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Cell Growth & Differentiation Vol. 11, 49-61, January 2000
© 2000 American Association for Cancer Research


Articles

Nonapoptotic Cell Death Associated with S-Phase Arrest of Prostate Cancer Cells via the Peroxisome Proliferator-activated Receptor {gamma} Ligand, 15-Deoxy-{Delta}12,14- prostaglandin J21

Rachel Butler2, Susan H. Mitchell, Donald J. Tindall and Charles Y. F. Young3

Departments of Urology [R. B., S. H. M., D. J. T., C. Y. F. Y.] and Biochemistry [D. J. T., C. Y. F. Y.] and Molecular Biology, Mayo Clinic/Foundation, Rochester, Minnesota 55905

Abstract

15-Deoxy-{Delta}12,14-prostaglandin J2 (15d-PGJ2) is a highly specific activator of the peroxisome proliferator-activated receptor {gamma} (PPAR-{gamma}). We investigated the effect of 15d-PGJ2 on three human prostate cancer cell lines, LNCaP, DU145, and PC-3. Western blotting demonstrated that PPAR-{gamma}1 is expressed predominantly in untreated prostate cancer cells. Treatment with 15d-PGJ2 caused an increase in the expression of PPAR-{gamma}2, whereas PPAR-{gamma}1 remained at basal levels. PPARs {alpha} and ß were not detected in these cells. Lack of lipid accumulation, increase in CCAAT/enhancer binding proteins (C/EBPs), or expression of aP2 mRNA indicated that adipocytic differentiation is not induced in these cells by 15d-PGJ2. 15d-PGJ2 and other PPAR-{gamma} activators induced cell death in all three cell lines at concentrations as low as 2.5 µM (similar to the Kd of PPAR-{gamma} for this ligand), coinciding with an accumulation of cells in the S-phase of the cell cycle. Activators for PPAR-{alpha} and ß did not induce cell death. Staining with trypan blue and propidium iodide suggested that, although the plasma membrane appears intact by electron microscopy, disturbances are evident as early as 2 h after treatment. Mitochondrial transmembrane potentials are significantly reduced by 15d-PGJ2 treatment. In addition, treatment with 15d-PGJ2 resulted in cytoplasmic changes, which are indicative of type 2 (autophagic), nonapoptotic programmed cell death.

Introduction

Prostate cancer is the most common cancer in American men and the second leading cause of male cancer death (1, 2, 3) . There is much experimental and epidemiological evidence to suggest that dietary fat can influence the incidence rate of prostate cancer. Essential fatty acids have been implicated in the development and progression of advanced prostate cancer (4, 5, 6, 7, 8) . In recent years, different series of prostaglandins, the products of arachidonic fatty acid metabolism, have been shown to have either positive or negative effects on cancer cell growth (9 , 10) . Diets rich in fish and olive oils, containing {omega}-3 fatty acids, appear to be linked to a lower incidence of prostate cancer compared with the high incidence in the Western world, where the diets include a high intake of corn oil, which contains {omega}-6 fatty acids.

Many unsaturated, long chain fatty acids and their metabolites, including prostanoids and synthetic analogues, have been demonstrated to act as ligands/hormones via the PPAR4 . PPAR is a member of the steroid hormone receptor superfamily involved in the ligand-inducible regulation of lipid metabolism (11, 12, 13, 14) . PPAR, retinoic acid, vitamin D, and thyroid hormone receptors belong to type II nuclear receptors (15 , 16) . There are three types of PPARs: PPAR-{alpha}, PPAR-ß (or NUC-1 and PPAR-{delta}), and PPAR-{gamma}. In addition, two isoforms of PPAR-{gamma}, PPAR-{gamma}1 and PPAR-{gamma}2, are expressed in human tissues as a result of alternate transcription start sites and alternative splicing (17) . These two isoforms differ only in their NH2 termini in that PPAR-{gamma}2 has an additional 30 amino acids (18, 19, 20) . The expression of the two isoforms is differentially regulated in a tissue-specific manner. PPAR-{gamma}2 is abundant in adipocytes and is thought to be relatively specific for this tissue (18 , 21 , 22) . PPAR-{gamma}1 is also highly expressed in adipocytes; however, lower levels of expression have been detected in a number of other tissues, including muscle and liver (21, 22, 23) . All PPAR isoforms require heterodimerization with the retinoid X receptor for optimal DNA binding and transcriptional activity (15 , 16 , 23) .

Although several long-chain fatty acids can activate PPARs, they are not effective activators for PPAR-{gamma} (23, 24, 25, 26, 27) . However, a fatty acid metabolite, 15d-PGJ2, the terminal metabolite of the prostaglandin J series, has been shown to be a highly specific activator of PPAR-{gamma} (25 , 26) . Prostaglandin D2 and prostaglandin J2 are endogenous metabolites of the prostaglandin J series that have already been shown to have antitumor activities (11) . These observations led us to study the effect of 15d-PGJ2 on the growth of prostate cancer cells with the hypothesis that the action of these prostaglandins may be occurring through PPAR-{gamma}.

Results

15d-PGJ2 Inhibits the Growth of Prostate Cancer Cell Lines.
Lipids that are known to be specific activators of the three isoforms of PPAR were used to establish the involvement of the particular isoforms in the growth response mechanism in human prostate adenocarcinoma cells. Hormone sensitive (LNCaP) and hormone refractory (PC-3 and DU145) prostate cancer cell lines were treated with ETYA (22 , 26) , an activator for PPAR-{alpha}; linoleic acid (23 , 27) , an activator for PPAR-ß; 15d-PGJ2 (24 , 25) , an activator for PPAR-{gamma}; and prostaglandin E2 (23 , 25, 26, 27) , an inert activator for PPARs over a range of concentrations (1 to 100 µM). Additionally, the synthetic PPAR-{gamma} agonist ciglitazone, as well as two {omega}-3 polyunsaturated fatty acids, EPA and DHA, which are natural PPAR-{gamma} activators, were included in the studies. The effect of each ligand on the growth response of the cells was determined using the MTS cell viability assay. Neither prostaglandin E2 nor linoleic acid had any apparent effect on the viability of any of the cell lines examined. ETYA had an inhibitory effect on all three cell lines but only at the highest concentration tested (100 µM; Fig. 1Citation ). In comparison, 15d-PGJ2 showed a potent inhibitory effect (at ~2.5 µM or less) on all cell lines tested (Fig. 1)Citation . It is important to note that the effective concentration of 15d-PGJ2 needed to reduce cell viability (2.5 µM) is comparable with that of 15d-PGJ2 required for PPAR-{gamma} activation in gene transfer studies (2 µM; Ref. 25 ). At higher concentrations of 15d-PGJ2 (up to 10 µM studied), a profound effect on cell viability was seen. Treatment with EPA, DHA, and ciglitazone also produced a potent growth-inhibitory effect. Ciglitazone produced an inhibitory pattern that was similar to 15d-PGJ2 at comparable concentrations. However, the concentrations of EPA and DHA needed to produce the effect were much higher than that of 15d-PGJ2. Transfection studies (28 , 29) also show that a higher concentration of EPA and DHA is required for PPAR {gamma} activation, although in vitro ligand binding studies demonstrate an affinity (Kd) of ~2 µM for EPA binding. One possible explanation for this is that EPA and DHA have differences in compound stability and binding to fatty acid binding protein compared with 15d-PGJ2. The transportation of EPA and DHA into cells may be mediated by fatty acid transporters (30) .



