Affiliations of authors: X. Song, A. J. Johnson, Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky, Lexington; H.-P. Lin, P.-H. Tseng, Y.-T. Yang, S. K. Kulp, Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, and The Comprehensive Cancer Center, The Ohio State University, Columbus; C.-S. Chen, Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky and Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, and The Comprehensive Cancer Center, The Ohio State University.
Correspondence to: Ching-Shih Chen, Ph.D., Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, The Ohio State University, 500 West 12th Ave., Columbus, OH 432101291 (e-mail: chen.844{at}osu.edu).
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Previously, we demonstrated that celecoxib (Celebrex®) induced apoptosis in prostate cancer cells by interfering with multiple signaling targets, including Akt, ERK2, and endoplasmic reticulum Ca2+-ATPases (10,25). Disruption of these signaling pathways leads rapidly to apoptosis, an apoptotic mechanism distinctly different from that of conventional anticancer agents. It is noteworthy that the effect of celecoxib on apoptosis was independent of androgen responsiveness, the level of Bcl-2 expression, and the functional status of p53 in cancer cells (10,25). Nevertheless, this rapid induction of apoptosis was unique to celecoxib, because the abilities of other COX-2 inhibitors, including rofecoxib (Vioxx®), NS398, and DuP697, to induce apoptosis were much lower than that of celecoxib (25). This observation underscores differences in the mechanisms by which these COX-2 inhibitors mediate apoptosis in prostate cancer cells. To determine whether COX-2 inhibitor-induced apoptosis required the inhibition of COX-2 enzyme activity, we examined the effect of COX-2 depletion on apoptosis in PC-3 clones carrying tetracycline-on (Tet-On) antisense COX-2 complementary DNAs (cDNAs) and performed a structureactivity analysis of various celecoxib derivatives in androgen-independent PC-3 cells.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We used the following three different prostate cancer cell lines to assess the impact of androgen responsiveness and p53 functional status on the induction of apoptosis by celecoxib and its derivatives: androgen-responsive LNCaP (p53+/+), androgen-nonresponsive PC-3 (p53/), and DU-145 (p53/). The antisense COX-2 construct was a gift from Drs. Rebecca Chinery and Jason Morrow (Vanderbilt University Medical School, Nashville, TN). It contained an almost complete human COX-2 cDNA insert (1.93 kilobases) that was cloned into the XbaI/EcoRV sites in the tetracycline response element (TRE)-response plasmid pUHD.2neo (26). This Tet-On antisense COX-2 construct has been used in colorectal cancer cells to assess the role of prostaglandins in cell proliferation (26). Celecoxib and rofecoxib were from commercial Celebrex® and Vioxx® (Amerisource Health, Malvern, PA) capsules, respectively, by solvent extraction followed by recrystallization. DuP697 was a gift from Professor Hsin-Hsiung Tai (University of Kentucky, Lexington), and NS398 was from Calbiochem (La Jolla, CA). The following compounds were synthesized according to published procedures (27): 4-[5-(4-chlorophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 1), 4-[5-phenyl-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 2), 4-[5-(4-aminophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 3), 4-[5-(4-ethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 4), 4-[5-(4-trifluoromethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 5), 4-[5-(2,5-dichlorophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 6), and 4-[5-(2,5-dimethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (compound 7). Rabbit anti-COX-2 antibodies were from Cayman Chemicals (Ann Arbor, MI). Rabbit polyclonal antibodies against Akt, phospho-Ser473-Akt, ERK, and phospho-ERK were from New England Biolabs (Beverly, MA), and mouse anti-actin monoclonal antibody was from ICN Pharmaceuticals (Costa Mesa, CA). Goat anti-rabbit immunoglobulin G (IgG)-horseradish peroxidase conjugates were from Jackson ImmunoResearch Laboratories (West Grove, PA). Rabbit anti-poly(ADP-ribose) polymerase (PARP) antibodies were from BD PharMingen (San Diego, CA).
