Affiliations of authors: J. Zhu, H.-P. Lin (Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy), D. C. Young (Biostatistics Core, Comprehensive Cancer Center), The Ohio State University, Columbus; X. Song, Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky, Lexington; S. Yan, V. E. Marquez, Laboratory of Medicinal Chemistry, Center for Cancer Research, National Cancer Institute at Frederick, Frederick, MD; C.-S. Chen, Division of Medicinal Chemistry and Pharmacognosy, College of Pharmacy, The Ohio State University, and Division of Pharmaceutical Sciences, College of Pharmacy, University of Kentucky.
Correspondence to: Ching-Shih Chen, Ph.D., College of Pharmacy, The Ohio State University, 336 Parks Hall, 500 W. 12th Ave., Columbus, OH 432101291 (e-mail: chen.844{at}osu.edu).
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ABSTRACT |
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INTRODUCTION |
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However, an expanding body of evidence suggests that COX-2 inhibition may not play a role in NSAID-mediated apoptotic cell death (19). For example, sulindac sulfide and sulindac sulfone, which are metabolites of the NSAID sulindac, have been reported to mediate apoptosis in cancer cells via the inhibition of cyclic GMP phosphodiesterase (2023). which is a COX-2-independent mechanism (24). In addition, we have used a tetracycline-inducible antisense COX-2 expression plasmid to demonstrate that the sensitivity of prostate cancer cells to COX-2 inhibitor-induced apoptosis is independent of the expression status of COX-2 (25). The finding that these two pharmacologic effects of NSAIDsCOX-2 inhibition and apoptosis inductionare separable has considerable therapeutic implications and provides molecular underpinnings for the design of a new class of anticancer compounds whose mode of action is different from that of conventional chemotherapeutic agents.
Celecoxib induces apoptosis in prostate cancer cells by interfering with multiple signaling targets, including the serine/threonine kinase Akt, extracellular signal-regulated kinase 2 (ERK2), and endoplasmic reticulum Ca2+-ATPases (26,27). Disruption of these signaling pathways results in the loss of regulation of cellular functions that govern cell growth and survival, leading to rapid apoptotic death. This rapid induction of apoptosis, however, is unique to celecoxib because other COX-2 inhibitors that have the same COX-2 inhibitory potencies as celecoxib, including rofecoxib (Vioxx®), NS398, and DuP697, display apoptosis-inducing activities that are nearly two orders of magnitude lower than that displayed by celecoxib (26,27). Here we use celecoxib and rofecoxib as molecular starting points from which to understand the structural basis underlying this discrepancy and to optimize the apoptosis-inducing potency of celecoxib in prostate cancer cells.
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MATERIALS AND METHODS |
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Celecoxib and rofecoxib were extracted with ethyl acetate followed by recrystallization in a mixture consisting of ethyl acetate and hexane from Celebrex® and Vioxx®, respectively, which were obtained from Amerisource Health (Malvern, PA). NS398 was purchased from Calbiochem (San Diego, CA). Rabbit polyclonal antibodies against Akt, phospho-473Ser Akt, p44/42 ERKs, and phospho-p44/42 ERKs were purchased from Cell Signaling Technologies (Beverly, MA). A rabbit polyclonal anti-poly(ADP-ribose) polymerase (PARP) antibody was obtained from Pharmingen (San Diego, CA). Other chemical and biochemical reagents used were obtained from Sigma-Aldrich (St. Louis, MO), unless otherwise mentioned.
Synthesis of Compounds
In this article, we discuss 50 compounds. The full chemical name of each of these compounds is provided in Table 1. We used published procedures to synthesize compounds 1, 1-NH2, 229, and 4046. (2831). Proton nuclear magnetic resonance (1H NMR) spectroscopy, high-resolution mass spectrometry (HRMS), and elemental analysis were used to validate the identity of each of the synthetic compounds. The procedures we used to synthesize compounds 3039 and 4750 are presented below. Our results and the properties of these chemicals are summarized in Table 2
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4-[5-(2,4-Dichlorophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 31) was synthesized from 2',4'-dichloroacetophenone using the two-step procedure described for the synthesis of compound 30, in which 2',4'-dichloroacetophenone was used in place of 4'-chloroacetophenone (52% overall yield).
4-[5-(2,5-Dichlorophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 32) was synthesized from 2',5'-dichloroacetophenone using the two-step procedure described for the synthesis of compound 30, in which 2',5'-dichloroacetophenone was used in place of 4'-chloroacetophenone (60% overall yield).
