Expression of peroxisome proliferator-activated receptors in
urinary tract of rabbits and humans
Youfei
Guan1,
Yahua
Zhang1,
Linda
Davis1, and
Matthew D.
Breyer1,2,3
1 Division of Nephrology,
2 Departments of Medicine and
Molecular Physiology and Biophysics,
3 Veterans Affairs Medical Center,
and Vanderbilt University School of Medicine, Nashville, Tennessee
37212
 |
ABSTRACT |
Peroxisome
proliferator-activated receptors (PPARs,
,
/
, and
) are members of the nuclear receptor superfamily of
ligand-activated transcription factors. PPARs regulate the expression
of genes involved in lipid metabolism.
8(S)-hydroxyeicosatetraenoic acid (8-S-HETE), leukotriene
B4
(LTB4), and hypolipidemic
fibrates activate PPAR
, whereas PPAR
is activated by
prostaglandin metabolites. The present studies examined the intrarenal
and tissue distribution of rabbit and human PPAR
, -
/
, and -
mRNAs. Nuclease protection showed PPAR
predominated in liver, heart,
and kidney, whereas PPAR
, a putative adipose-specific transcription
factor, was in white adipose tissue, bladder, and ileum, followed by
kidney and spleen. Lower expression levels of PPAR
/
were observed
in several tissues. In situ hybridization of kidney showed PPAR
mRNA
predominated in proximal tubules and medullary thick ascending limbs of
both rabbit and human. PPAR
was exclusively expressed in medullary collecting duct and papillary urothelium. Immunoblot confirmed the
expression of PPAR
protein in freshly isolated inner medullary collecting ducts. mRNAs for all the PPARs were expressed in the ureter
and bladder in both rabbit and human, but PPAR
expression was
greatest. This distinct distribution of PPAR isoforms has important
implications for lipid-activated gene transcription in urinary
epithelia.
prostaglandins; collecting duct; ureter; bladder; nephron
 |
INTRODUCTION |
PEROXISOME PROLIFERATORS comprise a group of
structurally diverse compounds including hypolipidemic fibrates (e.g.,
clofibrate) and leukotriene analogs. When administered to rodents,
these compounds induce proliferation of peroxisomes and upregulate
several enzymes involved in lipid metabolism (18, 36). Peroxisome
proliferators are now known to bind to a family of nuclear receptors
designated peroxisome proliferator-activated receptors (PPARs). PPARs
were originally identified as members of the steroid hormone receptor superfamily of nuclear transcription factors which includes the thyroid
hormone receptors and retinoic acid receptors (41). PPARs form
heterodimers with the 9-cis retinoic
acid receptor, RXR
(25). These heterodimers bind to characteristic
DNA sequences termed peroxisome proliferator response elements (PPRE)
located in the 5'-flanking region of target genes (12, 30, 44,
47). After binding the PPREs, PPARs activate transcription of several genes including acyl-CoA synthase, acyl-CoA oxidase (44), cytochrome P-450 fatty acid
-hydroxylase (27,
30) and phosphoenolpyruvate carboxykinase (40).
Since the first PPAR (PPAR
) was cloned from mouse liver (18), two
additional PPAR genes have been recognized (8). These genes are
designated PPAR
(also referred to as PPAR
or NUC1) and PPAR
(19, 24). These PPARs are differentially activated by a variety of
fatty acids (19, 24, 45). Whereas PPAR
is activated by fibrates,
8(S)-hydroxyeicosatetraenoic acid
(8-S-HETE), and leukotriene
B4
(LTB4) (7, 45), PPAR
is
activated by 15-deoxy-
12,14-prostaglandin
J2
(15-deoxy
12,14-PGJ2),
a metabolite of PGD2 (12, 23).
Importantly, PPAR
is also activated by the antidiabetic
thiazolidinediones. A PPAR
/
-selective ligand has not yet been
identified. A growing body of evidence demonstrates that PPAR
,
-
/
, and -
are not only activated by different ligands but that
they are also expressed in distinct tissues. Whereas PPAR
is
expressed in liver, heart, brain, muscle, and kidney (1, 19, 24),
PPAR
has been relatively selectively expressed in adipose tissue
(42). Lower expression levels of PPAR
have also been reported in
other tissues (9). In the present studies, we examined the expression
of PPAR
, -
/
, and -
in rabbit tissues and determined their
distribution along the urinary tract in both rabbits and humans.
 |
MATERIALS AND METHODS |
Partial cloning of rabbit PPAR
,
-
/
, and -
.
