# Activation of Penile Proadipogenic Peroxisome Proliferator-Activated Receptorγ with an Estrogen: Interaction with Estrogen Receptor Alpha during Postnatal Development

Authors: Mahmoud M. Mansour; Hari O. Goyal; Tim D. Braden; John C. Dennis; Dean D. Schwartz; Robert L. Judd; Frank F. Bartol; Elaine Coleman S.; Edward E. Morrison
Journal: PPAR Research
 (2008)

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## Abstract

Exposure to the estrogen receptor alpha (ERα) ligand diethylstilbesterol (DES) between neonatal days 2 to 12 induces penile adipogenesis and adult infertility in rats. The objective of this study was to investigate the in vivo interaction between DES-activated ERα and the proadipogenic transcription factor peroxisome proliferator-activated receptor gamma (PPARγ). Transcripts for PPARs α, β, and γ and γ1a splice variant were detected in Sprague-Dawley normal rat penis with PPARγ predominating. In addition, PPARγ1b and PPARγ2 were newly induced by DES. The PPARγ transcripts were significantly upregulated with DES and reduced by antiestrogen ICI 182, 780. At the cellular level, PPARγ protein was detected in urethral transitional epithelium and stromal, endothelial, neuronal, and smooth muscular cells. Treatment with DES activated ERα and induced adipocyte differentiation in corpus cavernosum penis. Those adipocytes exhibited strong nuclear PPARγ expression. These results suggest a biological overlap between PPARγ and ERα and highlight a mechanism for endocrine disruption.


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## Body

## 1. Introduction

Endocrine disruption, originally limited to steroid receptor signaling,
now extends to include other
members of the 48 reported nuclear receptor superfamily [1]. Both
peroxisome proliferator-activated receptor gamma (PPARγ) and estrogen receptor alpha (ERα) are targets for endocrine disrupting
chemicals [2–4].
Recently, Goyal et al. showed that neonatal exposure of rats to the estrogenic
endocrine disruptor diethylstilbestrol (DES) induced adipogenesis in penile
corpus cavernosum by activation of ERα [5–8]. In this model of DES-ERα activation, DES exposure at a dose of 0.1 to 0.12 mg/kg
bw/day, on alternate days, from postnatal days 2 to12, resulted in infertility
in 100% of the treated male rats. Loss of fertility was associated with
abnormal accumulation of fat cells in the corpus cavernosum penis, and the
associated loss of cavernous spaces apparent as early as postnatal day 18 (reviewed in [9]). It remains unknown, however, whether this penile ERα-induced adipogenesis is mediated by activation of a constitutively
expressed or DES-induced  PPARγ.Both ERα and PPARγ  pathways
are implicated in fat regulation. First, recent findings suggest that
PPARγ and ERα  pathways involve
shared coactivators that promote differentiation of preadipocytes into mature
fat cells. For example, constitutive
coactivator of PPARγ (CCPG) is described as a bona fide coactivator that cross reacts with ERα independent of its ligand and contains four LXXLL motifs that are
characteristic of nuclear receptor coactivators [10]. Second, studies have shown that forced
expression of PPARγ2 or PPARγ1 can trigger the differentiation of fibroblasts
to adipocytes resulting in the activation of adipocyte-specific genes and lipid
accumulation [11].The PPAR family
consists of three isotypes that include PPARα (NR1C1), PPARβ (also known as
PPARδ, NR1C2, FAAR, or NUC-1), and PPARγ  (NR1C3) [12–14].
A nuclear receptor, PPARγ, is known to play a central role in fat metabolism and adipocyte
differentiation [15, 16].
The PPARγ is present in two key
isoforms, PPARγ1 and PPARγ2. The two isoforms stem from alternate
promoters [17]. Compared to PPARγ1, PPARγ2
has an additional 30 amino acids at the N-terminal end and is distinctively
expressed in adipose tissue, where it plays a key role in adipogenesis [18]. These nonsteroidal receptors
(i.e., do not mediate effects of steroids) form part of a class I nuclear hormone
receptor superfamily [19] and function as
ligand-activated transcription factors [20–22].Each of
the three PPAR isotypes is constitutively expressed in certain reproductive and
nonreproductive rat tissues [23, 24],
but their temporal and cell-specific expression in penile tissue, with the
exception of a limited demonstration of PPARγ
in penile corporal smooth muscle cells [25], has not been shown. Further,
no specific link is known between neonatal activation of ERα and penile PPARγ. This is
important given the expanded definition of the term endocrine disruptors to
include activation of metabolic sensors such as PPARs. A number of findings
suggest involvement of PPARs in endocrine disruption either through direct receptor
activation or indirectly through crosstalk with other nuclear receptors.
First, in vitro studies demonstrated that PPARγ
and ERα (the iconic receptor involved
in endocrine disruption) are implicated in cross-talk [26–28].
Second, some endocrine disruptor chemicals, such as monethylhexyl phthalate
(MEHP), a primary metabolite of diethylhexyl phthalate (DEHP), mediate their
toxic effect by PPARγ activation [29, 30].
Third, several nonbiological xenobiotics compounds can activate PPARγ. For example, activation of PPARγ with synthetic
PPARγ activators, such as antidiabetic drugs thiazolidinediones (TZDs), improve insulin sensitivity but they undesirably increase
preadipocyte differentiation and white adipose tissue mass  [31–33]. Consistent with this
adipogenic effect, reduced PPARγ
level, as in mice with heterozygous (PPARγ+/−)
deficiency, is associated with
reduced white adipose tissue mass [34].Findings related to interaction
between ERα and PPARγ in the aforementioned DES-penile rat
model will illuminate a potential molecular mechanism by which estrogen exposure
at critical period of development
perturbs reproductive tissues. Therefore, we hypothesize that DES-induced
penile adipogenesis is associated with ERα-mediated
activation of PPARγ. Objectives of
this study were to (1) determine the basal expression of PPARs (α, β, and γ)
in rat penis and (2) evaluate the neonatal modulatory effect of ERα-activator DES on
penile PPARγ  as a marker of undesirable adipogenesis.