Development of PC-3 Tet-On Antisense COX-2 Clones
PC-3 cells were cultured in RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS) in T-25 flasks at 37 °C in a humidified CO2 incubator to 80% confluence. Each flask was washed with 6 mL of serum-free Opti-MEM (Invitrogen Life Technologies, Carlsbad, CA), and then 3 mL of serum-free Opti-MEM was added. Aliquots containing 0.12 µg of the Tet-On regulator plasmid pTet-On (Clontech, Palo Alto, CA) and 0.12 µg of the antisense COX-2 construct in 150 µL of serum-free Opti-MEM medium were preincubated with 3 µL of the Plus reagent from the LipofectAMINE Plus Reagent kit (Invitrogen Life Technologies) at 25 °C for 15 minutes, followed by 12 µL of the LipofectAMINE reagent in 150 µL of Opti-MEM medium. The resulting mixture was incubated at 25 °C for 15 minutes and then added to each flask with gentle mixing. After 5 hours at 37 °C, the transfection medium was replaced with 5 mL of RPMI-1640 medium containing 10% Tet-System-approved FBS (Clontech). After 48 hours, cells were cultured in fresh medium containing G418 (Calbiochem, La Jolla, CA) at 100 µg/mL to select for transfected clones. The G418-supplemented medium was changed every 4 days. After 3 weeks, G418-resistant cells were subcloned into 96-well plates by limiting dilution with a final cell density of approximately 0.5 cell per well. After 12 days with a change of G418-containing medium every 4 days, viable clones were further subcloned into 12-well plates. After 4 or 5 days, cells in each well were divided into three T-25 flasks. The level of COX-2 expression was determined 120 hours after cells were exposed to doxycycline (2 µg/mL) by western blot analysis. By this procedure, the following four independent clones (2F6, 1F2, 3D9, and 7D9) were selected for analyses; 2F6 was a COX-2-deficient clone, and 1F2, 3D9, and 7D9 expressed different levels of COX-2 in the absence of doxycycline.
Immunoblotting
For western blot analysis, cells were washed in phosphate-buffered saline (PBS), resuspended in sodium dodecyl sulfate (SDS) gel-loading buffer (50 mM TrisHCl [pH 6.8], 100 mM dithiothreitol, 2% SDS, 0.1% bromophenol blue, and 10% glycerol), sonicated with an ultrasonic sonicator for 5 seconds (Virsonic 300, 4.5 output; VirTis, Gardiner, NY), and boiled for 5 minutes. After a brief centrifugation, equivalent protein amounts (60100 µg) from the soluble fractions were resolved in 10% SDSpolyacrylamide gels on a Minigel (Amersham Pharmacia, Piscataway, NJ) apparatus and transferred to a nitrocellulose membrane in a semidry transfer cell. The transblotted membrane was washed twice with Tris-buffered saline (TBS) containing 0.05% Tween 20 (TBST). After blocking with TBS containing 5% nonfat milk for 60 minutes, the membrane was incubated with the appropriate primary antibody (anti-COX-2, anti-Akt, anti-P-473Ser Akt, anti-ERK, and anti-phospho-ERK antibodies diluted 1 : 1000; anti-actin monoclonal antibody, diluted 1 : 5000) in TBS1% nonfat milk at 4 °C for 12 hours and washed twice with TBST. The membranes were probed with goat anti-rabbit IgG-horseradish peroxidase conjugates (diluted 1 : 5000) for 1 hour at room temperature and washed twice with TBST. Bands were visualized by enhanced chemiluminescence.
Prostaglandin E2 (PGE2) Immunoassay
Parental and transfected cells, with or without a doxycycline (2 µg/mL) pretreatment, were grown to 10 x 106 cells in T-75 flasks in RPMI-1640 medium containing 10% FBS. Culture medium was changed, and 24 hours later, conditioned medium was collected to assay PGE2. Conditioned medium was centrifuged to remove particulate material, and then attached cells were collected by scraping to determine the protein concentration. PGE2 was assayed in 100 µL of medium in triplicate, according to the manufacturer's instruction (R&D Systems, Minneapolis, MN). PGE2 data were normalized to protein content.
Cell Viability
Parental or transfected PC-3 cells with or without doxycycline (2 µg/mL) pretreatment were plated in 12-well plates and cultured in RPMI-1640 medium supplemented with 10% FBS in the absence or presence of doxycycline (2 µg/mL) for 48 hours. COX-2 inhibitors (at various concentrations) were dissolved in dimethyl sulfoxide (DMSO; final concentration = 0.1% after addition to medium) and were then added to the cells in serum-free RPMI-1640 medium. Control cells received DMSO at the same concentration. During treatment, the percentage of floating cells increased over time. At the end of the treatment, adherent cells were harvested by trypsinization, and floating cells were recovered by centrifugation at 3200g for 5 minutes. Cell morphology was assessed with a light microscope at x400. Both adherent and floating cells were combined, and cell viability was assessed by trypan blue dye exclusion.