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Fig. 1. The effects of activators of PPARs on the viability of human prostatic adenocarcinoma cell lines, DU145, PC-3, and LNCaP. Cells were incubated with 0.01–100 µM linoleic acid (an activator of PPAR-ß), ETYA (an activator of PPAR-{alpha}), and prostaglandin E2 (an inert activator for PPARs); 1–10 µM, 15d-PGJ2; 5–100 µM EPA and DHA; and 2–20 µM, ciglitazone (activators of PPAR-{gamma}); or, vehicle controls (column 0) for 6 days. Viable cells were measured by MTS assay and expressed as a percentage of controls (n = 4–6); bars, SD.

 
To confirm the effect of 15d-PGJ2 on cell proliferation, the incorporation of bromodeoxyuridine into DNA was measured in a nonradioactive DNA synthesis assay. As shown in Fig. 2Citation , treatment with 2.5 and 10 µM 15d-PGJ2 decreases DNA synthesis significantly in these cells by 24 h. For PC-3 and DU145 cells, DNA synthesis decreased additionally between 24 and 48 h of treatment. Together these two assays suggest that the effect of 15d-PGJ2 on these cells (also see below) is to induce cell death but not terminal differentiation, as has been suggested for other tumor cell lines (31 , 32) . In addition, these data show that the end point of the cell death induced by 15d-PGJ2 is occurring between 24 and 48 h in the majority of cells. However, the initiation of the 15d-PGJ2 effect occurs very rapidly, with cells starting to round up and detach from the culture dish within 2 h of treatment (data not shown).



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Fig. 2. The effects of 15d-PGJ2 on prostate cancer cell growth analyzed by DNA synthesis. LNCaP, PC-3, and DU145 cells were treated with 0, 2.5, and 10 µM 15d-PGJ2 for 24 and 48 h. Additionally, cells were incubated with 10 µM bromodeoxyuridine solution for 18 h prior to harvesting. The amount of newly synthesized DNA was measured by nonradioactive bromodeoxyuridine ELISA (Roche) and expressed as percentage of controls (n = 4); bars, SD.

 
Expression of PPARs in Prostate Cancer Cell Lines and Tissues.
Because 15d-PGJ2 is a ligand for PPAR-{gamma}, it is reasonable to expect that this receptor is the mediator for the inhibitory growth effect of 15d-PGJ2 on prostate cancer cells. Western blotting analysis of total cell extracts from LNCaP, PC-3, and DU145 prostate cancer cell lines using a PPAR-{gamma}-specific polyclonal antibody demonstrated the expression of the receptor in all three human cell lines (Fig. 3a)Citation . The PPAR-{gamma} antibody clearly recognizes both the {gamma}1 and {gamma}2 isoforms of the receptor. In addition, when cells were treated with low concentrations of 15d-PGJ2 (2.5 µM) over 24 h, a marked increase in the level of PPAR-{gamma}2 was observed as early as 6 h. By 24 h, an even larger increase in the level of PPAR-{gamma}2 protein had occurred in PC-3 and DU145 cell lines (Fig. 3a)Citation . Moreover, when DU145 cells were treated at the highest dose of 15d-PGJ2 (10 µM), a significant increase in the expression of PPAR-{gamma}2 was seen, especially by 24 h of treatment. The expression level of PPAR-{gamma}1 remained unchanged with these treatments and therefore served as an internal control. A Western blot analysis of benign and cancerous prostate tissues was performed to see the expression patterns of PPAR-{gamma} (Fig. 3b)Citation . Tissues from three different patients were examined, and one representative blot is shown. The analysis showed that PPAR-{gamma}1 protein can be detected in all tissues tested at relatively constant levels, and PPAR-{gamma}2 is not present in these tissues. As controls for PPAR{gamma} expression, DU145 cells treated for 6 h with 10 µM 15d-PGJ2 and differentiated 3T3-L1 cells are shown in Fig. 3bCitation .



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Fig. 3. Expression of PPAR-{gamma}1 and PPAR-{gamma}2 isoforms in human prostate cancer cell lines showing the effect of 15d-PGJ2 treatment. a, cells were treated with 2.5 and 10 µM 15d-PGJ2 and harvested at 3-, 6-, and 24-h intervals. Western blots were prepared and probed with anti-PPAR-{gamma} antibody (1:2000; BioMol), which detects both isoforms of PPAR-{gamma}. b, Western blots of benign (B) and cancerous (T) prostate tissue, DU145 cells treated with 10 µM 15d-PGJ2 for 6 h, and differentiated 3T3-L1 cells. The same antibody shown in a was used in b.

 
We also carried out Western blot analysis on these cells to determine the expression levels of the {alpha} and ß isoforms of PPAR. There was no detectable level of expression of either PPAR-{alpha} or PPAR-ß protein in any of the prostate cancer cell lines studied (data not shown). Taken together, these results suggest that 15d-PGJ2 has a specific antiproliferative effect on the growth of both androgen-dependent and androgen-independent prostate cancer cell lines, and that this action is occurring through a pathway involving PPAR-{gamma}.

Treatment of Prostate Cancer Cells with 15d-PGJ2 Causes Changes in Plasma Membrane Integrity.
We observed that the cells round up in a short time after 15d-PGJ2 treatment but do not die completely until after 24 h of treatment (data not shown). This suggests that changes may also be occurring at the level of the plasma membrane. Two staining methods were used to assess whether the plasma membranes of cells treated with 15d-PGJ2 were undergoing permeability changes. Trypan blue staining was performed on cells treated with either 2.5 or 10 µM 15d-PGJ2 for 2, 4, and 6 h (Fig. 4a)Citation and cells treated with 50 µM EPA, 50 µM DHA, and 15 µM ciglitazone at 6 h of treatment. The number of cells staining blue in a total of 500 cells was counted and expressed as a percentage. Fig. 4aCitation shows that by 2 h of treatment with both 2.5 and 10 µM 15d-PGJ2, all three cell lines had a significant number of trypan blue-stained cells compared with controls. LNCaP cells were the most affected by 15d-PGJ2 treatment in this assay. By 6 h of treatment, ~80% of the LNCaP cells treated with 15d-PGJ2 are capable of taking up trypan blue dye. The effect of 15d-PGJ2 occurred in a dose-dependent manner. After 6 h of treatment with EPA, LNCaP, PC-3, and DU145 cells took up 99, 97, and 91% of trypan blue dye, respectively (data not shown). After treatment with DHA, 98, 97, and 95% of LNCaP, PC-3, and DU-145 cells took up trypan blue, respectively (data not shown). After ciglitazone treatment, 59, 92, and 51% of LNCaP, PC-3, and DU145 cells took up trypan blue dye (data not shown). Some cells appeared to be surrounded by a ring of trypan blue dye at the membrane surface. These cells were counted as trypan blue positive and may account for the discrepancy between the number of cells stained by trypan blue and PI (see below).