## 2. Materials and Methods

### 2.1. Animals and Treatments

This DES study was performed in collaboration with Dr. Hari Goyal at Tuskegee University using male pups from
pregnant female Sprague-Dawley (SD) rats (Harlan Sprague-Dawley, Indianapolis, Ind, USA).
All animal procedures were approved by Institutional Animal Care and Use
Committee at Tuskegee University. In all
experiments, rats were maintained using standard housing conditions including
constant temperature of 22°C, ad libitum water and feeding, and 12:12 hours light dark cycle. Two experiments were conducted. In
experiment 1, three groups of male pups (n=5 per group, all were littermates)
received subcutaneous injections of 25 μL
of olive oil (control), oil containing DES (0.1 mg/kg, Sigma-Aldrich, St. Louis, Miss, USA),
or DES plus ICI 182, 780 (16.6 mg/kg, ICI; Tocris Bioscience, Ellisville, Miss, USA)
daily on postnatal day 2 to 6. Rats in experiment 1 were sacrificed at 28 day of age. ICI 182, 780 is a high-affinity estrogen
receptor antagonist (IC50= 0.29 nM) and is also considered a high-affinity ligand for the membrane estrogen receptor GPR30 (Tocris Bioscience).
In experiment 2, two groups of male
pups (n=4 per group) received DES (1 mg/kg) or olive oil (control) every
other day for 6 days starting at postnatal day 2. Penile tissues were
collected from rats sacrificed at 120 days of age (adulthood). Small sections
of the penile shaft tissue from each rat in experiment 1 and 2  were fixed overnight in 4% paraformaldehyde
for IHC or fat staining, and the remainder of the shaft tissue was frozen in
liquid nitrogen and stored at −80°C for RNA extraction and PCR
analysis. The doses used for end-point evaluation at 28 and 120 days
post-treatment were based on previous publications from our group that showed
DES prenatal exposure (between postnatal days 2 to12) at a dose range of 0.1 to
0.12 mg/kg/day, or higher (1 mg/kg/day) result in similar abnormal penile
development and adipogenesis [5, 8].


### 2.2. Total RNA Isolation

Total RNA was isolated from the body of the penis using
TRIZOL reagent (Invitrogen-Life Technologies Inc., Carlsbad, Calif, USA),
according to the manufacturer's protocol. RNA concentrations were estimated at 260 nm and
the ratio of 260/280 was determined using UV spectrophotometry (DU640, Beckman
Coulter Fullerton, Calif, USA).
The integrity of each RNA sample, indicated by the presence of intact 28S and
18S ribosomal RNA, was verified by denaturing agarose gel electrophoresis. RNA
samples were treated with DNase (Ambion Inc.) to remove possible genomic DNA
contamination. Samples with 260/280 ratio of ≥1.8 were used.


### 2.3. Conventional End-Point and Real-Time PCR

Expression of mRNA for PPAR (α, β, and γ) isotypes
was initially determined by conventional end-point RT-PCR with primers
designed using primer quest software and synthesized
by Integrated DNA Technology (IDT Inc, Coralville, Iowa, USA) from
previously published rat sequences (see Table 1). Subsequently,
semiquantitative RT-PCR for coamplification of PPARs and S-15 (known as Rig;
small subunit ribosomal protein used as a house keeping gene) was performed to
determine the relative expression levels of PPAR isotypes. Verification of
accurate PCR products was confirmed by determination of the expected size of
PCR bands and by sequence analysis of generated amplicons at Auburn University sequencing
facility. The resulting sequences for the three PPAR isotypes were
matched with previously published rat sequences in GenBank (accession number NM013196, U40064,
and NM013124 for PPARα, PPARβ, and PPARγ, resp.) using Chromas 2.31 software (Technelysium
Pty ltd, Tewantin Qld 4565, Australia). PPARγ
splice variants or subtypes were identified using specific primers
designed for rat PPARγ1a and PPARγ1b synthesized by IDT Inc. (Table 1).
Liver and white adipose tissues from adult Sprague-Dawley rats in experiment 2
were used as positive controls for PPARγ1
[35] and PPARγ2 [18], respectively. The amplification protocol was as follows:
initial cycle for 3 minutes at 95°C, and 30 cycles each at
(95°C for 30 seconds, 55°C for 30 seconds, and 72°C
for 30 seconds) followed by a final extension cycle at 72°C for 7
minutes. PCR reactions were performed on a Robocycler (Stratagene Inc, La Jolla, Calif, USA)
and products were analyzed electrophoretically on 2% (w/v) agarose gels. The
intensity of the PCR bands was determined using Fluor-S multi-imaging
analysis system (Bio-Rad, Hercules, Calif, USA). Level of mRNA for PPARs was
normalized to the levels of S-15 housekeeping gene.Table 1
PCR primer sets, sequence, product size (bp), nucleotide (nt) location, and GenBank accession numbers for rat
PPARs used in this study. Note that a common antisense oligoprimer (sequence in bold) was used for PPARγ1a, PPARγ1b, and PPARγ2.
Product/Sense primerAntisense primerProduct sizentaccession#(bp)locationPPARα5-TTG TGA CTG GTC AAG CTC AGG ACA-35-TCG TAC GCC AGC TTT AGC CGA ATA-3492296–787NM013196PPARβ5-TAA CGC ACC CTT CAT CAT CCA CGA-35-TTG ACA GCA AAC TCG AAC TTG GGC-3390873–1262U40064PPARγ5-TCT CCA GCA TTT CTG CTC CAC ACT-35-ATA CAA ATG CTT TGC CAG GGC TCG-3533257–789NM013124PPARγ1a5-CTG ACG AGG TCT CTC TC G GCT G-35-AGC AAG GCA CTT CT GAA ACC GA-365821–679AF246458PPARγ1b5-CAG CGC TAA ATT CAT CTT AAC T-35-AGC AAG GCA CTT CTG AA A CCG A-361821–639AF246457PPARγ25-GAG CAT GGT GCC TTC GCT GA-35-AGC AAG GCA CTT CTG AA A CCG A-356337–600AB01956185–648AF15666686–649Y12882PPARγ/ERα(primers for real-time PCR were obtained from Superarray Inc)190/179[PPARγ/ERα]NM013124 [PPARγ](Sequence are not disclosed by the Company)respectivelyNM012689 [ERα]Gapdh5-ATG ATT CTA CCC ACG GCA AG-35-CTG GAA GAT GGT GAT G CGT T-38971–159DQ403053184–272BC087743216–304BC059110Rig/S15 (Ambion)5-TTC CGC AAG TTC ACC TAC C-35-CGG GGC CGG CCA TGC T TTA CG-336174–433BC105810Quantitative real-time PCR (Bio-Rad, MyiQTM) for determination of
expression levels of PPARγ and ERα mRNA was performed in 25-μL reaction
mixture containing 12.5 μL RT2 real-time
SYBR/Fluorescein Green PCR master mix, 1 μL
first strand cDNA, 1 μL RT2 validated PCR primer set for PPARγ
or ERα (Super Array Bioscience
Corporation, Frederic, Md, USA), and 10.5 μL
PCR-grade water (Ambion Inc). Samples were run in 96-well PCR plates (Bio-Rad,
Hercules, Calif, USA) in duplicates, and the results were normalized to
GAPDH (see primer set in Table 1) expression. The
amplification protocol was set at 95°C for 15 minutes for one cycle,
and 40 cycles each at (95°C for 30 seconds, 55°C for 30
seconds, and 72°C for 30 seconds) followed by melting curve determination between 55°C and 95°C to
ensure detection of a single PCR product. Template RNA from rat white adipose
tissue and penis were used for determination of amplification efficiencies for
(ERα/PPARγ) targets and GAPDH
by generating standard curves. Curves were generated by using serial 10-fold dilutions total RNA and plotting the log dilution against CT (threshold cycle) value obtained for each dilution. The
Pearson's correlation coefficient (r) value for each generated standard curve
was ≥0.98, and the calculated amplification efficiency was between 98.5 to 99%.