Analysis of Apoptosis
In addition to the morphologic changes of intact cells observed by phase-contrast microscopy, four methods were used to assess drug-induced apoptotic cell death.
Phosphatidylserine externalization. Approximately 2.5 x 105 cells were grown on glass coverslips for 24 hours. At various times after drug treatment, cells were washed gently with PBS and then exposed to 0.5 mL of binding buffer (10 mM HEPES [pH 7.4], containing 150 mM NaCl, 2.5 mM CaCl2, 1 mM MgCl2, and 4% bovine serum albumin), followed by 0.6 mL of annexin V-fluorescein isothiocynate (FITC) (200 µg/mL) for 30 minutes. After washing with binding buffer, apoptotic cells were identified directly as cells with annexin V-FITC on their outer membrane by fluorescence microscopy. In a set of controls, cells received medium containing DMSO vehicle in lieu of the test agent.
4`,6-Diamidino-2-phenylindole (DAPI) staining of nuclei.
At various times after treatment with various test agents, morphologic changes were detected in the nuclei of apoptotic cells by staining with the DNA-binding fluorochrome DAPI. For adherent PC-3 cells, cells were grown on glass coverslips until approximately 70% confluent and exposed to the test agent at 50 µM for various times. Supernatants then were carefully removed, adherent cells were washed with PBS, DAPI (0.5 µg/mL) was added in a fixation solution (4% paraformaldehyde, 2 mM EGTA [ethylene glycol bis(-aminoethyl ether)-N,N,N`,N`-tetraacetic acid], and 13.7% sucrose in PBS), and the mixture was incubated at room temperature for 20 minutes in the dark. Cells were then washed for two 20-minute periods with PBS. Floating PC-3 cells were examined by a modification of the above method as follows. PC-3 cells were cultured in T-25 flasks and treated with the test agent. Floating cells then were collected, washed, and stained with DAPI as described above. Cells were allowed to attach to poly-L-lysine-coated coverslips and viewed by microscopy at a magnification of x400.
Apoptosis detection by an enzyme-linked immunosorbent assay (ELISA). Induction of apoptosis was also assessed by using a Cell Death Detection ELISA (Roche Diagnostics, Mannheim, Germany) by following the manufacturer's instructions. This test is based on the quantitative determination of cytoplasmic histone-associated DNA fragments in the form of mononucleosomes and oligonucleosomes after induced apoptotic death. In brief, 2.5 x 106 PC-3 cells were cultured in a T-75 flask 24 hours before the experiment. Cells were washed twice in 5 mL of serum-free RPMI-1640 medium and then treated with a test agent or the DMSO vehicle, as indicated. Both floating and adherent cells were collected, and cell lysates equivalent to 104 cells were used in the ELISA.
Western blot analysis of PARP cleavage. Drug-treated cells were collected, washed with ice-cold PBS, and resuspended in lysis buffer [20 mM TrisHCl (pH 8), 137 mM NaCl, 1 mM CaCl2, 10% glycerol, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS, 100 µM 4-(2-aminoethyl)benzenesulfonyl fluoride, leupeptin at 10 µg/mL, and aprotinin at 10 µg/mL]. Soluble cell lysates were collected after centrifugation at 1500g for 5 minutes. Equivalent amounts of protein (60100 µg) from each lysate were resolved in 10% SDSpolyacrylamide gels. Bands were transferred to nitrocellulose membranes and analyzed by immunoblotting with anti-PARP antibodies, as described above.