4-[5-(3,4-Dichlorophenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 33) was synthesized from 3',4'-dichloroacetophenone using the two-step procedure described for the synthesis of compound 30, in which 3',4'-dichloroacetophenone was used in place of 4'-chloroacetophenone (55% overall yield).
4-[5-(4-Methylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl] benzenecarboxyamide (compound 34) was synthesized from 4'-methylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 4'-methylacetophenone was used in place of 4'-chloroacetophenone (65% overall yield).
4-[5-(4-Trifluoromethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 35) was synthesized from 4'-trifluoromethylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 4'-trifluoromethylacetophenone was used in place of 4'-chloroacetophenone (53% overall yield).
4-[5-(4-Ethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl] benzenecarboxyamide (compound 36) was synthesized from 4'-ethylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 4'-ethylacetophenone was used in place of 4'-chloroacetophenone (44% overall yield).
4-[5-(2,4-Dimethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 37) was synthesized from 2',4'-dimethylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 2',4'-dimethylacetophenone was used in place of 4'-chloroacetophenone (62% overall yield).
4-[5-(2,5-Dimethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 38) was synthesized from 2',5'-dimethylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 2',5'-dimethylacetophenone was used in place of 4'-chloroacetophenone (58% overall yield).
4-[5-(3,4-Dimethylphenyl)-3-(trifluoromethyl)-1H-pyrazol-1-yl]benzenecarboxyamide (compound 39) was synthesized from 3',4'-dimethylacetophenone using the two-step procedure described for the synthesis of compound 30, in which 3',4'-dimethylacetophenone was used in place of 4'-chloroacetophenone (56% overall yield).
3-(4-Methylsulfonylphenyl)-4-phenyl-2(5H)-furanone (compound 47) was synthesized in two steps. In the first step, a mixture of 4'-(methylsulfonyl)acetophenone (5.5 g, 27.8 mmol), morpholine (2.5 mL), and sulfur (0.89 g, 27.8 mmol) was refluxed for 10 hours and then poured into ice where it formed a precipitate. The precipitate was collected by filtration and washed with cold ethyl acetate. The precipitate was added to 10% sodium hydroxide (55 mL), and the mixture was then heated to 84 °C for 12 hours, forming an alkaline solution that was acidified to pH 3 with 12 N HCl. The solid that resulted from acidification was collected by filtration, dried, and recrystallized from a solution containing equal volumes of hexane and ethyl acetate to give 4-methylsulfonylphenylacetic acid (a white solid; 4.2 g, 52% overall yield). In the second step, 2-bromoacetophenone (1.02 g, 5.12 mmol) dissolved in acetonitrile (28 mL) was added to triethylamine (1.74 mL), followed by the addition of 4-methylsulfonylphenylacetic acid (1 g, 4.67 mmol). The mixture was stirred at room temperature for 1.5 hours and then 1,8-diazabicyclo[5,4,0]undec-7-ene (1.67 mL) was added. The mixture was stirred for another hour, after which 1 N HCl (35 mL) was added. The end product was extracted from the mixture with ethyl acetate, dried over sodium sulfate, and recrystallized from ethyl acetatehexane (vol/vol) to give compound 47 (880 mg, 60% overall yield).
3-(4-Sulfamoylphenyl)-4-phenyl-2(5H)-furanone (compound 48) was synthesized from 4-sulfamoylphenylacetic acid and 2-bromoacetophenone in a manner similar to that described for compound 47, with a 40% yield.
3-(4-Sulfamoylphenyl)-4-(2,4-dichlorophenyl)-2(5H)-furanone (compound 49) was synthesized from 4-sulfamoylphenylacetic acid and 2-bromo-1-(2'4-dichlorophenyl)acetophenone in a manner similar to that described for compound 47, with a 32% yield.
3-(4-sulfamoylphenyl)-4-(3,4-dichlorophenyl)-2(5H)-furanone (compound 50) was synthesized from 4-sulfamoylphenylacetic acid and 2-bromo-1-(3'4-dichlorophenyl)acetophenone in a manner similar to that described for compound 47, with a 30% yield.