### 2.4. Immunohistochemistry (IHC)

Immunolocalization of PPARγ in penile tissue was performed using mouse anti-PPARγ
IgG1 monoclonal antibody (sc7273, Santa Cruz Biotechnology Inc, Santa Cruz, Calif, USA)
raised against a C-terminus sequence of human and mouse PPARγ  (similar to the corresponding rat sequence).
The antibody detects PPARγ1, PPARγ2 and, to a lesser extent, PPARα and PPARβ of rat, mouse, and
human by IHC using paraplast-embedded tissues. Approximately 5-mm-long penis sections from the middle
of the body of the penis were fixed in 4% paraformaldehyde
for 48 hours, embedded in Paraplast (Sigma-Aldrich), and cut at 5-μm thickness [7].
Mounted penis sections were deparaffinized in Hemo-D (Scientific Safety
Solvents, Keller, Tex, USA) and hydrated to distilled water (dH2O).
The slides were transferred to a rack and placed in 1 L of 10 mM sodium citrate
(pH 6.0). The beaker was placed on a hot plate, allowed to come to a boil and
tissues were boiled for 20 minutes. When the citrate solution cooled to near
room temperature (RT), the slides were transferred to a glass staining dish and
equilibrated in phosphate buffered saline (PBS) (Sigma-Aldrich, ST Louis, Miss, USA). After 20 minutes incubation in blocker (5%
normal goat serum, Sigma-Aldrich) and 2.5% BSA (Sigma) in PBS, slides were washed
briefly in PBS. Anti-PPARγ, diluted 1:20 in blocker, was applied and the
sections were left to incubate overnight at RT. Next day, slides were washed 3x in PBS, 3 minutes each, and tissues were
incubated with Alexa 488-conjugated goat antimouse IgG (Molecular Probes, Eugene, Ore, USA)
for 1 hour at RT. After washing two times in PBS, 3 minutes each, slides were
mounted with VectaShield (Vector Laboratories, Burlingame, Calif, USA),
and the coverslips were sealed. The sections were examined using a Nikon
TE2000E microscope and digital images were generated using an attached Retiga
EX CCD digital camera (Q Imaging, Burnaby, BC, Canada).
Penile tissue sections from all 28-day treated rats were examined. Representative
micrographs from different penile histological structures were shown for
untreated control rats, and for rats treated with DES or DES + ICI.


### 2.5.  Fat Staining

Histochemical demonstration of fat was performed as previously described [7]. Briefly, tissue sections from penile body, approximately 5 mm-long, were
fixed for 24 hours in 4% formaldehyde, followed by en bloc staining of fat for
8 hours with 1% osmium tetroxide dissolved in 2.5% potassium dichromate
solution. Specimens were then processed for paraplast embedding and cut at 5-μm thickness. Deparaffinized sections were examined for
black staining indicative of fat cells using light microscopy.


### 2.6. Statistical Analyses

Analysis of real-time PCR data for relative gene
expression level (fold change of target relative to control) was performed
using a modification of the
delta delta Ct method (ΔΔ CT) as described previously [36]. Statistical differences
between treatment groups were performed using Sigma Stat statistical software
(Jandel Scientific, Chicago, Ill, USA). Δ CT for real-time PCR data [37],
and intensity values (for semi-quantitative RT-PCR data) were subjected to
analyses of variance. Experimental groups with means significantly
different (P<.05) from controls were identified using Holm-Sidak
and Tukey tests. When data were not distributed normally, or heterogeneity of
variance was identified, analyses were performed on transformed or ranked data.


## 2.1. Animals and Treatments

This DES study was performed in collaboration with Dr. Hari Goyal at Tuskegee University using male pups from
pregnant female Sprague-Dawley (SD) rats (Harlan Sprague-Dawley, Indianapolis, Ind, USA).
All animal procedures were approved by Institutional Animal Care and Use
Committee at Tuskegee University. In all
experiments, rats were maintained using standard housing conditions including
constant temperature of 22°C, ad libitum water and feeding, and 12:12 hours light dark cycle. Two experiments were conducted. In
experiment 1, three groups of male pups (n=5 per group, all were littermates)
received subcutaneous injections of 25 μL
of olive oil (control), oil containing DES (0.1 mg/kg, Sigma-Aldrich, St. Louis, Miss, USA),
or DES plus ICI 182, 780 (16.6 mg/kg, ICI; Tocris Bioscience, Ellisville, Miss, USA)
daily on postnatal day 2 to 6. Rats in experiment 1 were sacrificed at 28 day of age. ICI 182, 780 is a high-affinity estrogen
receptor antagonist (IC50= 0.29 nM) and is also considered a high-affinity ligand for the membrane estrogen receptor GPR30 (Tocris Bioscience).
In experiment 2, two groups of male
pups (n=4 per group) received DES (1 mg/kg) or olive oil (control) every
other day for 6 days starting at postnatal day 2. Penile tissues were
collected from rats sacrificed at 120 days of age (adulthood). Small sections
of the penile shaft tissue from each rat in experiment 1 and 2  were fixed overnight in 4% paraformaldehyde
for IHC or fat staining, and the remainder of the shaft tissue was frozen in
liquid nitrogen and stored at −80°C for RNA extraction and PCR
analysis. The doses used for end-point evaluation at 28 and 120 days
post-treatment were based on previous publications from our group that showed
DES prenatal exposure (between postnatal days 2 to12) at a dose range of 0.1 to
0.12 mg/kg/day, or higher (1 mg/kg/day) result in similar abnormal penile
development and adipogenesis [5, 8].


## 2.2. Total RNA Isolation

Total RNA was isolated from the body of the penis using
TRIZOL reagent (Invitrogen-Life Technologies Inc., Carlsbad, Calif, USA),
according to the manufacturer's protocol. RNA concentrations were estimated at 260 nm and
the ratio of 260/280 was determined using UV spectrophotometry (DU640, Beckman
Coulter Fullerton, Calif, USA).
The integrity of each RNA sample, indicated by the presence of intact 28S and
18S ribosomal RNA, was verified by denaturing agarose gel electrophoresis. RNA
samples were treated with DNase (Ambion Inc.) to remove possible genomic DNA
contamination. Samples with 260/280 ratio of ≥1.8 were used.