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Fig. 4. Effects of 15d-PGJ2 treatment on plasma membrane integrity assessed by trypan blue exclusion assay and PI staining. LNCaP, PC-3, and DU145 cells treated with 0, 2.5, and 10 µM 15d-PGJ2 were stained with: a, trypan blue dye; or b, PI and Hoechst 33258 at 2, 4, and 6 h after treatment. The number of blue (a) or red (b) cells per total of 500 was counted and expressed as a percentage of the total (n = 4).

 
The second assay involved staining with two DNA-binding dyes, PI and Hoechst 33258, which can be viewed with fluorescence microscopy. In addition to showing nuclear morphology by binding DNA, PI can also be used to determine plasma membrane integrity, because this dye is a polar molecule and cannot enter the cell unless the plasma membrane is perturbed in some manner. In contrast, Hoechst 33258 can freely enter cells and stain DNA to show nuclear morphology. Therefore, these two dyes were used to test the plasma membrane integrity and to visualize any changes in nuclear morphology (see below) at various time points after 15d-PGJ2 treatment. Staining with PI (Fig. 4Citation b) correlated well with trypan blue, taking into account the discrepancy described above. As with the trypan blue staining, 15d-PGJ2 treatment had a profound effect on the ability of LNCaP cells to take up PI, indicating an effect on the integrity of the plasma membrane. Together, these data suggest that a relatively early effect of the PPAR-{gamma} ligand, 15d-PGJ2, is to cause disturbances in the plasma membrane, resulting in a loss of integrity and eventually cell death. However, the disturbance of the plasma membrane may not be a direct effect of 15d-PGJ2, because this effect occurs gradually and over an extended period of time,.

15d-PGJ2 Treatment Causes S-Phase Cell Cycle Arrest in Prostate Cancer Cell Lines.
Because of the profound inhibition of cellular proliferation caused by 15d-PGJ2, we wanted to determine whether the cell cycle was altered. LNCaP, PC-3, and DU145 cells were treated with 2.5 and 10 µM 15d-PGJ2 for 24 and 48 h, at which time cell cycle changes were studied by FACS analysis. All three cell lines showed a significant accumulation of cells in the S-phase by 24 h of treatment with 15d-PGJ2 compared with control cells (Table 1)Citation . However, the androgen-dependent cell line, LNCaP, exhibited a lesser effect than the androgen-independent cell lines DU145 and PC-3; although compared with control cells, there was approximately three times the number of treated cells in S-phase. LNCaP cells grow very slowly in culture and therefore may not have completed an entire cell cycle before the effects of 15d-PGJ2 occur.


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Table 1 Cell cycle analysis of prostate cancer cell lines treated with 15d-PGJ2

Cells were treated with 2.5 and 10 µM 15d-PGJ2 for 24 and 48 h and then analyzed by FACS. n = 4 for 10 µM 15d-PGJ2 treatment and n = 2 for 2.5 µM 15d-PGJ2 treatment. Values are mean ± SD.

 
The completion of cell cycle events in an orderly manner, from the G1 stage to DNA replication in S-phase or from the G2 stage to mitosis, is controlled by many factors, which act as checkpoints to ensure that no mistakes occur (33) . Therefore, in addition to carrying out FACS analysis to study the cell cycle, we also studied changes in the protein expression of certain cell cycle regulating factors over a time course of 15d-PGJ2 treatment. The cell cycle regulators studied were p53, p21, and p27, all of which are involved in controlling various checkpoints in the cell cycle. However, no changes were observed in the expression levels of any of these proteins over the time period in which cell cycle arrest occurred (data not shown).

After observing the 15d-PGJ2-induced cell cycle changes mentioned above, we were interested to see whether 15d-PGJ2 could induce differentiation of prostate cancer cells into adipocytes, as has been described for other cell types. The C/EBPs, C/EBP-{alpha} and C/EBP-ß, are nuclear transcription factors, which are vital components for the differentiation of preadipocytes into adipocytes (34, 35, 36, 37) . It has been demonstrated that some compounds, such as the thiazolidinediones (31 , 37, 38, 39) , generally up-regulate the expression of these proteins. When we examined the levels of C/EBP-{alpha} and C/EBP-ß, there was no effect of 15d-PGJ2 by 3 h in all cell lines studied (Fig. 5a)Citation . Moreover, by 6 and 24 h, the levels of both C/EBP-{alpha} and C/EBP-ß were significantly decreased compared with controls. To confirm that there is no adipocytic program initiated by 15d-PGJ2 a Northern blot analysis for aP2 was performed (Fig. 5b)Citation . Differentiated 3T3-L1 cells were used as a positive control. The figure shows that there is no up-regulation of the aP2 mRNA by 15d-PGJ2 treatment in either LNCaP or DU145 cells. PC-3 cells also exhibited no up-regulation (data not shown). In addition, we tested for the accumulation of neutral lipids with oil red O dye. When LNCaP, PC-3, and DU145 cells were treated for 4 days with a range of concentrations of 15d-PGJ2, no differences were detected in the amount of oil red O staining between treated cells and controls (data not shown). These results suggest that 15d-PGJ2 does not induce differentiation of these cells into adipocytes. A Northern blot analysis of PSA was also performed to prove there is no further differentiation with 15d-PGJ2 treatment (Fig. 5c)Citation . The Northern blot results show that after the up-regulation of PSA by mibolerone in LNCaP cells, 15d-PGJ2 treatment down-regulates the expression of PSA mRNA.



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Fig. 5. Expression of differentiation markers. a, the expression of C/EBP{alpha} and C/EBPß was studied by Western blotting of LNCaP, PC-3, and DU145 cells treated with 0, 2.5, and 10 µM 15d-PGJ2 for 3, 6, and 24 h. Anti-C/EBP{alpha} and anti-C/EBPß antibodies (1:500; Santa Cruz) were used for immunodetection. b, a Northern blot of mRNA in LNCaP, DU145, and differentiated 3T3-L1 cells using a probe for aP2, PSA, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH).