Cell Culture
The human prostate cancer cell lines LNCaP and PC-3 were purchased from the American Type Culture Collection (ATCC; Manassas, VA). Bcl-2-overexpressing PC-3 cells were prepared as previously described (26). Cells were cultured in RPMI-1640 medium (Gibco, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS; Gibco) at 37 °C in a humidified incubator containing 5% CO2. Cells were replenished daily with fresh medium and were harvested by trypsinization and split at a 1 : 4 ratio with fresh medium every 3 days.
Cell Viability Analysis
Prostate cancer cells were grown in 10% FBS-supplemented RPMI-1640 medium for 48 hours to approximately 60% confluency. The cells were then washed in serum-free RPMI-1640 and incubated in serum-free RPMI-1640 medium that contained various concentrations of celecoxib, rofecoxib, or test agent, each dissolved in 0.1% dimethyl sulfoxide (DMSO). Control cell cultures were washed in serum-free RPMI-1640 and then incubated in serum-free RPMI-1640 that contained the same concentration of DMSO as the celecoxib-treated cells. Floating cells were recovered from culture medium by centrifugation at 3200g for 5 minutes, and adherent cells were harvested by trypsinization. Both the floating and adherent cells were observed for morphologic changes with a light microscope at x200 magnification. We combined the adherent and floating cells and measured their viability by using a trypan blue dye exclusion assay.
Assessment of Apoptosis
Enzyme-linked immunosorbent assay (ELISA) to detect DNA fragmentation. We used the Cell Death Detection ELISA kit (Roche Diagnostics, Mannheim, Germany), according to the manufacturers instructions, to measure the induction of apoptosis in human prostate cancer cells treated with various compounds. This assay quantifies cytoplasmic histone-associated DNA fragments (both mono- and oligonucleosomes) that result from the induction of apoptosis. In brief, 2.5 x 106 PC-3 cells were plated in T-75 flasks and incubated for 24 hours. The cells were washed twice with 5 mL of serum-free RPMI-1640 medium and then incubated with serum-free medium containing the test compounds as described above. We then collected and pooled the floating and adherent cells, as described above, and counted them. Cell lysates equivalent to 104 cells were used for the ELISA analysis. Histone-associated DNA fragments were quantitated spectrophotometrically using antibodies against DNA and histones in a colorimetric assay.
Western blot analysis of PARP cleavage. PC-3 cells treated with DMSO or the various compounds as described above were collected, washed with ice-cold phosphate-buffered saline (PBS), and resuspended in 50 µL of lysis buffer (20 mM TrisHCl [pH 8], 137 mM NaCl, 1 mM CaCl2, 10% glycerol, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 100 µM 4-(2-aminoethyl)benzenesulfonyl fluoride, 10 µg/mL leupeptin, and 10 µg/mL aprotinin) for 10 minutes. Soluble cell lysates were collected after centrifugation at 1500g for 5 minutes. Protein concentrations of the lysates were determined by using a Bradford protein assay kit (Bio-Rad, Hercules, CA); equivalent amounts of protein from each lysate were resolved in 10% SDSpolyacrylamide gels and then transferred to nitrocellulose membranes. Western blotting with an anti-PARP antibody was carried out as described below, and apoptosis was detected by monitoring proteolysis of the 116-kd native PARP enzyme to the apoptosis-specific 85-kd fragment.
Western Blot Analysis of Apoptosis Signaling Components
Treated cells collected as described above, washed with PBS, resuspended in SDS gel-loading buffer (100 mM TrisHCl [pH 6.8], 4% [wt/vol] SDS, 0.2% [wt/vol] bromophenol blue, 20% [vol/vol] glycerol, and 200 mM dithiothreitol), sonicated with an ultrasonic sonicator for 5 seconds, and boiled for 5 minutes. After brief centrifugation, equivalent amounts of soluble protein, as determined by the Bradford method, were resolved in 10% SDSpolyacrylamide minigels and transferred to nitrocellulose membranes with the use of a semidry transfer cell (Bio-Rad). The membranes were washed twice with TBS (0.3% [wt/vol] Tris, 0.8% [wt/vol] NaCl, and 0.02% [wt/vol] KCl) containing 0.05% Tween 20 (TBST) and then incubated with TBS containing 5% nonfat dry milk for 60 minutes to block nonspecific antibody binding. Each membrane was then incubated at 4 °C for 12 hours with a primary antibody specific for Akt, phospho-Akt, ERKs, phospho-ERKs, or PARP, which was diluted 1 : 1000 in TBS containing 1% nonfat dry milk. The membranes were washed twice with TBST and then incubated at room temperature for 1 hour with a horseradish peroxidase-conjugated goat anti-rabbit immunoglobulin G (IgG) diluted 1 : 5000 in TBS containing 1% nonfat dry milk. The membranes were washed twice with TBST, and bound antibody was visualized by enhanced chemiluminescence using ECLTM western blotting detection reagents (Amersham Pharmacia Biotech, Little Chalfont, U.K.). Unphosphorylated Akt and ERK2, as immunostained by anti-Akt and anti-ERK2 antibodies, were used as internal standards for the comparison of phospho-Akt and phospho-ERK2 levels among samples of different exposure intervals.