Statistical Analysis
Each experiment was performed in triplicate. All experiments were carried out at least two times on different occasions. Where appropriate, the data are presented as the mean ± 95% confidence interval.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
To assess the effect of COX-2 enzyme activity on cell growth, we prepared Tet-On antisense COX-2 clones by transfecting parental PC-3 cells with an antisense COX-2 cDNA construct under the control of a tetracycline-inducible promoter. Four Tet-On antisense COX-2 clones, 2F6, 1F2, 3D9, and 7D9, were selected after subcloning the G418-resistant cells by limiting dilution. 2F6 is a COX-2-deficient clone in which the COX-2 protein was virtually undetectable, and 1F2, 3D9, and 7D9 are antisense COX-2 clones that express COX-2 protein at different levels. In 1F2, 3D9, and 7D9 in the absence of doxycycline, COX-2 is expressed, but in the presence of doxycycline, COX-2 is depleted by antisense inhibition (Fig. 1, B). Fig. 1, A
, shows a western blot analysis of the effect of doxycycline (2 µg/mL) on COX-2 expression in the Tet-On clone 3D9 over a 10-day period. By 4 days after the addition of doxycycline, COX-2 protein had been depleted in 3D9 cells, and as long as doxycycline was present, COX-2 was not detected. Because COX-1 expression was negligible in these clones (data not shown), COX-2 produced most of the prostaglandins. Accordingly, the level of COX-2 expression reflected the level of PGE2 production. Although the levels of PGE2 in 1F2 cells and parental PC-3 cells were comparable, those in 3D9 were twofold higher and those in 7D9 cells were 10-fold higher than levels in parental cells (Fig. 1, C
). However, when treated with rofecoxib, NS398, or DuP697 at 50 µM, PGE2 production was reduced to the same extent in 1F2, 3D9, 7D9, and parental PC-3 cells (data not shown; it could not be determined in celecoxib-treated cells whether PGE2 was produced by rapid apoptosis). Because the decreased expression of the antiapoptotic protein Bcl-2 has been implicated in the apoptotic mechanism of the COX-2 inhibitors SC-58125 and NS398 (1,2), it is also noteworthy that COX-2 ablation had essentially no impact on the expression of Bcl-2 (Fig. 1, D
).
|
Using these COX-2 antisense clones, we obtained two lines of evidence that the effect of COX-2 inhibitors on apoptosis was independent of their COX-2-inhibitory activity. First, although both antisense COX-2 cDNA and COX-2 inhibitors completely blocked prostaglandin production, their effects on cell viability were markedly different. Treatment of PC-3 cells or any of the four clones with individual COX-2 inhibitors led to apoptotic death, whereas depletion of COX-2, and thus inhibition of PGE2 production with the antisense cDNA, did not adversely affect the viability of these antisense clones, i.e., this treatment did not induce cell death. Second, although the basal levels of COX-2 in these four clones varied, all clones and parental PC-3 cells were equally susceptible to apoptosis induced by COX-2 inhibitors, and this susceptibility did not change after doxycycline-induced COX-2 depletion. In other words, susceptibility to COX-2-inhibitor-induced apoptosis was independent of the level of COX-2 expression. Fig. 2, A, shows the time course of cell death in the presence of 50 µM celecoxib in PC-3 cells, COX-2-deficient 2F6 cells, and COX-2-overexpressing 7D9 cells with and without COX-2 depletion, i.e., grown in the presence and absence of doxycycline, respectively. For all three of these clones incubated with 50 µM celecoxib, the time required for 50% cell death (T1/2) was approximately 2 hours. Similar times were noted with 1F2 and 3D9 cells.
Previously, we demonstrated that celecoxib induced rapid apoptotic death in both androgen-responsive LNCaP (p53+/+) and androgen-nonresponsive PC-3 (p53/) prostate cancer cells by inhibiting the Akt and ERK signaling pathways (10,25). In this study, the apoptotic death induced in all four clones was also associated with decreased phosphorylation of Akt and ERK2, as observed in parental PC-3 cells. In addition, the time course for the dephosphorylation of Akt and ERK2 in 7D9 cells (Fig. 2, B) was consistent with that for cell death. Similar results were obtained with the three other clones.
|
|
Structural modifications of celecoxib were carried out to dissociate COX-2 inhibition and the induction of apoptosis. We synthesized a series of celecoxib derivatives with different substituents at the terminal phenyl ring and examined the apoptosis-inducing potency of each. Fig. 4 summarizes the structures, the COX-2 inhibitory activity (IC50 = concentration of drug-inhibiting COX-2 activity by 50%), and the apoptosis-inducing activity (T1/2) of celecoxib and seven representative analogues.
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Use of the Tet-On antisense COX-2 clones was advantageous because these clones represented syngeneic cell lines, thereby abating concerns about the effects of genetic variations among different cell lines, and they displayed differential COX-2 expression that could be shut off by doxycycline treatment. We determined that the effect of COX-2 inhibitors on apoptosis was independent of the COX-2 inhibitory activity. Although both COX-2 depletion and COX-2 inhibitors inhibited PGE2 production, these two treatments had very different effects on cell viability. Treatment with COX-2 inhibitors led to cell death, but COX-2 depletion did not. The sensitivity to COX-2-inhibitor-induced apoptosis was independent of the level of COX-2 expression in the antisense clones and was, in fact, unaltered from that of parental PC-3 cells.