 
Cell Death Induced by 15d-PGJ2 Occurs via a Nonapoptotic Mechanism.
A number of apoptotic assays were performed to investigate the mode of cell death occurring in response to 15d-PGJ2. Nuclear morphology changes, DNA fragmentation, and PARP cleavage were studied at various time points after treatment with 10 µM 15d-PGJ2. No DNA fragmentation or PARP cleavage, both of which are classic events known to occur during apoptosis, was observed in any of the cell lines (data not shown; Refs. 40 and 41 ). Moreover, the use of caspase inhibitors to block or delay apoptosis had no effect on the induction of cell death by 15d-PGJ2 (data not shown). Fig. 6Citation shows an example of Hoechst 33258 and PI staining of LNCaP cells after 6 h of 10 µM 15d-PGJ2 treatment. Clearly, none of the nuclear morphology changes associated with apoptosis have occurred by this time point. Ciglitazone treatment also showed no nuclear morphology changes associated with apoptosis by Hoechst 33258 staining (data not shown). The nuclei from all three cell lines were still intact at 72 h of treatment (data not shown). Moreover, the majority of cells (80%) stained with both PI and Hoechst 33258 (Fig. 6)Citation , and the cells with PI staining also had more intense Hoechst 33258 staining, suggesting that the plasma membrane may have become more permeable even after short exposures (6 h) to 15d-PGJ2.



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Fig. 6. Nuclear staining of LNCaP cells. LNCaP cells were treated with 10 µM 15d-PGJ2 for 6 h and stained with Hoechst 33258 and PI. Cells were photographed from a double-stained population by confocal microscopy. a, Hoechst 33258; b, PI.

 
To delineate whether any further alterations in the plasma membrane occurred and to look for additional ultrastructural changes, transmission electron microscopy was performed on cells from all three cell lines treated with 2.5 and 10 µM 15d-PGJ2 for 24 h. Negative control cells received no treatment. Positive controls for necrosis and apoptosis (42 , 43) were treated with high (20 µM) and low (2 µM) concentrations of the calcium ionophore A23187, respectively. Electron micrographs of representative cells can be seen in Fig. 7Citation . The majority of cells maintained an intact plasma membrane after treatment with 10 µM 15d-PGJ2 for 24 h, unlike those seen in the necrosis control in which almost all cells have completely disrupted membranes and have lost their cellular contents (Fig. 7c)Citation . A small percentage of 15d-PGJ2-treated cells also have this morphology, but this is likely to be attributable to mechanical damage incurred while preparing the cells for electron microscopy.



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Fig. 7. Transmission electron micrographs showing morphological changes in prostate cancer cells treated with 15d-PGJ2. a, LNCaP control; b, DU145 control, note the location of the mitochondria (m) throughout the cytoplasm and the extensive microvilli (mv) in the plasma membrane; c, necrosis control, LNCaP cells treated with 20 µM A23187 for 2 h; d, apoptosis control, DU145 cells treated with 2 µM A23187 for 6 h; e, LNCaP cells treated with 10 µM 15d-PGJ2 for 24 h, showing perinuclear position of mitochondria, vacuolization of the cytoplasm (vac), and distension of endoplasmic reticulum into autophagic vesicles (av); f, DU145 cells treated with 10 µM 15d-PGJ2 for 24 h, showing loss of microvilli (arrow with asterisk), perinuclear location of mitochondria (m) and chromatin condensation; g, LNCaP cells treated with 2.5 µM 15d-PGJ2 for 24 h, showing vacuolization (vac), distended endoplasmic reticulum, autophagic vesicles (av) and chromatin condensation; and h, DU145 cells treated with 2.5 µM 15d-PGJ2, showing loss of microvilli, chromatin condensation, and extensive vacuolization of the cytoplasm (vac).

 
LNCaP, PC-3, and DU145 cells all originate from prostate epithelia and as such have distinct microvilli on their surfaces (particularly clear in DU145 cells). Cells treated with 15d-PGJ2 showed a pronounced loss of plasma membrane microvilli (Fig. 7Citation , compare b to f and h). These changes at the cell surface may be enough to allow the uptake of polar dyes such as PI.

The nuclei of cells remained intact after 15d-PGJ2 treatment in correlation with the Hoechst 33258 nuclear staining and DNA fragmentation data. Chromatin condensation was still evident, although complete segregation has not occurred to give the pyknotic structures seen in apoptotic cells. This can be seen in the apoptosis control cells treated with 2 µM A23187 (Fig. 7d)Citation .

The cytoplasmic changes seen in cells treated with 15d-PGJ2 are possibly more dramatic than the nuclear events. Cytoplasmic changes included extensive vacuolization and distension of the endoplasmic reticulum (Fig. 7, e–h)Citation . In a number of cells, the endoplasmic reticulum was distended to such an extent that "whorl-like" structures formed, and some of these appear to contain organelles, possibly mitochondria, which may be undergoing degradation by autophagy. In addition, a pronounced phenomenon involving the movement of mitochondria from a diffuse cytoplasmic pattern of localization to a tightly packed grouping in a perinuclear position occurred in the majority of cells treated with 10 µM 15d-PGJ2 (Fig. 7, e and fCitation compared with a and b). PC-3 cells showed similar cytoplasmic changes (data not shown).

15d-PGJ2 Causes a Decrease in Mitochondrial Membrane Potential.
The prominent change in mitochondrial position, together with the apparent autophagic degradation of a number of mitochondria in 15d-PGJ2-treated cells, suggests that mitochondrial function may be impaired in these cells. In addition, changes in plasma membrane integrity can also have a profound effect on the stability of mitochondrial function. Therefore, the mitochondrial abnormalities observed by electron microscopy were evaluated further using a specific fluorescent mitochondrial probe, JC-1. JC-1 is a lipophilic cation that normally exists as a monomer emitting green fluorescence. In a reaction driven by mitochondrial transmembrane potential, JC-1 forms dimeric "J-aggregates" that emit a red fluorescence (44 , 45) . In this manner, the use of JC-1 enables the dual analysis of mitochondrial mass (green fluorescence) and mitochondrial transmembrane potential (red fluorescence). After treatment with 15d-PGJ2, we observed a progressive increase in green fluorescence, which correlated with a decrease in red fluorescence at both concentrations of the ligand in all cell lines studied (Fig. 8)Citation . As expected, this effect was more profound at 10 µM 15d-PGJ2 than at the lower concentration. However, at 24 h, the extent of the loss of mitochondrial transmembrane potential inferred by the decreasing red fluorescence and increasing green fluorescence was much greater in PC-3 and DU145 cells than in the LNCaP cell line. These data, together with the red:green fluorescence ratio, which represents the net transmembrane potential per mitochondrion (46) , are presented in Table 2Citation . The fact that the profound reduction in mitochondrial transmembrane potential correlates with the time frame in which cell viability is lost after 15d-PGJ2 treatment suggests that this endogenous ligand for PPAR-{gamma} has a direct effect on mitochondrial function.