Molecular Modeling Experiments
Molecular structures of compounds 4750, as well as celecoxib and rofecoxib, were initially subjected to 1000 steps of Monte Carlo simulation using the Merck Molecular Force Field program available as part of Macromodel 7.0 (Schrodinger, Portland, OR; http://www.schrodinger.com). The minimum conformation reached by the Monte Carlo simulations was then fully optimized at a density functional theory level of B3LYP/631G* with Gaussian 98A7 (Gaussian, Inc., Pittsburgh, PA) (32). All the fully optimized structures were confirmed by normal mode analysis; no negative frequencies were found. Computations for electrostatic potential and electron density were then carried out for each of the fully optimized structures with a grid of 216 000 points using Gaussian 98A7. The electrostatic potential maps for each compound were generated by gOpenMol (http://staff.csc.fi/~laaksone/gopenmol/gopenmol.html) (33,34) and are presented with the electrostatic potential mapped onto the electron density. The electron density isosurface value was 0.0004 with a range of 0.03 to 0.03 for the electrostatic potential.
Statistical Analysis
Each experiment was performed in triplicate and was repeated at least two times on different occasions. Analyses included nonparametric and parametric techniques to include analysis of variance (ANOVA) for linear models of dose response, ANOVA with Scheffé post hoc comparisons, KruskalWallis nonparametric ANOVA, and Spearman rank correlation. A two-sided alpha of 0.05 was considered statistically significant.
We used two descriptive termsthe T50% and the apoptosis indexto express the apoptosis-inducing activity of individual compounds in PC-3 cells. T50% denotes the time required for eliciting apoptotic death in 50% of the cells when they are exposed to a specific concentration of the test compound. The apoptosis index is a semiquantitative indication of the apoptosis-inducing activity of a compound and was defined according to the T50% for cells exposed to test compound at 50 µM. Compounds were classified into the following four categories of apoptosis index (in descending order of apoptosis-inducing activity): +++, T50% is less than or equal to 2 hours; ++, T50% is greater than 2 hours but less than or equal to 4 hours; +, T50% is greater than 4 hours but less than or equal to 24 hours; and -, induced no appreciable apoptosis at 24 hours. Cut points to define the categories of apoptosis indices were determined by first performing ANOVA for all T50% values for the compounds that elicited an apoptotic effect. Although there were statistically significant differences in apoptosis index among the different compounds (P<.001), no differences were seen among LNCaP, PC-3, and Bcl-2-overexpressing PC-3 cells (P = .49). To simplify the identification of the apoptosis index categories, we separated groups of compounds into those that had an apoptosis index of 2 hours or less and those that had an apoptosis index of greater than 2 hours, using the rounded group midpoint value of 2 hours that was determined from a conservative Scheffé post hoc comparison. Because compounds were evaluated in all three cell lines only at a dose of 50 µM, no doseresponse relationship was analyzed.
The purpose of using these two descriptive terms was twofold. First, these two parameters, along with the IC50 (concentration for 50% inhibition) for COX-2 inhibition, allowed us to confirm our previous finding that COX-2 inhibition and apoptosis induction were independent. Second, the use of these terms allowed us to identify compounds that had high apoptosis-inducing potencies.