## 2.3. Conventional End-Point and Real-Time PCR

Expression of mRNA for PPAR (α, β, and γ) isotypes
was initially determined by conventional end-point RT-PCR with primers
designed using primer quest software and synthesized
by Integrated DNA Technology (IDT Inc, Coralville, Iowa, USA) from
previously published rat sequences (see Table 1). Subsequently,
semiquantitative RT-PCR for coamplification of PPARs and S-15 (known as Rig;
small subunit ribosomal protein used as a house keeping gene) was performed to
determine the relative expression levels of PPAR isotypes. Verification of
accurate PCR products was confirmed by determination of the expected size of
PCR bands and by sequence analysis of generated amplicons at Auburn University sequencing
facility. The resulting sequences for the three PPAR isotypes were
matched with previously published rat sequences in GenBank (accession number NM013196, U40064,
and NM013124 for PPARα, PPARβ, and PPARγ, resp.) using Chromas 2.31 software (Technelysium
Pty ltd, Tewantin Qld 4565, Australia). PPARγ
splice variants or subtypes were identified using specific primers
designed for rat PPARγ1a and PPARγ1b synthesized by IDT Inc. (Table 1).
Liver and white adipose tissues from adult Sprague-Dawley rats in experiment 2
were used as positive controls for PPARγ1
[35] and PPARγ2 [18], respectively. The amplification protocol was as follows:
initial cycle for 3 minutes at 95°C, and 30 cycles each at
(95°C for 30 seconds, 55°C for 30 seconds, and 72°C
for 30 seconds) followed by a final extension cycle at 72°C for 7
minutes. PCR reactions were performed on a Robocycler (Stratagene Inc, La Jolla, Calif, USA)
and products were analyzed electrophoretically on 2% (w/v) agarose gels. The
intensity of the PCR bands was determined using Fluor-S multi-imaging
analysis system (Bio-Rad, Hercules, Calif, USA). Level of mRNA for PPARs was
normalized to the levels of S-15 housekeeping gene.Table 1
PCR primer sets, sequence, product size (bp), nucleotide (nt) location, and GenBank accession numbers for rat
PPARs used in this study. Note that a common antisense oligoprimer (sequence in bold) was used for PPARγ1a, PPARγ1b, and PPARγ2.
Product/Sense primerAntisense primerProduct sizentaccession#(bp)locationPPARα5-TTG TGA CTG GTC AAG CTC AGG ACA-35-TCG TAC GCC AGC TTT AGC CGA ATA-3492296–787NM013196PPARβ5-TAA CGC ACC CTT CAT CAT CCA CGA-35-TTG ACA GCA AAC TCG AAC TTG GGC-3390873–1262U40064PPARγ5-TCT CCA GCA TTT CTG CTC CAC ACT-35-ATA CAA ATG CTT TGC CAG GGC TCG-3533257–789NM013124PPARγ1a5-CTG ACG AGG TCT CTC TC G GCT G-35-AGC AAG GCA CTT CT GAA ACC GA-365821–679AF246458PPARγ1b5-CAG CGC TAA ATT CAT CTT AAC T-35-AGC AAG GCA CTT CTG AA A CCG A-361821–639AF246457PPARγ25-GAG CAT GGT GCC TTC GCT GA-35-AGC AAG GCA CTT CTG AA A CCG A-356337–600AB01956185–648AF15666686–649Y12882PPARγ/ERα(primers for real-time PCR were obtained from Superarray Inc)190/179[PPARγ/ERα]NM013124 [PPARγ](Sequence are not disclosed by the Company)respectivelyNM012689 [ERα]Gapdh5-ATG ATT CTA CCC ACG GCA AG-35-CTG GAA GAT GGT GAT G CGT T-38971–159DQ403053184–272BC087743216–304BC059110Rig/S15 (Ambion)5-TTC CGC AAG TTC ACC TAC C-35-CGG GGC CGG CCA TGC T TTA CG-336174–433BC105810Quantitative real-time PCR (Bio-Rad, MyiQTM) for determination of
expression levels of PPARγ and ERα mRNA was performed in 25-μL reaction
mixture containing 12.5 μL RT2 real-time
SYBR/Fluorescein Green PCR master mix, 1 μL
first strand cDNA, 1 μL RT2 validated PCR primer set for PPARγ
or ERα (Super Array Bioscience
Corporation, Frederic, Md, USA), and 10.5 μL
PCR-grade water (Ambion Inc). Samples were run in 96-well PCR plates (Bio-Rad,
Hercules, Calif, USA) in duplicates, and the results were normalized to
GAPDH (see primer set in Table 1) expression. The
amplification protocol was set at 95°C for 15 minutes for one cycle,
and 40 cycles each at (95°C for 30 seconds, 55°C for 30
seconds, and 72°C for 30 seconds) followed by melting curve determination between 55°C and 95°C to
ensure detection of a single PCR product. Template RNA from rat white adipose
tissue and penis were used for determination of amplification efficiencies for
(ERα/PPARγ) targets and GAPDH
by generating standard curves. Curves were generated by using serial 10-fold dilutions total RNA and plotting the log dilution against CT (threshold cycle) value obtained for each dilution. The
Pearson's correlation coefficient (r) value for each generated standard curve
was ≥0.98, and the calculated amplification efficiency was between 98.5 to 99%.


## 2.4. Immunohistochemistry (IHC)

Immunolocalization of PPARγ in penile tissue was performed using mouse anti-PPARγ
IgG1 monoclonal antibody (sc7273, Santa Cruz Biotechnology Inc, Santa Cruz, Calif, USA)
raised against a C-terminus sequence of human and mouse PPARγ  (similar to the corresponding rat sequence).
The antibody detects PPARγ1, PPARγ2 and, to a lesser extent, PPARα and PPARβ of rat, mouse, and
human by IHC using paraplast-embedded tissues. Approximately 5-mm-long penis sections from the middle
of the body of the penis were fixed in 4% paraformaldehyde
for 48 hours, embedded in Paraplast (Sigma-Aldrich), and cut at 5-μm thickness [7].
Mounted penis sections were deparaffinized in Hemo-D (Scientific Safety
Solvents, Keller, Tex, USA) and hydrated to distilled water (dH2O).
The slides were transferred to a rack and placed in 1 L of 10 mM sodium citrate
(pH 6.0). The beaker was placed on a hot plate, allowed to come to a boil and
tissues were boiled for 20 minutes. When the citrate solution cooled to near
room temperature (RT), the slides were transferred to a glass staining dish and
equilibrated in phosphate buffered saline (PBS) (Sigma-Aldrich, ST Louis, Miss, USA). After 20 minutes incubation in blocker (5%
normal goat serum, Sigma-Aldrich) and 2.5% BSA (Sigma) in PBS, slides were washed
briefly in PBS. Anti-PPARγ, diluted 1:20 in blocker, was applied and the
sections were left to incubate overnight at RT. Next day, slides were washed 3x in PBS, 3 minutes each, and tissues were
incubated with Alexa 488-conjugated goat antimouse IgG (Molecular Probes, Eugene, Ore, USA)
for 1 hour at RT. After washing two times in PBS, 3 minutes each, slides were
mounted with VectaShield (Vector Laboratories, Burlingame, Calif, USA),
and the coverslips were sealed. The sections were examined using a Nikon
TE2000E microscope and digital images were generated using an attached Retiga
EX CCD digital camera (Q Imaging, Burnaby, BC, Canada).
Penile tissue sections from all 28-day treated rats were examined. Representative
micrographs from different penile histological structures were shown for
untreated control rats, and for rats treated with DES or DES + ICI.