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Fig. 8. Effect of 15d-PGJ2 treatment on the mitochondrial transmembrane potential of prostate cancer cells. LNCaP, PC-3, and DU145 prostate cancer cells were treated with 15d-PGJ2 for 6 and 24 h. Mitochondrial transmembrane potential was measured using a mitochondria-specific fluorescent probe, JC-1. Red and green fluorescence were measured simultaneously by FACS analysis [excitation at 488 nm; emission 530 nm (green fluorescence) and 585 nm (red fluorescence)]. Events (20,000) were measured, and the number of red or green events is expressed as a percentage of the total (n = 3). P <= 0.05. , red fluorescence; {blacksquare}, green fluorescence.

 

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Table 2 Mean fluorescence values of mitochondrial mass and {Delta}{psi}m in 15d-PGJ2-treated cells (relative to control cells)

Green and red fluorescence are expressed as a ratio of treated:control. Decreasing red fluorescence indicates a loss of mitochondrial transmembrane potential in correlation with increasing green fluorescence ratio.

 
Discussion

A principle role for PPAR-{gamma} is to trigger the differentiation cascade resulting in the formation of mature adipocytes from preadipocytes. However, the fact that this receptor is expressed in a variety of normal and tumor tissues suggests an additional role. Because of the antitumor effects observed by other members of the J series of prostaglandins, we were interested in the effect of 15d-PGJ2 treatment on the growth of prostate cancer cell lines. We also wanted to elucidate whether the mechanism of action was occurring through the PPAR-{gamma}, for which 15d-PGJ2 is a ligand. In our study, treatment with 15d-PGJ2 results in cell death at high and low concentrations in both hormone-sensitive and hormone-resistant human prostate cancer cells.

Our studies have demonstrated that PPAR-{gamma} is expressed in three of the commonly studied metastatic prostate cancer cell lines, LNCaP, PC-3, and DU145 (47) , as well as in human benign and cancerous prostatic tissues. Interestingly, we have found that PPAR-{gamma}2 is up-regulated by low concentrations of 15d-PGJ2 in a time-dependent manner. PPAR-{gamma}2 usually is expressed more specifically in fat cells. However, we could not demonstrate any adipogenic differentiation of treated prostate cancer cells. In addition, at 10 µM of 15d-PGJ2, PPAR-{gamma}2 was only inducible in DU145 cells. Although the phenomenon seems to be very interesting, no plausible explanation can be offered at the present time. Thus, the role of differentially expressed PPAR-{gamma}2 in prostatic cancer cell death is not clear.

The obvious difference between PPAR-{gamma}2 and PPAR-{gamma}1 is 30 additional amino acids at the NH2 terminus of the {gamma}2 isoform (18 , 19) . Werman et al. (48) showed that the NH2 terminus of PPAR-{gamma}2 activates a heterologous promoter to a greater degree than PPAR-{gamma}1. However, the activation domain was mapped to a common region in both isoforms. As for many nuclear receptors, the expression of specific coactivators in a particular cell type has been suggested as a mechanism to confer functional specificity to receptors or isoforms of receptors that bind the same response element. Therefore, it is likely that two isoforms of PPAR-{gamma} are used to enable different factors to regulate their expression and/or activity to meet specific biological requirements. Of the two isoforms, the expression of PPAR-{gamma}2 is influenced more by changes in the nutritional status of an animal, such as obesity or starvation (48) . Because we showed that PPAR-{gamma}2 can be differentially regulated by different concentrations of 15d-PGJ2 in prostate cancer cells, the regulatory mechanisms may deserve further investigation.

Recently, high levels of PPAR-{gamma} were reportedly expressed in prostate cancer tumors (49) ; therefore the availability of ligand may be an important factor for the action of PPAR-{gamma} as a negative regulator of prostate cancer cell growth. Production of certain prostaglandins may be modulated by the amount of {omega}-3 versus {omega}-6 fatty acids ingested in the diet. Although the amount of 15d-PGJ2 in the prostate has not been determined, the prostate gland does produce large amounts of prostaglandins (50, 51, 52) . In fact, the prostate has the capacity to produce prostaglandin D2, which can be spontaneously converted to 15d-PGJ2, the terminal metabolite of this prostaglandin series (53) . Therefore, the modulation of 15d-PGJ2 via "good" fat in the diet presents a novel therapeutic and potential preventive method for treating prostate cancer. However, PPAR-{gamma} can be activated by a number of ligands; therefore, the level of 15d-PGJ2 alone may not be as important as the total amount of available ligand.

PPAR-{gamma} is known to cause cell cycle withdrawal preceding adipocyte differentiation (54) . Cell cycle withdrawal in preadipocytes correlates with a large decrease in the DNA binding and transcriptional activity of E2F/DP, a growth-related transcription factor. Because the discovery of natural and synthetic ligands for PPAR-{gamma}, a potential therapy for a number of cancers has been suggested. This therapy uses a mechanism of growth inhibition by terminal differentiation of the cancer cells into adipocytes via PPAR-{gamma} activation. Although this theory has shown potential in breast and liposarcoma cancers (31, 32 , 55) , we and others have found no evidence that terminal differentiation is occurring in prostate cancer cells (49) . However, the accumulation of cells in the S-phase of the cell cycle after 15d-PGJ2 treatment (more pronounced for PC-3 and DU145 cell lines) was unexpected. Because other members of the prostaglandin J series have been shown to cause G1 arrest at lower doses and at higher concentrations cause G2-M arrest leading to apoptosis, we thought similar events would occur with 15d-PGJ2 treatment (56 , 57) . Many assays used to detect apoptotic events showed that this was not the case. In fact, the effect of 15d-PGJ2 on prostate cancer cells is the induction of rapid detachment of the cells by 6 h of treatment and ending with the majority of cells dead at 24 h.