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RESULTS |
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The objective of this structurefunction analysis of celecoxib and rofecoxib was twofold: to elucidate the structural basis underlying the discrepancy in apoptosis-inducing potency between the COX-2 inhibitors celecoxib and rofecoxib and to optimize the activity of celecoxib with regard to apoptosis induction. To attain these goals, we systematically altered the structure of celecoxib using the strategies to modify the moieties depicted in Fig. 1, A. In strategy A, we either modified the terminal aromatic ring of celecoxib with various substituents to produce compounds 223, or replaced the terminal aromatic ring with different ring systems to produce compounds 2429 (see Fig. 3
for structures). In strategy B, we substituted the carboxamide group for the sulfonamide group of various apoptosis-active celecoxib derivatives to produce compounds 3039 (see Fig. 5
for structures). In strategy C, we modified the heterocyclic system of celecoxib to produce compounds 4046 (Fig. 1, B
). Molecular structures of celecoxib and rofecoxib were also subjected to computer modeling to examine the effect of surface potential on apoptosis; compounds 4750 were prepared, on the basis of computer modeling of those compounds (see Fig. 1, B
, and 6, C
, for structures).
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Role of the Sulfamoyl Moiety of Celecoxib in the Induction of Apoptosis
We tested COX-2 inhibitors that have similar IC50 values for COX-2 inhibition for their apoptosis-inducing activities in PC-3 cells and found that these COX-2 inhibitors exhibited widely discrepant apoptosis-inducing activities (Fig. 2, A). On the basis of their respective apoptosis indices (AIs) in PC-3 cells, we classified these COX-2 inhibitors into two groupsthose that induced apoptosis (e.g., celecoxib) and those that did not (e.g., rofecoxib, NS398, DuP697, and the terphenyl derivative of DuP697, compound 1) (Fig. 2
). Structurally, these COX-2 inhibitors could be classified into two groups according to the type of sulfonyl (-SO2-) functionality they contained, i.e., sulfonamide (-SO2NH2) or methylsulfone (-SO2CH3). As shown in Fig. 2, A
, celecoxib contains a sulfonamide group, whereas rofecoxib, NS398, DuP697, and compound 1 all contain a methylsulfone group. This structural difference suggests that the sulfamoyl moiety of celecoxib may play a role in its apoptosis-inducing activity. This possibility was strengthened by our demonstration that the methylsulfone-containing counterpart of celecoxib was a less potent inducer of apoptosis than celecoxib (Fig. 2, B
, left panel). Moreover, modification of compound 1 to compound 1-NH2 (28) resulted in an increase in apoptosis-inducing activity by an order of magnitude (Fig. 2, B
, right panel). The conversion of rofecoxib and DuP697 to their sulfonamide counterparts, however, did not substantially affect the apoptosis-inducing activities of those compounds (data not shown), indicating that structural elements other than the sulfonamide group contributed to the activation of the apoptosis machinery by celecoxib and compound 1-NH2.
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We synthesized a series of celecoxib derivatives (i.e., compounds 229) to assess the role of the terminal phenyl ring on the apoptosis-inducing activity of celecoxib (Fig. 3). We found that the structural requirements for the induction of apoptosis by these compounds included a certain degree of bulkiness and hydrophobicity in the 5-aryl ring and were distinct from those required for COX-2 inhibition (29). For example, when we reduced the size of the 5-aryl ring (i.e., from a CH3 group in celecoxib to an H group in compound 2 or an F group in compound 3) or increased its polarity (e.g., from a CH3 group to an OH group in compound 11, an NH2 in compound 12, or an NO2 group in compound 13), the apoptosis indices of the resulting compounds decreased precipitously. Among the 28 derivatives we examined, compounds 8, 10, 14, and 1921 had an apoptosis index of +++ in PC-3 cells; compounds 6, 7, 9, 15, and 18 had an apoptosis index of ++; and the remaining compounds displayed no appreciable apoptosis induction at 24 hours (i.e., an apoptosis index of -).