## 2.5.  Fat Staining

Histochemical demonstration of fat was performed as previously described [7]. Briefly, tissue sections from penile body, approximately 5 mm-long, were
fixed for 24 hours in 4% formaldehyde, followed by en bloc staining of fat for
8 hours with 1% osmium tetroxide dissolved in 2.5% potassium dichromate
solution. Specimens were then processed for paraplast embedding and cut at 5-μm thickness. Deparaffinized sections were examined for
black staining indicative of fat cells using light microscopy.


## 2.6. Statistical Analyses

Analysis of real-time PCR data for relative gene
expression level (fold change of target relative to control) was performed
using a modification of the
delta delta Ct method (ΔΔ CT) as described previously [36]. Statistical differences
between treatment groups were performed using Sigma Stat statistical software
(Jandel Scientific, Chicago, Ill, USA). Δ CT for real-time PCR data [37],
and intensity values (for semi-quantitative RT-PCR data) were subjected to
analyses of variance. Experimental groups with means significantly
different (P<.05) from controls were identified using Holm-Sidak
and Tukey tests. When data were not distributed normally, or heterogeneity of
variance was identified, analyses were performed on transformed or ranked data.


## 3. Results

### 3.1. Detection and Sequence Analysis of PPAR and ERα Transcripts in the Body of the Penis

Primer sets used in this study are shown in Table1. Transcripts for three PPAR
isoforms (α, β, and γ) were detected, albeit at different levels, in penile
tissue from normal control adult (120 days) rats (Figure 1, parts A1 and A2). Semiquantitative
RT-PCR analysis of PPARs indicated predominant expression of PPARγ
mRNA when compared with PPAR (α and β)
isoforms (Figure 1(B)). Sequence analysis and alignment with published sequence
data confirmed the identity of all three PPAR isoforms. Treatment with DES
induced over three-fold-increase (3.38) in ERα transcripts  in 28-day-old
rats compared to over two-fold-increase (2.5) in 120-day-old adult rats when
each age group was compared with its respective untreated controls (Figure 2).
Similarly, DES induced slightly over seven-fold-increase (7.1) in PPARγ  transcription level in
28-day-old rats compared with over six-fold-increase (6.8) in 120-day-old
adult rats (Figure 3). The upregulation of PPARγ expression by DES in
28-day-old rats was abrogated when rats were cotreated with DES and ICI 182,
780 (Figure 4). The differences in the transcriptional level of penile
ERα and PPARγ between the DES-treated rat groups (28 versus 120-day-old
rats) were not significantly different. Because of
the relatively high expression of penile PPARγ in the 28-day-old rats
subsequent studies for determination of splice variants and PPARγ protein expression were performed in the 28-day-old rats.(A1) and (A2) RT-PCR amplification of three PPARs (α, β, and γ) from the
body of the penis of three (1, 2, and 3) normal adult (120 days) control rats.
(A1) Shows coamplification of PPARs (α, β, and γ (upper bands) and S15 (small
ribosomal subunit protein as housekeeping gene, lower bands) in two
representative rats (1 and 2). PCR markers were included in lane 1. Expected band sizes for S-15, PPARα, PPARβ, and PPARγ were 361, 492, 390, and 533 bp, respectively. Identities
of amplicons were further confirmed by sequence analysis (see Section 2). Note
that the ampilicons for PPARβ and S15 in
lane 5 and 6 were overlapped (compare run for PPARα, PPARβ, and PPARγ without S15 shown for rat 3 in (A2). In all rats note the
predominant expression of PPARγ. 15 μL PCR products were loaded per each lane. (B) Graphic
representation of signal intensity for PPARs showing
predominant expression of PPARγ. Transcript levels were normalized to
the levels of S15 housekeeping gene. To calculate the intensity for PPARβ the mean intensity of S-15 in lanes
2, 3, 8, and 9 in Figure (A1) was subtracted from the combined intensity of PPARβ + S15 in lanes 5 and 6 to obtain the
intensity of penile PPARβ
for rat 1 and 2, respectively. *P<.05.Figure 2
Real-time PCR showing 3.38 and 2.5 fold increase in ERα mRNA in penile tissue of 28-day-old
(DES-28 d) and adult rats (DES-Adult) neonatally treated with DES,
respectively. Fold change was calculated relative to respective controls
(CONT-28 d and CONT-Adult). Data (n= 4-5) are expressed as mean ±SE. *P<.05.Figure 3
Real-time PCR showing 7.1 and 6.8 fold increase in PPARγ mRNA in penile tissue of 28-day-old
(DES-28 d) and adult rats (DES-Adult) neonatally treated with DES, respectively.
Fold change was calculated relative to respective controls (CONT-28 d and
CONT-Adult). Data (n= 4-5) are expressed as mean ±SE. *P<.05.Figure 4
Real-time PCR data showing attenuation of the effect of DES on PPARγ
mRNA by ER blocker ICI 182, 780 in 28-day-old rats treated neonatally with
either 25 μL of olive oil (CONT-28 d), oil containing DES (DES-28 d; 0.1 mg/kg
bw), or DES plus ICI (DES + ICI; 16.6 mg/kg). ICI treatment significantly
inhibited DES-induced PPARγ
mRNA [DES-28 d versus DES + ICI]. Comparison between control and DES treated rats
showed 7.1 fold increase in expression [CONT-28 d versus DES-28 d]. Letter [a] indicates no significant differences between
CONT-28 d and DES + ICI. Data (n=5) are expressed as mean ±SE. *P<.05.


### 3.2. Detection of PPARγ Splice Variants  and Real-time PCR Data

In order to determine which PPARγ splice variant is expressed in the body
of normal and DES-treated rats, primers (Table 1) were designed to amplify the
two known rat PPARγ1a and  PPARγ1b splice variants using conventional
end-point RT-PCR. Splice variant analyses revealed
expression of PPARγ1a in normal 28-day-old
rat penis. However, in addition to PPARγ1a, PPARγ1b
and PPARγ2 were newly induced by DES treatment (Figure 5).Figure 5
RT-PCR amplification of PPARγ (γ, γ1a, γ1b, and γ2) splice variants in the body of the penis (P) of control (lanes,
2–5) and DES-treated (lanes, 7–10) 28-day-old rats. Lanes 12–15 were
amplification products from RNA template obtained from rat liver (L)
(used as positive control for PPARγ1b). Lanes 16–19 were RNA template from rat white adipose
tissue (F) (used as positive control for PPARγ1a and PPARγ2). Note that only PPARγ and PPARγ1a were detected in (P) of
normal rats. In contrast, in DES-treated rats enhanced expression of all PPARγ splice variants can be noted. In
addition to PPARγ
and PPARγ1a expression (seen in normal rats),
PPARγ1b and PPARγ2 were induced by DES-treatment. As
expected, PPARγ1b and PPARγ2 were strongly expressed in (L)
and (F), respectively. S-15 (S, lane 20) is a housekeeping gene
amplified from (P) as control for
RT-PCR conditions. The expected amplicon
sizes for S, PPARγ1a, PPARγ1b, PPARγ2, and PPARγ are 361, 658, 618, 563, and 533 bp, respectively.