PI and trypan blue staining assays showed that polar dyes were able to be taken up by a large majority of the cells at about the same time that detachment from the culture dish was occurring, which taken alone could be indicative of necrosis. Studies at the electron microscopy level showed that many of the typical signs of necrosis are not occurring in these prostate cancer cells treated with 15d-PGJ2. In fact, many of the morphological changes occurring in these cells as a result of 15d-PGJ2 treatment, for example, loss of microvilli, vacuolization of the cytoplasm, chromatin condensation without nuclear fragmentation, and loss of cytoplasmic structures are indicative of type 2 (autophagic) programmed cell death (58) . This form of cell death is characterized primarily by the formation of autophagic vacuoles, together with the occasional dilation of mitochondria and endoplasmic reticulum (59 , 60) . Although autophagic vacuoles were seen in only a small percentage of cells treated with 15d-PGJ2, dilated endoplasmic reticulum was observed in almost all cells. Autophagy does not directly destroy the plasma membrane (by definition) or the intact nucleus, probably because of its size. However, plasma membrane changes are seen in cells undergoing autophagic cell death. This is more pronounced in epithelial cells that lose microvilli and/or junctional complexes (60, 61, 62, 63) and can clearly be seen to occur in our cell culture system upon treatment with 15d-PGJ2. This may account for the loss of plasma membrane integrity inferred by the uptake of PI and trypan blue dyes. The nuclear degradation reported in autophagic cell death is by no means as prevalent or striking as that seen in apoptosis, although in some cases of these types of cell death pyknotic nuclei are reported (60 , 64) . In all cases, the nuclei of the treated cells in our study remained intact, although a certain amount of chromatin condensation was seen to occur. The lack of DNA fragmentation after nuclear condensation in these cells suggests that they may already have lost their ATP because of decreased mitochondrial function (see below) and plasma membrane integrity. Therefore, they are not able to further process chromatin, which requires energy, and proceed along the apoptotic pathway (65) .

One interpretation of the role of autophagy is that it protects the cell rather than destroying it by degrading restricted parts of the cytoplasm through autolysis and segregation, thus protecting the rest of the cell (59) . This protective mechanism may be occurring in these prostate cancer cells treated with 15d-PGJ2 because this treatment also induces the expression of heat shock protein 70,5 which plays a role in the stress response of cells and may be up-regulated in an attempt to evade death (66, 67, 68, 69) .

For many years, it has been known that mitochondria play a central role in the mechanism of necrotic cell death, but only recently has this role been extended to include apoptosis (reviewed in Refs. 70 and 71 ). Although we believe that the mechanism of cell death occurring in our system is neither necrosis nor apoptosis, the involvement of mitochondria in the mechanism of cell death cannot be ruled out. Mitochondria are the primary organelles targeted for autophagy in type 2 cell death and appear to be engulfed by dilated endoplasmic reticulum in some of the cells in our study. Therefore, we looked at the functioning of mitochondria after treatment with 15d-PGJ2 by studying their transmembrane potential. JC-1 is a fluorogenic molecule widely used for the purpose of measuring mitochondrial transmembrane potential (44 , 45) . Treatment with 15d-PGJ2 does indeed greatly reduce the mitochondrial transmembrane potential in all three cell lines, starting at 6 h of treatment. By 24 h, the effect is greatly pronounced, particularly in DU145 and PC-3 cells. It has been shown that the expression of an important group of proteins, the uncoupling proteins, involved in the uncoupling of oxidative phosphorylation from ATP synthesis, is regulated by PPAR-{gamma}. The uncoupling proteins are transmembrane proteins found in the inner mitochondrial membrane, predominantly in brown adipose tissue and skeletal muscle (72) . Whether these uncoupling proteins play a role in the decreased mitochondrial transmembrane potential observed in our cells remains a subject of ongoing investigation in our laboratory.

In addition to decreased mitochondrial function in cells after 15d-PGJ2 treatment, we also observed the clustering of mitochondria in a perinuclear position rather than the dispersed cytoplasmic distribution as seen in control cells. Mitochondria and microtubules have long been documented to colocalize in many cell types (reviewed in Ref. 73 ). The involvement of microtubules in the positioning of mitochondria is supported by evidence that mitochondria redistribute in mammalian cells treated with microtubule destabilizing agents (74 , 75) . The microtubule-based motor proteins, kinesin and cytoplasmic dyenin, bind microtubules and transduce chemical energy into mechanical work as they hydrolyze ATP to enable polarized movement of "cargo" along microtubules (76) . Recently, disruption of kinesin motor activity has been shown to cause a perinuclear pattern of mitochondrial localization attributable to the loss of polarized movement to the periphery of the cell (77 , 78) . Therefore, we suggest that 15d-PGJ2 may affect mitochondrial functioning by causing mitochondrial membrane depolarization, leading to the uncoupling of oxidative phosphorylation from ATP synthesis and ultimately to cell death.

In summary, we demonstrate that a ligand for PPAR-{gamma}, 15d-PGJ2, induces nonapoptotic cell death in human prostate cancer cells. We suggest that PPAR-{gamma} is a negative regulator of prostate cancer cell growth. It is apparent from our studies that both mitochondrial and plasma membrane disturbances are involved in the mechanism of cell death. However, further investigation into the downstream events activated by PPAR-{gamma} to induce cell death is required if modulation of ligands for this receptor is to form a significant means of preventing or treating prostate cancer.

Materials and Methods

Cell Culture
LNCaP, DU145, PC-3, and 3T3-L1 cells were obtained from the American Type Culture Collection. All cells were maintained in a humidified atmosphere of 95% air and 5% CO2 at 37°C. Cells (7.5 x 104 per ml) were seeded in 24-well culture plates in RPMI 1640 supplemented with 5% serum and incubated for 3 days. The medium was changed to serum-free/phenol red-free RPMI 1640 24 h prior to treatment. To differentiate 3T3-L1 cells (79) , the cells were treated with 0.25 µM dexamethasone, 0.5 mM 1-methyl-3-isobutylxanthine, and 1 µg/ml insulin. After 48 h, cells were switched to 10% FCS medium containing 1 µg/ml insulin. Then cells were collected, as described below, for Western blotting.

Chemicals
All prostaglandins and fatty acids were purchased from Sigma Chemical Co. (St. Louis, MO) with the exception of 15d-PGJ2 (Caymen Chemicals, Ann Arbor, MI) and ciglitazone (Biomol, Plymouth Meeting, PA). Stock solutions were prepared and stored according to the manufacturer’s specifications.

Assays of Growth Inhibition
Cell Viability Assay.
After treatment with ligands at varying concentrations (described in the text), the cells were incubated for an additional 6 days. Cell viability was determined using the MTS colorimetric assay (Promega, Madison, WI). MTS assay reagents (mixed at 1 part phenazine methosulfate to 20 parts MTS, according to manufacturer’s instructions) were added to the culture medium at a 1:6 dilution (200 µl/well for 24-well plates and 20 µl/well for 96-well plates). The cells were incubated for 90 min at 37°C, and the absorbance was measured at 490 nm using a plate reader (80 , 81) . Each experiment was carried out in quadruplicate and repeated at least three times.