We were able to dissociate the apoptosis-inducing activity of celecoxib from the COX-2 inhibitory activity via structural modification of this drug. This finding is reminiscent of the previous report that sulindac metabolitessulindac sulfide and sulindac sulfonecould induce apoptosis in prostate cancer cells via a COX-independent mechanism (22). Several celecoxib derivatives, although lacking COX-2 inhibitory activity, were as potent in eliciting apoptosis in PC-3 cells as the parent compound. These compounds induce apoptosis in both hormone-responsive and hormone-nonresponsive prostate cancer cells. Furthermore, the mechanism by which these compounds and the parental compound celecoxib induce apoptosis remained the same, i.e., facilitating the dephosphorylation of Akt and ERK2.
From a clinical perspective, the separation of the COX-2 inhibitory activity of celecoxib from the apoptosis-inducing effect provides a molecular basis for the design of new classes of apoptosis-inducing agents. It is noteworthy that these molecules mediate apoptosis independent of the cell's androgen sensitivity, p53 functional status, and level of Bcl-2 expression. These features make these apoptosis-inducing agents potential candidates for the prevention and treatment of human prostate cancer.
![]() |
NOTES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
X. Song and H.-P. Lin contributed equally to this paper.
We thank Dr. Rebecca Chinery and Dr. Jason Morrow at Vanderbilt University Medical School for providing the antisense COX-2 cDNA construct and Dr. Hsin-Hsiung Tai at the University of Kentucky for providing DuP697.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1 Sheng H, Shao J, Morrow JD, Beauchamp RD, DuBois RN. Modulation of apoptosis and Bcl-2 expression by prostaglandin E2 in human colon cancer cells. Cancer Res 1998;58:3626.[Abstract]
2 Liu XH, Yao S, Kirschenbaum A, Levine AC. NS398, a selective cyclooxygenase-2 inhibitor, induces apoptosis and down-regulates bcl-2 expression in LNCaP cells. Cancer Res 1998;58:42459.[Abstract]
3
Chan TA, Morin PJ, Vogelstein B, Kinzler KW. Mechanisms underlying nonsteroidal antiinflammatory drug-mediated apoptosis. Proc Natl Acad Sci U S A 1998;95:6816.
4 Piazza GA, Rahm AL, Krutzsch M, Sperl G, Paranka NS, Gross PH, et al. Antineoplastic drugs sulindac sulfide and sulfone inhibit cell growth by inducing apoptosis. Cancer Res 1995;55:31106.[Abstract]
5 Hanif R, Pittas A, Feng Y, Koutsos MI, Qiao L, Staiano-Coico L, et al. Effects of nonsteroidal anti-inflammatory drugs on proliferation and on induction of apoptosis in colon cancer cells by a prostaglandin-independent pathway. Biochem Pharmacol 1996;52:23745.[Medline]
6 Thompson HJ, Jiang C, Lu J, Mehta RG, Piazza GA, Paranka NS, et al. Sulfone metabolite of sulindac inhibits mammary carcinogenesis. Cancer Res 1997;57:26771.[Abstract]
7 Jones MK, Wang H, Peskar BM, Levin E, Itani RM, Sarfeh IJ, et al. Inhibition of angiogenesis by nonsteroidal anti-inflammatory drugs: insight into mechanisms and implications for cancer growth and ulcer healing. Nat Med 1999;5:141823.[Medline]
8
Williams CS, Tsujii M, Reese J, Dey SK, DuBois RN. Host cyclooxygenase-2 modulates carcinoma growth. J Clin Invest 2000;105:158994.
9
Zhang X, Morham SG, Langenbach R, Young DA. Malignant transformation and antineoplastic actions of nonsteroidal antiinflammatory drugs (NSAIDs) on cyclooxygenase-null embryo fibroblasts. J Exp Med 1999;190:4519.
10
Hsu AL, Ching TT, Wang DS, Song X, Rangnekar VM, Chen CS. The cyclooxygenase-2 inhibitor celecoxib induces apoptosis by blocking Akt activation in human prostate cancer cells independently of Bcl-2. J Biol Chem 2000;275:11397403.
11
Marx J. Cancer research. Anti-inflammatories inhibit cancer growthbut how? Science 2001;291:5812.
12 Hla T, Ristimaki A, Appleby S, Barriocanal JG. Cyclooxygenase gene expression in inflammation and angiogenesis. Ann N Y Acad Sci 1993;696: 197204.[Medline]
13
Dubois RN, Abramson SB, Crofford L, Gupta RA, Simon LS, Van De Putte LB, et al. Cyclooxygenase in biology and disease. FASEB J 1998;12:106373.
