden, P. Pyrko, J. Uddin, F. M. Hofman, T. C. Chen, S. G. Louie, N. A. Petasis, and A. H. Schonthal
Aggravated Endoplasmic Reticulum Stress as a Basis for Enhanced Glioblastoma Cell Killing by Bortezomib in Combination with Celecoxib or Its Non-Coxib Analogue, 2,5-Dimethyl-Celecoxib
Cancer Res., February 1, 2008; 68(3): 843 - 851.
[Abstract] [Full Text] [PDF]

Home page
 Molecular Cancer TherapeuticsHome page
P. Pyrko, A. Kardosh, Y.-T. Liu, N. Soriano, W. Xiong, R. H. Chow, J. Uddin, N. A. Petasis, A. K. Mircheff, R. A. Farley, et al.
Calcium-activated endoplasmic reticulum stress as a major component of tumor cell death induced by 2,5-dimethyl-celecoxib, a non-coxib analogue of celecoxib
Mol. Cancer Ther., April 1, 2007; 6(4): 1262 - 1275.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Pharmacol.Home page
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Up-Regulation of 150-kDa Oxygen-Regulated Protein by Celecoxib in Human Gastric Carcinoma Cells
Mol. Pharmacol., March 1, 2007; 71(3): 860 - 870.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
M. Aburaya, K.-I. Tanaka, T. Hoshino, S. Tsutsumi, K. Suzuki, M. Makise, R. Akagi, and T. Mizushima
Heme Oxygenase-1 Protects Gastric Mucosal Cells against Non-steroidal Anti-inflammatory Drugs
J. Biol. Chem., November 3, 2006; 281(44): 33422 - 33432.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Pharmacol.Home page
I. Alloza, A. Baxter, Q. Chen, R. Matthiesen, and K. Vandenbroeck
Celecoxib Inhibits Interleukin-12 {alpha}beta and beta2 Folding and Secretion by a Novel COX2-Independent Mechanism Involving Chaperones of the Endoplasmic Reticulum
Mol. Pharmacol., May 1, 2006; 69(5): 1579 - 1587.
[Abstract] [Full Text] [PDF]

Home page
 Mol. Pharmacol.Home page
H. Ushijima, K.-I. Tanaka, M. Takeda, T. Katsu, S. Mima, and T. Mizushima
Geranylgeranylacetone Protects Membranes against Nonsteroidal Anti-Inflammatory Drugs
Mol. Pharmacol., October 1, 2005; 68(4): 1156 - 1161.
[Abstract] [Full Text] [PDF]

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