Celecoxib induces apoptosis by a mechanism that involves the concomitant dephosphorylation of Akt and ERK2 (2527). We therefore examined whether the compounds with the most potent apoptotic indices induced cell death by a mechanism similar to that of celecoxib. Fig. 4, A, depicts the time- and/or dose-dependent effects of one of the active compounds generated by strategy A, compound 10, on cell viability. ELISA analysis of the lysates from drug-treated cells revealed the time-dependent formation of oligonucleosomes as a result of DNA degradation (Fig. 4, B
, left panel). In addition, immunoblot analysis of PARP indicated that exposure of PC-3 cells to compound 10 led to the rapid cleavage of the 116-kd native enzyme to form the apoptosis-specific 85-kd fragment (Fig. 4, B
, right panel). It is also noteworthy that the mechanism by which compound 10 caused apoptosis, i.e., dephosphorylation of Akt and ERK2 (Fig. 4, C
), was the same as that by which celecoxib causes apoptosis. Similar results were obtained with other active compounds in this group. The doseresponse relationship for compound 10, with respect to cell viability, was statistically significant (ANOVA, P<.001 for time, dose, and cell viability). Fig. 4, B
, shows that PC-3 cells treated with compound 10 displayed statistically significantly higher levels of ELISA-detectable nucleosome formation, an indicator of apoptotic cell death, over time than did PC-3 cells treated with DMSO (ANOVA; P<.001; mean readings [n = 3] at OD450 at 10, 20, and 30 minutes were 2.6 [95% CI = 2.5 to 2.7], 4.1 [95% CI = 4.0 to 4.2], and 4.1 [95% CI = 4.0 to 4.2], respectively, for drug-treated cells and 0.21 [95% CI = 0.18 to 0.24], 0.22 [95% CI = 0.20 to 0.24], and 0.16 [95% CI = 0.15 to 0.17], respectively, for DMSO-treated cells.
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Apoptosis-Inducing Activities of Celecoxib Analogues That Contain a Carboxamide Moiety in Place of the Sulfamoyl Moiety
Although both the sulfonamide and methylsulfone pharmacophores showed comparable potency in COX-2 inhibition (29), the apoptosis-inducing activity of celecoxib was abrogated when the sulfamoyl moiety was replaced by a methylsulfonyl group. The result suggested that the sulfonamide pharmacophore conferred optimal potency with regard to apoptosis induction. We further investigated whether this functional group could be replaced by a carboxamide moiety without abrogating apoptosis-inducing activities of the resulting compounds. Accordingly, we determined the apoptosis indices for compounds 3039, which possess a carboxamide group in place of the sulfonamide group present in compound 6, compounds 810, celecoxib, compound 15, and compounds 1821. As shown in Fig 5, A, replacement of the sulfonamide group in compounds 8, 9, 10, and 19 with a carboxamide group to produce compounds 31, 32, 33, and 37, respectively, had no substantial effect on the potency of the resulting compounds in apoptosis induction. However, for the rest of the compounds examined (i.e., compounds 30, 34, 35, 36, 38, and 39), the replacement of the sulfonamide group with a carboxamide group resulted in a substantial reduction in their apoptosis-inducing activities. This observation suggested that these two pharmacophores (i.e., carboxamide and sulfonamide) may cause some compounds that contain them to interact differently with the target protein(s) that affect apoptosis. With compound 37 as a representative of this class of derivatives, Fig. 5, B
, shows evidence of drug-induced apoptotic death, which included time-dependent effects on nucleosomal formation (treated versus control, ANOVA, P<.001) and PARP cleavage. Moreover, the structural modification of compound 19 to compound 37 did not alter the mechanism by which this carboxamide-containing compound mediated apoptosis, i.e., by facilitating Akt and ERK2 dephosphorylation (Fig. 5, C
). Similar results were obtained with the other active compounds in this group.
Contributions of the Heterocyclic System to the Apoptosis-Inducing Activity of Celecoxib
The sulfonamide-containing counterparts of rofecoxib and DuP697 did not show substantial apoptosis-inducing activities in PC-3 cells (data not shown). This finding indicated that structural components other than the sulfonamide group may play a role in interacting with the target protein(s) responsible for apoptosis. To shed light on this issue, we further examined the effect of a number of benzenesulfonamides with different heterocyclic rings (i.e., compounds 4046) on the viability of PC-3 cells. Despite the presence of the sulfonamide group, none of these compounds displayed appreciable apoptosis induction at 24 hours (data not shown). Both replacement of the pyrazole ring with other heterocyclic systems and removal of the trifluoromethyl moiety from the pyrazole ring of celecoxib eliminated the apoptosis-inducing activity of the resulting compounds. This finding indicates the effect of the heterocyclic system on the apoptosis-inducing activity.
Molecular Modeling
The above data prompted us to examine how the heterocyclic ring might contribute to the interaction of celecoxib with the signaling target(s) responsible for apoptosis. We therefore conducted a molecular modeling analysis of the two prototypic drugs celecoxib and rofecoxib to examine the electrostatic potentials that surround the heterocyclic systems in these compounds (Fig. 6, A and B). The electron density of individual areas is colored blue to indicate negative electrostatic potentials and red to indicate positive electrostatic potentials. Changes in electrostatic potential from negative to positive are seen in transition from blue to red. As shown in Fig. 6, A and B
, the pyrazole and lactone rings had opposite electron density profiles. This finding suggests that the heterocyclic ring in rofecoxib is more electropositive than the heterocyclic ring in celecoxib.