14
Taketo MM. Cyclooxygenase-2 inhibitors in tumorigenesis (part I). J Natl Cancer Inst 1998;90:152936.
15 Hla T, Bishop-Bailey D, Liu CH, Schaefers HJ, Trifan OC. Cyclooxygenase-1 and -2 isoenzymes. Int J Biochem Cell Biol 1999;31:5517.[Medline]
16 Prescott SM, Fitzpatrick FA. Cyclooxygenase-2 and carcinogenesis. Biochim Biophys Acta 2000;1470:M6978.[Medline]
17 Tsujii M, DuBois RN. Alterations in cellular adhesion and apoptosis in epithelial cells overexpressing prostaglandin endoperoxide synthase 2. Cell 1995;83:493501.[Medline]
18 DuBois RN, Shao J, Tsujii M, Sheng H, Beauchamp RD. G1 delay in cells overexpressing prostaglandin endoperoxide synthase-2. Cancer Res 1996;56:7337.[Abstract]
19
McGinty A, Chang YW, Sorokin A, Bokemeyer D, Dunn MJ. Cyclooxygenase-2 expression inhibits trophic withdrawal apoptosis in nerve growth factor-differentiated PC12 cells. J Biol Chem 2000;275:12095101.
20 Oshima M, Dinchuk JE, Kargman SL, Oshima H, Hancock B, Kwong E, et al. Suppression of intestinal polyposis in Apc delta716 knockout mice by inhibition of cyclooxygenase 2 (COX-2). Cell 1996;87:8039.[Medline]
21
Liu CH, Chang SH, Narko K, Trifan OC, Wu MT, Smith E, et al. Overexpression of cyclooxygenase-2 is sufficient to induce tumorigenesis in transgenic mice. J Biol Chem 2001;276:185639.
22 Lim JT, Piazza GA, Han EK, Delohery TM, Li H, Finn TS, et al. Sulindac derivatives inhibit growth and induce apoptosis in human prostate cancer cell lines. Biochem Pharmacol 1999;58:1097107.[Medline]
23
Narko K, Ristimaki A, MacPhee M, Smith E, Haudenschild CC, Hla T. Tumorigenic transformation of immortalized ECV endothelial cells by cyclooxygenase-1 overexpression. J Biol Chem 1997;272:2145560.
24
Trifan OC, Smith RM, Thompson BD, Hla T. Overexpression of cyclooxygenase-2 induces cell cycle arrest. Evidence for a prostaglandin-independent mechanism. J Biol Chem 1999;274:341417.
25 Johnson AJ, Song X, Hsu A, Chen C. Apoptosis signaling pathways mediated by cyclooxygenase-2 inhibitors in prostate cancer cells. Adv Enzyme Regul 2001;41:22135.[Medline]
26
Chinery R, Coffey RJ, Graves-Deal R, Kirkland SC, Sanchez SC, Zackert WE, et al. Prostaglandin J2 and 15-deoxy-delta12,14-prostaglandin J2 induce proliferation of cyclooxygenase-depleted colorectal cancer cells. Cancer Res 1999;59:273946.
27 Penning TD, Talley JJ, Bertenshaw SR, Carter JS, Collins PW, Docter S, et al. Synthesis and biological evaluation of the 1,5-diarylpyrazole class of cyclooxygenase-2 inhibitors: identification of 4-[5-(4-methylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenesulfonamide (SC-58635, celecoxib). J Med Chem 1997;40:134765.[Medline]
28
Cao Y, Pearman AT, Zimmerman GA, McIntyre TM, Prescott SM. Intracellular unesterified arachidonic acid signals apoptosis. Proc Natl Acad Sci U S A 2000;97:112805.
29 He TC, Chan TA, Vogelstein B, Kinzler KW. PPARdelta is an APC-regulated target of nonsteroidal anti-inflammatory drugs. Cell 1999;99:33545.[Medline]
30 Yin MJ, Yamamoto Y, Gaynor RB. The anti-inflammatory agents aspirin and salicylate inhibit the activity of I(kappa)B kinase-beta. Nature 1998;396:7780.[Medline]
Manuscript received September 27, 2001; revised February 13, 2002; accepted February 26, 2002.
This article has been cited by other articles in HighWire Press-hosted journals:
![]() |
||||
|
Oxford University Press Privacy Policy and Legal Statement |