### 3.3. Immunohistochemistry and Fat Staining

Immunohistochemistry results revealed PPARγ protein localization in transitional epithelium of the
urethra, and the surrounding corpus spongiosum penis. It is also expressed in
stromal, endothelial, neuronal, and smooth muscular cells of the cavernous
sinuses located in the corpus cavernousm region of normal 28-day-old rat penis
(Figures 6(a) and 6(b)). Treatment with DES induced a strong staining intensity for PPARγ protein in the peripherally located nuclei of newly
induced adipocytes (Figure 6(a), Panel (c) with a magnified inset-box view in
C2). PPARγ immunostaining was markedly reduced by ICI 182,780 treatment
(Figure 6(b)). In unstained penile
sections from 28-day-old and adult DES-treated rats, the new adipocytes were
seen as empty spaces similar to fat cells and were specifically localized in
the corpus cavernosum region of the penis (Figure 7, panels (b) and (d)). In
addition, staining with 1% osmium tetroxide confirmed that the empty spaces
were cluster of fat cells (stained as black granules in Figure 7, panels (c) and
(e)). No fat cells were seen in penile sections from rats treated with DES + ICI
(Figure 7, panels (f) and (g)).(a) Representative immunohistochemical staining for PPARγ protein in the body of the penis of
28-day-old DES-treated (A), (B), and (C) and control untreated rats (D), (E), and (F). Note that PPARγ protein is expressed in DES-treated (DES-28 d) and normal
rats (CONT-28 d) with increased intensity and fat cells in DES-treated rats
(see panel (C)). Note expression in transitional epithelium of the penile
urethra (PU) and the surrounding
corpus spongiousm (CS) in (A) and (D) and in the endothelium
of blood vessels and smooth muscle cells in the dorsal artery (a) and vein (v), and in nerve fibers of the dorsal nerve (n) of the penis (B) and (E). Similar staining intensity can be seen in the endothelium and
smooth muscles of the vascular lacunae (Vl)
in the corpus cavernosum penis (CC) in control normal rats (F). Note
one contrasting difference is that the cavernous spaces in DES-treated rats in
panel (C) are replaced with fat
cells (Fc) that show increased
staining intensity in the cell nucleus located at cell periphery. Panels (C2) and (F2) show a closer view of area outlined by insert box. Control
sections (minus primary antibody) were in panels (G), (H), and (I). Scale
bar = 30 μm. (b) IHC staining for PPARγ protein was
significantly reduced by ICI 182, 780 treatment [compare staining in panels (J) and (M) with (L) and (N)]. Panel (P) is a negative control (minus
primary antibody). Scale bar 30 μm.
(a)(b)Micrograph sections from penile body of normal rat (a) and rats
treated neonatally with DES (b)–(e) or DES + ICI (f)-(g). Panel (a) was from a normal adult rat stained
with H and E for demonstration of normal histological structures of the penis
(a: dorsal artery; v: dorsal vein; CC: corpus cavernosum; CS: corpus
spongiosum; PU: penile urethra; TA: Tunica albuginea). Panels ((b), unstained) and ((c), stained for fat with 1% osmium tetroxide) were from a 28-day-old rat. Panels ((d), unstained) and ((e),
stained for fat with 1% osmium tetroxide and presented as a magnified view of
CC and CS regions) were from adult rat (120 days) treated neonatally with DES.
Note the empty appearing spaces of fat cells in CC regions in unstained
sections (panels (b) and (d)). In sections stained with 1% osmium tetroxide (to confirm presence of fat)
fat cells appear as black granules, *. Panels ((f),
unstained) and ((g), stained with 1%
osmium tetroxide) were from a 28-day-old rat treated neonatally with DES + ICI.
Note the absence of empty appearing fat cells and lack of fat staining in CC
region. Sections from these rats were used for immunolocalization of PPARγ in Figure 6
parts (a) & (b). Scale bars = 30 (E) and 200 μm in other panels.
(a)(b)(c)(d)(e)(f)(g)


## 3.1. Detection and Sequence Analysis of PPAR and ERα Transcripts in the Body of the Penis

Primer sets used in this study are shown in Table1. Transcripts for three PPAR
isoforms (α, β, and γ) were detected, albeit at different levels, in penile
tissue from normal control adult (120 days) rats (Figure 1, parts A1 and A2). Semiquantitative
RT-PCR analysis of PPARs indicated predominant expression of PPARγ
mRNA when compared with PPAR (α and β)
isoforms (Figure 1(B)). Sequence analysis and alignment with published sequence
data confirmed the identity of all three PPAR isoforms. Treatment with DES
induced over three-fold-increase (3.38) in ERα transcripts  in 28-day-old
rats compared to over two-fold-increase (2.5) in 120-day-old adult rats when
each age group was compared with its respective untreated controls (Figure 2).
Similarly, DES induced slightly over seven-fold-increase (7.1) in PPARγ  transcription level in
28-day-old rats compared with over six-fold-increase (6.8) in 120-day-old
adult rats (Figure 3). The upregulation of PPARγ expression by DES in
28-day-old rats was abrogated when rats were cotreated with DES and ICI 182,
780 (Figure 4). The differences in the transcriptional level of penile
ERα and PPARγ between the DES-treated rat groups (28 versus 120-day-old
rats) were not significantly different. Because of
the relatively high expression of penile PPARγ in the 28-day-old rats
subsequent studies for determination of splice variants and PPARγ protein expression were performed in the 28-day-old rats.(A1) and (A2) RT-PCR amplification of three PPARs (α, β, and γ) from the
body of the penis of three (1, 2, and 3) normal adult (120 days) control rats.
(A1) Shows coamplification of PPARs (α, β, and γ (upper bands) and S15 (small
ribosomal subunit protein as housekeeping gene, lower bands) in two
representative rats (1 and 2). PCR markers were included in lane 1. Expected band sizes for S-15, PPARα, PPARβ, and PPARγ were 361, 492, 390, and 533 bp, respectively. Identities
of amplicons were further confirmed by sequence analysis (see Section 2). Note
that the ampilicons for PPARβ and S15 in
lane 5 and 6 were overlapped (compare run for PPARα, PPARβ, and PPARγ without S15 shown for rat 3 in (A2). In all rats note the
predominant expression of PPARγ. 15 μL PCR products were loaded per each lane. (B) Graphic
representation of signal intensity for PPARs showing
predominant expression of PPARγ. Transcript levels were normalized to
the levels of S15 housekeeping gene. To calculate the intensity for PPARβ the mean intensity of S-15 in lanes
2, 3, 8, and 9 in Figure (A1) was subtracted from the combined intensity of PPARβ + S15 in lanes 5 and 6 to obtain the
intensity of penile PPARβ
for rat 1 and 2, respectively. *P<.05.Figure 2
Real-time PCR showing 3.38 and 2.5 fold increase in ERα mRNA in penile tissue of 28-day-old
(DES-28 d) and adult rats (DES-Adult) neonatally treated with DES,
respectively. Fold change was calculated relative to respective controls
(CONT-28 d and CONT-Adult). Data (n= 4-5) are expressed as mean ±SE. *P<.05.Figure 3
Real-time PCR showing 7.1 and 6.8 fold increase in PPARγ mRNA in penile tissue of 28-day-old
(DES-28 d) and adult rats (DES-Adult) neonatally treated with DES, respectively.
Fold change was calculated relative to respective controls (CONT-28 d and
CONT-Adult). Data (n= 4-5) are expressed as mean ±SE. *P<.05.Figure 4
Real-time PCR data showing attenuation of the effect of DES on PPARγ
mRNA by ER blocker ICI 182, 780 in 28-day-old rats treated neonatally with
either 25 μL of olive oil (CONT-28 d), oil containing DES (DES-28 d; 0.1 mg/kg
bw), or DES plus ICI (DES + ICI; 16.6 mg/kg). ICI treatment significantly
inhibited DES-induced PPARγ
mRNA [DES-28 d versus DES + ICI]. Comparison between control and DES treated rats
showed 7.1 fold increase in expression [CONT-28 d versus DES-28 d]. Letter [a] indicates no significant differences between
CONT-28 d and DES + ICI. Data (n=5) are expressed as mean ±SE. *P<.05.