On the basis of this computer modeling data, we attempted to alter the surface potential of rofecoxib to mimic that of celecoxib by repositioning the lactone carbonyl group in the opposite orientation (Fig. 6, C). The total electrostatic potential map of the resulting isomer, compound 47, was similar to that of celecoxib (Fig. 6, A versus C
). However, because compound 47, which contained a methylsulfone group, showed poor activity in eliciting apoptosis in PC-3 cells (apoptosis index, -) (data not shown), we also modeled the surface electrostatic potentials of the corresponding sulfonamide-containing compound, compound 48, and its dichloro- analogues, compounds 49 and 50. Compound 49, which had a substantially higher apoptosis index (T50% at 50 µM = 15 hours; apoptosis index, +) than rofecoxib, compound 48, or compound 50 (each of which showed no appreciable apoptosis induction at 24 hours; apoptosis index, [data not shown]), had an electrostatic potential map that strongly resembled that of its pyrazole counterpart, compound 8 (apoptosis index, +++) (Fig. 3
). In addition, the similarity in the electrostatic potential profile of compound 49 and compound 8 (Fig. 6, D and E
) was akin to the similarity in the profiles of celecoxib and compound 47, but improved on the 5-aryl moiety. These data demonstrate that a clear understanding of the stereo-electronic characteristics of the entire conjugated system may provide a novel way of associating structural changes with the apoptosis-inducing activities of these compounds.
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DISCUSSION |
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Our statistical analysis of data from three different prostate cancer cell lines suggests that the induction of apoptosis by the active compounds was not dependent on androgen sensitivity, p53 functional status, or the level of Bcl-2 expression. The effectiveness of these agents against androgen-independent prostate cancer (i.e., PC-3) cells is especially noteworthy. Metastatic prostate cancers are lethal because they are heterogeneously composed of both androgen-dependent and androgen-independent malignant cells (35,36). Because androgen-independent prostate cancer cells are resistant to the induction of apoptosis by androgen ablative therapy, an important strategy in developing effective chemotherapy for metastatic prostate cancer is to specifically eliminate androgen-independent cells by targeted apoptosis (35,36). The apoptotic action of these celecoxib derivatives against androgen-independent prostate cancer cells underscores their unique signaling mechanism in disrupting multiple signaling pathways (i.e., Akt and ERK2) that are essential to cancer cell survival (26,27).
In addition to its effects on apoptosis induction, celecoxib has effects on angiogenesis (3740). It has been reported that celecoxib suppresses corneal blood vessel formation in a rat model via a COX-2-dependent mechanism (38). However, our preliminary data indicate that celecoxib derivatives that can induce apoptosis in prostate cancer cells but that lack COX-2 inhibitory activity can also inhibit angiogenesis in the yolk sac of chicken embryos, with potencies comparable with or higher than that of celecoxib (Kulp S, Lin H, Zhu J, Ward P, Chen K, and Chen C: unpublished results). This finding suggests that the anti-angiogenic activity of celecoxib may, in part, be attributable to a COX-2-independent pathway. In view of the crucial role of Akt and ERK2 signaling in embryonal angiogenesis (41), it is possible that the anti-angiogenic effects of celecoxib and its derivatives are mediated through a mechanism similar to the one that induces apoptosis, i.e., Akt and ERK2 dephosphorylation.
It is important to note that, when given at therapeutic doses (oral administration of 400800 mg per day), celecoxib reaches peak plasma concentrations of 38 µM (42), severalfold lower than the concentrations required to induce apoptosis in prostate cancer cells in serum-free medium (i.e., 25 µM or higher). It is conceivable that the observed in vivo antitumor activity of celecoxib may arise from the concerted action of multiple mechanisms that include both the induction of apoptosis and the inhibition of angiogenesis. Consequently, our current research focuses on discerning the relative contributions of these mechanisms to the in vivo effects of celecoxib and its derivatives on tumor growth. In addition, investigations of the pharmacokinetic, pharmacodynamic, and toxicity profiles of these apoptosis-inducing agents are under way.
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NOTES |
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REFERENCES |
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Manuscript received March 11, 2002; revised August 27, 2002; accepted September 19, 2002.
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