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Ligand, 15-Deoxy-
12,14- prostaglandin J21
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-
12,14-prostaglandin J2
(15d-PGJ2) is a highly specific activator of the peroxisome
proliferator-activated receptor 
(PPAR-). We investigated the
effect of 15d-PGJ2 on three human prostate cancer cell
lines, LNCaP, DU145, and PC-3. Western blotting demonstrated that
PPAR-1 is expressed predominantly in untreated prostate cancer
cells. Treatment with 15d-PGJ2 caused an increase in the
expression of PPAR-2, whereas PPAR-1 remained at basal levels.
PPARs 
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- activators induced cell death in all three cell
lines at concentrations as low as 2.5 µM (similar to the
Kd of PPAR- for this ligand), coinciding
with an accumulation of cells in the S-phase of the cell cycle.
Activators for PPAR- 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 
-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 -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-, PPAR-ß (or
NUC-1 and PPAR-
), and PPAR-. In addition, two isoforms of
PPAR-, PPAR-1 and PPAR-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-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-2 is abundant in adipocytes and is thought to be relatively
specific for this tissue (18
, 21
, 22)
. PPAR-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- (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- (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-.
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-; linoleic acid
(23
, 27)
, an activator for PPAR-ß;
15d-PGJ2 (24
, 25) , an activator for
PPAR-; 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- agonist ciglitazone, as well as two -3 polyunsaturated
fatty acids, EPA and DHA, which are natural PPAR- 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. 1
). In comparison, 15d-PGJ2 showed a potent
inhibitory effect (at 
2.5 µM or less) on all cell
lines tested (Fig. 1)
. 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- 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 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)
.
|
, 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).
|
, 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--specific polyclonal antibody demonstrated the expression of
the receptor in all three human cell lines (Fig. 3a)
. The PPAR- antibody clearly recognizes both the 1
and 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-2 was observed as early as 6 h. By 24 h, an
even larger increase in the level of PPAR-2 protein had occurred in
PC-3 and DU145 cell lines (Fig. 3a)
. Moreover, when DU145
cells were treated at the highest dose of
15d-PGJ2 (10 µM), a
significant increase in the expression of PPAR-2 was seen,
especially by 24 h of treatment. The expression level of PPAR-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- (Fig. 3b)
. Tissues from three different patients
were examined, and one representative blot is shown. The analysis
showed that PPAR-1 protein can be detected in all tissues tested at
relatively constant levels, and PPAR-2 is not present in these
tissues. As controls for PPAR expression, DU145 cells treated for
6 h with 10 µM 15d-PGJ2 and differentiated
3T3-L1 cells are shown in Fig. 3b
.
|
and ß isoforms of PPAR. There was no
detectable level of expression of either PPAR- 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-.
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)
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. 4a
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).
|
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- 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)
. 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.
|
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- 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- and C/EBP-ß,
there was no effect of 15d-PGJ2 by 3 h in
all cell lines studied (Fig. 5a)
. Moreover, by 6 and 24 h, the levels of both
C/EBP- 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)
. 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)
.
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.
|
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)
, 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.
|
. 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)
. 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.
|
, 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)
.
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)
.
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 f
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)
. 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 2
. 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- has a direct effect on mitochondrial
function.
|
|
A principle role for PPAR- 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-, 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- 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-2 is up-regulated by low concentrations of
15d-PGJ2 in a time-dependent manner. PPAR-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-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-2 in prostatic cancer cell
death is not clear.
The obvious difference between PPAR-2 and PPAR-1 is 30 additional
amino acids at the NH2 terminus of the 2
isoform (18
, 19)
. Werman et al.
(48)
showed that the NH2 terminus of
PPAR-2 activates a heterologous promoter to a greater degree than
PPAR-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- 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-2 is
influenced more by changes in the nutritional status of an animal, such
as obesity or starvation (48)
. Because we showed that
PPAR-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- were reportedly expressed in prostate
cancer tumors (49)
; therefore the availability of ligand
may be an important factor for the action of PPAR- as a negative
regulator of prostate cancer cell growth. Production of certain
prostaglandins may be modulated by the amount of -3
versus -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- 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- 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-, 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- 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-. 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-,
15d-PGJ2, induces nonapoptotic cell death in
human prostate cancer cells. We suggest that PPAR- 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- 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- polyclonal antibody
(BioMol, Plymouth Meeting, PA; 1:2000 dilution), anti-C/EBP
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. 
2 Present address: Department of Clinical
Neurosciences, Institute of Psychiatry, De Crespigny Park, Denmark
Hill, London SE5 8AF, United Kingdom.
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
4 The abbreviations used are: PPAR, peroxisome
proliferator-activated receptor; 15d-PGJ2,
15-deoxy-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.
5 R. Butler and C. Y-F. Young, unpublished
observation.
Received for publication 5/28/99. Revision received 12/ 8/99. Accepted for publication 12/ 8/99.
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0.05. 
, red
fluorescence; 
, green fluorescence.
m
in 15d-PGJ2-treated cells (relative to control cells)