## 3.2. Detection of PPARγ Splice Variants  and Real-time PCR Data

In order to determine which PPARγ splice variant is expressed in the body
of normal and DES-treated rats, primers (Table 1) were designed to amplify the
two known rat PPARγ1a and  PPARγ1b splice variants using conventional
end-point RT-PCR. Splice variant analyses revealed
expression of PPARγ1a in normal 28-day-old
rat penis. However, in addition to PPARγ1a, PPARγ1b
and PPARγ2 were newly induced by DES treatment (Figure 5).Figure 5
RT-PCR amplification of PPARγ (γ, γ1a, γ1b, and γ2) splice variants in the body of the penis (P) of control (lanes,
2–5) and DES-treated (lanes, 7–10) 28-day-old rats. Lanes 12–15 were
amplification products from RNA template obtained from rat liver (L)
(used as positive control for PPARγ1b). Lanes 16–19 were RNA template from rat white adipose
tissue (F) (used as positive control for PPARγ1a and PPARγ2). Note that only PPARγ and PPARγ1a were detected in (P) of
normal rats. In contrast, in DES-treated rats enhanced expression of all PPARγ splice variants can be noted. In
addition to PPARγ
and PPARγ1a expression (seen in normal rats),
PPARγ1b and PPARγ2 were induced by DES-treatment. As
expected, PPARγ1b and PPARγ2 were strongly expressed in (L)
and (F), respectively. S-15 (S, lane 20) is a housekeeping gene
amplified from (P) as control for
RT-PCR conditions. The expected amplicon
sizes for S, PPARγ1a, PPARγ1b, PPARγ2, and PPARγ are 361, 658, 618, 563, and 533 bp, respectively.


## 3.3. Immunohistochemistry and Fat Staining

Immunohistochemistry results revealed PPARγ protein localization in transitional epithelium of the
urethra, and the surrounding corpus spongiosum penis. It is also expressed in
stromal, endothelial, neuronal, and smooth muscular cells of the cavernous
sinuses located in the corpus cavernousm region of normal 28-day-old rat penis
(Figures 6(a) and 6(b)). Treatment with DES induced a strong staining intensity for PPARγ protein in the peripherally located nuclei of newly
induced adipocytes (Figure 6(a), Panel (c) with a magnified inset-box view in
C2). PPARγ immunostaining was markedly reduced by ICI 182,780 treatment
(Figure 6(b)). In unstained penile
sections from 28-day-old and adult DES-treated rats, the new adipocytes were
seen as empty spaces similar to fat cells and were specifically localized in
the corpus cavernosum region of the penis (Figure 7, panels (b) and (d)). In
addition, staining with 1% osmium tetroxide confirmed that the empty spaces
were cluster of fat cells (stained as black granules in Figure 7, panels (c) and
(e)). No fat cells were seen in penile sections from rats treated with DES + ICI
(Figure 7, panels (f) and (g)).(a) Representative immunohistochemical staining for PPARγ protein in the body of the penis of
28-day-old DES-treated (A), (B), and (C) and control untreated rats (D), (E), and (F). Note that PPARγ protein is expressed in DES-treated (DES-28 d) and normal
rats (CONT-28 d) with increased intensity and fat cells in DES-treated rats
(see panel (C)). Note expression in transitional epithelium of the penile
urethra (PU) and the surrounding
corpus spongiousm (CS) in (A) and (D) and in the endothelium
of blood vessels and smooth muscle cells in the dorsal artery (a) and vein (v), and in nerve fibers of the dorsal nerve (n) of the penis (B) and (E). Similar staining intensity can be seen in the endothelium and
smooth muscles of the vascular lacunae (Vl)
in the corpus cavernosum penis (CC) in control normal rats (F). Note
one contrasting difference is that the cavernous spaces in DES-treated rats in
panel (C) are replaced with fat
cells (Fc) that show increased
staining intensity in the cell nucleus located at cell periphery. Panels (C2) and (F2) show a closer view of area outlined by insert box. Control
sections (minus primary antibody) were in panels (G), (H), and (I). Scale
bar = 30 μm. (b) IHC staining for PPARγ protein was
significantly reduced by ICI 182, 780 treatment [compare staining in panels (J) and (M) with (L) and (N)]. Panel (P) is a negative control (minus
primary antibody). Scale bar 30 μm.
(a)(b)Micrograph sections from penile body of normal rat (a) and rats
treated neonatally with DES (b)–(e) or DES + ICI (f)-(g). Panel (a) was from a normal adult rat stained
with H and E for demonstration of normal histological structures of the penis
(a: dorsal artery; v: dorsal vein; CC: corpus cavernosum; CS: corpus
spongiosum; PU: penile urethra; TA: Tunica albuginea). Panels ((b), unstained) and ((c), stained for fat with 1% osmium tetroxide) were from a 28-day-old rat. Panels ((d), unstained) and ((e),
stained for fat with 1% osmium tetroxide and presented as a magnified view of
CC and CS regions) were from adult rat (120 days) treated neonatally with DES.
Note the empty appearing spaces of fat cells in CC regions in unstained
sections (panels (b) and (d)). In sections stained with 1% osmium tetroxide (to confirm presence of fat)
fat cells appear as black granules, *. Panels ((f),
unstained) and ((g), stained with 1%
osmium tetroxide) were from a 28-day-old rat treated neonatally with DES + ICI.
Note the absence of empty appearing fat cells and lack of fat staining in CC
region. Sections from these rats were used for immunolocalization of PPARγ in Figure 6
parts (a) & (b). Scale bars = 30 (E) and 200 μm in other panels.
(a)(b)(c)(d)(e)(f)(g)