DNA Synthesis Assay.
Cell proliferation was determined by measurement of bromodeoxyuridine incorporation during DNA synthesis via a nonradioactive colorimetric assay (Ref. 82 ; Roche, Indianapolis, IN). Cells were treated with 2.5 and 10 µM 15d-PGJ2, and the amount of bromodeoxyuridine incorporation over an 18-h period was measured at 24 and 48 h. The assay was carried out according to the manufacturer’s instructions. The substrate reaction was measured without stop solution at a wavelength of 370 nm on an ELISA plate reader.

Staining Assays for Cell Death and Plasma Membrane Integrity
For all staining assays, cells were treated with 2.5 and 10 µM 15d-PGJ2, stained, and counted at 2, 4, and 6 h.

PI Staining.
Stock solutions of PI (Sigma; 1 mg/ml) were dissolved in PBS and stored at 4°C. Cells were incubated with PI at a final concentration of 1 µg/ml for 5 min at 37°C.

Hoechst 33258 Staining.
Stock solutions of Hoechst 33258 (Sigma; 10 mg/ml) were dissolved in PBS and stored at -20°C. Cells were incubated with Hoechst 33258 at a final concentration of 5 µg/ml for 5 min at 37°C. Cells were washed gently with PBS, and red cells (PI) or blue cells (Hoechst 33258) were visualized by fluorescence microscopy using an Axiophot microscope (Zeiss, Inc.). Appropriate excitation filters were used (PI: 546 nm excitation and 590 nm emission; Hoechst 33258: 365 nm excitation and 420 nm emission). The number of red cells per 500 blue cells was counted and expressed as a percentage of the total.

Trypan Blue Staining.
Adherent cells were harvested by trypsinization and collected by centrifugation. Nonadherent cells were collected from spent media by centrifugation. Cell pellets were resuspended in 100 µl of fresh media, and trypan blue solution (Sigma) was added at a ratio of 1:1. Blue cells were counted as a percentage of a total of 500 cells.

Western Blotting.
Total cell lysates were prepared from each cell line after treatment with 15d-PGJ2. Adherent and nonadherent cells were collected by low-speed centrifugation and washed with PBS. Cell pellets were gently resuspended in RIPA buffer [PBS containing 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS plus freshly added protease inhibitors, 100 µg/ml phenylmethylsulfonyl fluoride, 30 µl/ml aprotinin (from Sigma stock solution), and 1 mM sodium orthovanadate]. Benign and cancerous tissues were homogenized in RIPA buffer. All samples were disrupted and homogenized by passage through a 21-gauge needle and incubated for 30 min on ice. Additional phenylmethylsulfonyl fluoride was added to a final concentration of 100 µg/ml. Samples were centrifuged for 20 min at 15,000 x g in a microcentrifuge at 4°C, and the supernatants (total cell lysates) were collected and stored at -20°C. Protein concentration was determined using a detergent-compatible protein assay (Bio-Rad, Hercules, CA), according to the manufacturer’s instructions. Equivalent amounts of protein were separated in precast SDS-polyacrylamide gels (Novex, San Diego, CA) and transferred to nitrocellulose membranes (Bio-Rad). Transfer and protein loading were checked by Ponceau S staining prior to blocking the membrane with 5% milk/TBS-Tween 20 (1 h at room temperature). Blocked membranes were incubated overnight with either anti-PPAR-{gamma} polyclonal antibody (BioMol, Plymouth Meeting, PA; 1:2000 dilution), anti-C/EBP{alpha} polyclonal antibody (Santa Cruz; 1:500 dilution), or anti-C/EBPß polyclonal antibody (Santa Cruz; 1:500 dilution). Three 10-min washes in TBS-Tween 20 (20 mM Tris-HCI, 137 mM NaCl, and 0.1% Tween 20, pH 7.6) were carried out between antibody steps, followed by incubation with antirabbit secondary antibody conjugated to horseradish peroxidase (Amersham, Arlington Heights, IL; 1:2000 dilution) for 1 h at room temperature. Immunoreactivity was detected using the enhanced chemiluminescence development method (Renaissance; DuPont NEN, Boston, MA).

Cell Cycle Analysis.
Cell cycle analysis was performed on cells treated with 2.5 and 10 µM 15d-PGJ2 for 24 and 48 h. Adherent and nonadherent cells were collected by centrifugation, washed with PBS, and fixed in 95% ethanol for 10 min on ice. The cells were pelleted, washed with PBS, and resuspended in 20 µg/ml PI in PBS containing 200 µg/ml RNase. Samples were incubated for 1 h at 37°C and subjected to FACS analysis (Becton Dickinson, Bedford, MA).

Northern Blotting.
Cells were treated with varying amounts of 15d-PGJ2 and 1 nM mibolerone as indicated, and RNA was collected by the guanidinium isothiocyanate method (83) . An RNA gel was run and transferred onto a nylon membrane according to the GeneScreen protocol by New England Nuclear. Twenty µg of total RNA were loaded in each lane. cDNAs for PSA, aP2, and glyceraldehyde-3-phosphate dehydrogenase were used as probes labeled with [P32]dCTP by random priming. The hybridization was performed by prehybridizing the membranes for 4 h with a hybridization buffer containing 7% SDS, 1 mM EDTA, and 0.25 M sodium phosphate, pH 7.2. The probes were added to fresh buffer and hybridized overnight. The membranes were washed with a washing buffer containing 0.1x SSC + 0.1% SDS. The films were autoradiographed at -70°C.

Oil Red O Staining.
Oil red O staining of neutral lipid accumulation within cells was measured using a spectrophotometric method (84) . Cells were grown in 24-well plates and treated with a range of concentrations of 15d-PGJ2 and incubated at 37°C for 4 days. Nonadherent cells were pelleted by centrifugation in a centrifuge with adaptors for 24-well plates. The medium was removed, and cells were fixed in 3% paraformaldehyde for 1 h at 4°C. Cells were washed with PBS and stained with a 0.5% solution of oil red O (dissolved in methanol and filtered) for 15 min at room temperature. Cells were washed with PBS, and oil red O was extracted by addition of 200 µl isopropanol. The extracted samples were transferred to a clean, 96-well plate, and the absorbance was measured at 510 nm using an ELISA plate reader. The experiment was carried out in quadruplicate and repeated at least three times.

Characterization of Nuclear Morphology.
Cells (7.5 x 104 per ml) were seeded onto 10-cm dishes and incubated as above. After 24 h in serum-free/phenol red-free medium, cells were treated with 10 µM 15d-PGJ2 and incubated for 48 h. The cells were collected by centrifugation, washed twice with PBS, and fixed for 5 min at 4°C with 3% paraformaldehyde in PBS. The cells were collected and resuspended in H2O for 1 min for rehydration and repelleted. The cells were stained with 0.3 µg/ml Hoechst 33258 for 5 min at room temperature. The stained cells were centrifuged and washed once with PBS and resuspended in an appropriate volume of PBS (200 µl; Ref. 85 ). The stained nuclei were viewed by fluorescent microscopy using an Axiophot microscope (Zeiss, Inc.). Appropriate excitation filters were used (365 nm excitation; 420 nm emission).