## 4. Discussion

This study demonstrated that three PPAR transcripts (α, β, and γ) are constitutively coexpressed
in normal rat penis with PPARγ as the
predominant isotype. In addition, it established that some ERα synthetic ligands, such as DES, can
activate PPARγ subtypes when administered at early perinatal days. Further, upregulation
of ERα by DES was associated with a corresponding increase in PPARγ suggesting
a synergistic interaction between the two receptors.Previous studies that used in situ hybridization to determine the distribution of PPARs in rat tissues, including
reproductive organs, showed expression of PPARα and PPARβ in somatic (Sertoli and
Leydig) and in germ cells of the testis, but did not address expression of
these two receptors in penile tissue [23, 24].
The role of PPARα and PPARβ in the testis, however, remains unknown.
Detailed study addressing expression of PPARγ isotypes in penile tissue is also
lacking, with the exception of a study that showed limited penile PPARγ
expression in corporal smooth muscle cells [25].In this study, PPARγ and PPARγ1a were detected in normal rat penis. However, DES as ERα
activator distinctively induced expression of PPARγ1b and PPARγ2
splice variants that were not present in control untreated penile
tissue. The induction of splice variant PPARγ1b is in agreement with previous in vitro studies that demonstrated
activation of PPARγ1 by the endocrine disruptor monoethyl-hexyl-phthalate in C2C12 mouse skeletal muscle cell line [2],
and with MCF-7 breast cancer cells stimulated with E2, the natural ERα ligand [38]. Further, the induction of
PPARγ2 concurs with increased adipogenesis observed in the corpus cavernousm penis as PPARγ2 is considered a unique marker for mature
adipocytes, and its forced induction is associated with terminal
differentiation of preadipocytes or fibroblast cells to functional mature
adipocytes [11, 22].
The upregulation of PPARγ was abrogated by
coadministration of the type-II antiestrogen ICI 182,780, indicating that DES
effects were mediated, at least in part, via the estrogen receptor system. It
is possible, however, that ICI may have directly repressed activation of PPARγ as
ICI was previously shown to inhibit the action of the selective PPARγ
agonist BRL 48, 482 in MDA-MB 231 breast cancer cell culture in the absence of
ER [38].One important difference between this study and previous in vitro studies that addressed
signal cross-talk between PPARγ and
ERα using MCF-7 cells [38–40] is that the activation of ERα by DES
in our study is associated with selective induction of PPARγ1b and
PPARγ2. This unique effect resulted in generation of de novo adipocytes that provide
direct functional proof for PPARγ2 induction. In contrast to our study, activated ERα
by E2 lowers both basal and ligand-stimulated PPARγ-mediated gene reporter activity in MCF-7 cancer cell culture
[38]. Likewise, activation of PPARγ in MCF-7 cell culture with the natural
PPARγ ligand cyclopentenone 15-deoxy-Δ12,14 prostaglandin J2 (15d-PGJ2)
inhibited estrogen-responsive elements [40]. Consequently, the MCF-7 cell
culture studies suggest that ERα and PPARγ negatively regulate each other. The
reason for the difference between our study and the aforementioned in vitro data could be related to differences
between in vitro and in vivo milieu
or to the deletional mutants used in the in vitro studies compared with the in vivo wild type receptors in our
study. Another reason for the disagreement could be due to differences in
coactivators and corepressors present in MCF-7 and penile tissue cells or more
importantly to differences in the ligands used. One plausible hypothesis,
however, for the increased transcriptional activation of PPARγ1b and PPARγ2 by DES-activated ERα is that
exposure of rats to DES at a critical neonatal period of days 1 to 12 is
uniquely associated with reprogramming of penile stromal or preadipocytes to
mature adipocytes [5–8].
In support of this concept, it is known that postnatal days 1 to 5 in rodents
coincide with a period for reproductive tract and adipocyte differentiation [41]. Further, data from other laboratories indicated that
neonatal exposure of rodents to DES is associated with increased whole body fat
at adulthood [42]. This novel adipogenic
effect of DES was proposed as a model for the study of what is called
developmental obesity mediated by early exposure to endocrine
disruptors [43].The molecular mechanism involved in DES-ERα-PPARγ transactivation could be related to two
factors. First, activated-ERα could
directly bind to PPAR response elements (PPREs) because the two receptors share
the capacity to bind to the AGGTCA half-sites consensus sequences contained as
palindrome or direct repeat in estrogen response elements (EREs) and PPRE
sequences, respectively [44].
This mechanism could result in bidirectional activation of shared target
sequences between ERα
and PPARγ
depending on activated receptor involved. Second, it is known that estrogen
could induce enzymatic conversion of prostaglandin D2 (PGD2) and the endogenous
metabolites of the latter can directly activate PPARγ [45].
The latter effect, however, was not associated with induced PPARγ mRNA [46] suggesting that the first
mechanism could be in play in our study.The strong PPARγ protein expression in normal transitional
epithelium of the urethra and the dorsal artery and vein of the penis indicates
possible physiological role for PPARγ in the penis vasculature and the
urothelium of the urinary tract. Although this study did not address
functionality of PPARγ in the penis,
current evidence suggests that its constitutive expression in some tissues is
linked to eicosanoids and prostaglandins (PGs) actions [47, 48]. In this regard, the terminal metabolite of the J series of PG, 15d-PGJ2,
is considered the natural activator of PPARγ [48]. Sources of penile PGs could include
synthesis by local penile cells and/or
cells of the renal medulla where PGs can be transported via the ureter and
pelvic urethra to the penis [49]. Among other functions, PGs are important mediators of inflammation, vascular
homoeostasis, and pain all
of which may be relevant to the pathophysiology of the penis.Staining with osmium confirmed the
presence of new lipid-laden adipocytes in penile tissues of DES-treated rats.
Previously, our group showed that Sprague-Dawley rats treated neonatally with
DES accumulated fat in the corpus cavernous penis [5–8]
just as observed for the rats in the present study. The histological demonstration
of DES-induced lipid buildup in the corpus cavernosum penis concurs with the newly induced adipocyte marker PPARγ2 detected with
RT-PCR.In penile tissue direct pharmacological
activation of PPARγ by the
antidiabetic TZD pioglitazone reportedly blocked corporal veno-occlusive
dysfunction in rat model of type 2 diabetes mellitus [25].
However, this effect was associated with fat buildup suggesting that direct
activation of penile PPARγ by
TZDs or indirectly by ERα ligands, as in this study,
could be a potential pathway for development of undesirable adipogenesis.  In conclusion, PPARs are currently
considered potential drug targets for diverse conditions including, vascular
homoeostasis, diabetes mellitus, hyperlipidemia, inflammation, cancer, and
infertility [50–54].
This study furthers our knowledge of mechanisms of endocrine disruption
mediated by PPARγ in
male subjects. The ERα-PPARγ signal pathway activation by DES is
analogous in some way to mechanisms postulated for endocrine disruptor MEHP and
other phthalates esters and organotins which directly activates PPARγ and promotes
adipogenesis in cell culture models [2, 3, 55].



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*Source: 651419-2008-08-21.xml*