DNA Fragmentation Gel Electrophoresis.
DNA fragmentation in cells treated with 2.5 and 10 µM 15d-PGJ2 was analyzed by gel electrophoresis at 24-, 48-, and 72-h time points using the method of Gunji et al. (86) . Adherent and nonadherent cells were collected by scraping, followed by centrifugation at 1000 rpm for 10 min. The cell pellets were resuspended in 20 µl of 50 mM Tris-HCl (pH 8.0), 10 mM EDTA, and 0.5 mg/ml proteinase K solution and incubated for 1 h at 50°C. RNase A solution (10 µl of a 0.5 µg/ml stock) was added, and the samples were incubated for 1 h at 50°C. Samples were loaded onto a 1% (w/v) agarose gel after addition of 10 µl of preheated (70°C) loading buffer containing 10 mM EDTA (pH 8.0), 1% (w/v) low-melting point agarose, 0.25% (w/v) bromphenol blue, and 40% (w/v) sucrose. The wells were sealed with 1% (w/v) low-melting point agarose prior to the addition of running buffer to the tank. Samples were electrophoresed at 25 V overnight at 4°C. DNA was visualized by ethidium bromide staining.

Analysis of PARP Cleavage.
Cell lysates from 15d-PGJ2-treated cells were prepared according to the method of Shah et al. (87) with minor modifications. Cells were harvested at various time points, collected by centrifugation, and washed with PBS. Repelleted cells were lysed in sample buffer [62.5 mM Tris-HCl (pH 6.8), 6 M urea, 10% glycerol, 2% SDS, 0.00125% bromphenol blue, and 5% ß-mercaptoethanol, freshly added]. Analysis of the Mr 116,000 PARP protein and its Mr 89,000 cleavage product was carried out by separation on 8% SDS-PAGE gels, followed by Western blotting, as described above. Anti-PARP polyclonal antibody (1:2000 dilution; Roche) was used to visualize specific bands.

Inhibition of Caspase Activity.
Cells were treated with a caspase inhibitor, over a range of concentrations (10–200 µM), for 1 h prior to treatment with 5 and 10 µM 15d-PGJ2. The caspase inhibitor used was Z-Val-Ala-DL-Asp-fluoromethylketone dissolved in DMSO (Bachem, Torrance, CA; Ref. 88 ). Inhibition of cell death was assessed by the MTS assay at various time points as described previously.

Transmission Electron Microscopy.
Cells were treated with 2.5 and 10 µM 15d-PGJ2 for 24 h prior to preparation for electron microscopy. Apoptosis and necrosis control samples were prepared by treating the cells with the calcium ionophore A23187 for 2 h at 2 and 20 µM, respectively. Cells were rinsed in warm PBS, harvested by gentle scraping, and pelleted by centrifugation. Cell pellets were resuspended in Trump’s fixative (1% gluteraldehyde and 4% formaldehyde in 0.1 M phosphate buffer, pH 7.2; Ref. 89 ), preheated to 37°C for 1 h at room temperature. Pelleted cells were rinsed three times with 0.1 M phosphate buffer (pH 7.2) and incubated in 1% osmium tetraoxide (phosphate buffered) for 1 h. Cells were washed with rinse buffer, followed by three washes with double-distilled H2O. Samples were incubated with 1% uranyl acetate for 1 h at room temperature, dehydrated in graded ethanol, infiltrated, and embedded in 100% Spurr’s resin (90) . Thin (90-nm) sections were cut on a Reichert Ultracut E ultramicrotome, placed on mesh copper grids, and stained with lead citrate. Micrographs were taken on a JEOL 1200 EXII transmission electron microscope (JEOL, Peabody, MA) operating at 60 kV.

Mitochondrial Membrane Potential Analysis.
Mitochondrial function was assessed indirectly by measuring the variation in mitochondrial transmembrane potential measured by JC-1 (Molecular Probes, Eugene, OR) red fluorescence using flow analysis (44 , 45) . Both red and green fluorescence emissions were analyzed after JC-1 staining using the method of Mancini et al. (46) . Cells treated with 2.5 and 10 µM 15d-PGJ2 for 6 and 24 h were collected by trypsinization, followed by centrifugation. Cell pellets were resuspended in 500 µl of medium containing 10 µg/ml JC-1 and incubated for 10 min at 37°C before flow analysis. FACScan (Becton Dickinson, Bedford, MA) was used to establish size gates and exclude cellular debris. For each experimental time point, 15d-PGJ2-treated and control cells were analyzed. The excitation wavelength was 488 nm. The emission wavelengths were 530 nm for green fluorescence and 585 nm for red fluorescence. Events (20,000) were analyzed per sample, and the relative change in mean fluorescence was calculated as the ratio of the 15d-PGJ2-treated to control samples.

Footnotes

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.

1 R. B., S. H. M., and C. Y-F. Y. are supported by NIH Grants DK41995 and CA70892 and by Department of Defense Grant DMAD 17-98-107-523. D. J. T. is supported by a National Cancer Institute grant to the Mayo Comprehensive Cancer Center and a grant from the T. J. Martell Foundation. Back

2 Present address: Department of Clinical Neurosciences, Institute of Psychiatry, De Crespigny Park, Denmark Hill, London SE5 8AF, United Kingdom. Back

3 To whom requests for reprints should be addressed, at Department of Urology, 17 Guggenheim, Mayo Clinic, 200 First Street SW, Rochester, MN 55905. Phone: (507) 284-8336; Fax: (507) 284-2384; E-mail: youngc{at}mayo.edu Back

4 The abbreviations used are: PPAR, peroxisome proliferator-activated receptor; 15d-PGJ2, 15-deoxy-{Delta}12,14-prostaglandin J2; ETYA, 5,8,11,14-eicosatetraenoic acid; EPA, eicosapentaenoic acid; DHA, docosahexanaenoic acid; MTS, 3-(4,5-dimethylthiazol-2-yl)-5-(carboxymethoxyphenyl)-2(4-sulophenyl)-2H-tetrazolium; FACS, fluorescence-activated cell sorter; C/EBP, CCAAT/enhancer binding proteins; PI, propidium iodide; PARP, poly(ADP-ribose) polymerase; JC-1, 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidoazolcarbocyanine iodide; PSA, prostate-specific antigen. Back

5 R. Butler and C. Y-F. Young, unpublished observation. Back

Received for publication 5/28/99. Revision received 12/ 8/99. Accepted for publication 12/ 8/99.

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