Reverse transcription-polymerase chain reaction (RT-PCR) was used to
amplify a portion of rabbit PPAR
, -
/
, and -
from RNA
isolated from female New Zealand White rabbits (1.5-2.0 kg) as
described below. Primers were selected from conserved sequences in the
human, rat, mouse, and Xenopus
homologs. For rabbit PPAR
, a cDNA comprising a portion of the DNA
binding and ligand binding domains (D and E/F) (8) was obtained by
RT-PCR using liver RNA as a template and primers derived from human
PPAR
cDNA sequence (5' AGA ACT TCA ACA TGA ACA AGG TCA
3' for sense and 5' GCC AGG ACG ATC TCC ACA GCA AAT
3' for antisense) (28). A cDNA comprising a portion of the
transactivation and DNA binding (A/B and C) domains of rabbit
PPAR
/
was amplified from kidney cDNA using the primers derived
from mouse PPAR
/
(upstream primer, 5' CGG GAA GAG GAG AAA
GAG GAA GTG 3'; downstream primer, 5' CTT GTT GCG GTT CTT CTT CTG GAT 3') (24). For rabbit PPAR
, a pair of primers based on human homolog (sense, 5' CCC TCA TGG CAA TTG AAT GTC GTG
3'; and antisense, 5' TCG CAG GCT CTT TAG AAA CTC CCT
3') were used to amplify a cDNA sequence comprising part of the
DNA binding and ligand binding (C and E/F) domains (14). Total RNA was
purified from rabbit kidney and liver using Trizol-Reagent (GIBCO-BRL) and reverse transcribed to single-stranded cDNAs using Moloney murine
leukemia virus reverse transcriptase and 2.5 µM of random hexamers
according to manufacturer's instructions (GeneAmp RNA PCR kit;
Perkin-Elmer Cetus, Norwalk, CT). The cDNAs were then amplified using
PPAR-selective primers. PCR reactions were carried out in 10 mM
tris(hydroxymethyl)aminomethane (Tris) hydrochloride (pH 8.3), 50 mM
KCl, 2.5 mM MgCl2, 0.2 mM dNTPs,
and 1 µM primers at 94°C for 0.5 min, 58°C for 0.5 min, and 72°C for 0.5 min for 35 cycles in a thermal
cycler (model 9600, Perkin-Elmer Cetus). Amplified cDNAs were ligated
into pCR II vector (Invitrogen) and sequenced. Nucleotide and predicted
amino acid sequence were compared using the GenBank database and BLAST
and CLUSTAL programs at the National Institutes of Health data bank.
Preparation of human PPAR
, -
/
, and
-
probes. Three human PPAR cDNA fragments were
generated by RT-PCR using human liver and kidney total RNA (Clontech,
Palo Alto, CA) and human PPAR isoform-specific oligonucleotides, as
follows: 5' AGA ACT TCA ACA TGA ACA AGG TCA 3' (sense) and
5' GCC AGG ACG ATC TCC ACA GCA AAT 3' (antisense) for human
PPAR
; 5' AGC AGC CTC TTC CTC AAC GAC CAG 3' (sense) and
5' GGT CTC GGT TTC GGT CTT CTT GAT 3' (antisense) for
PPAR
/
; and 5' CCC TCA TGG CAA TTG AAT GTC GTG 3'
(sense) and 5' TCG CAG GCT CTT TAG AAA CTC CCT 3'
(antisense) for PPAR
. After amplification, a 524-bp PPAR
cDNA, a
471-bp PPAR
/
cDNA, and 761-bp PPAR
cDNA were sequenced and
subcloned in Bluescript SK(
) vector (Stratagene). Antisense and
sense probes were transcribed using appropriate RNA polymerases
(MAXIscript; Ambion, Austin, TX) and
35S-UTP as labeled isotope for in
situ hybridization.
Solution hybridization/ribonuclease protection
assays. Total RNA from various rabbit tissues was
isolated by using Trizol-Reagent (GIBCO-BRL). Briefly, 1 mg of tissue
sample was homogenized in 10 ml Trizol reagent, and 1/10 vol of
chloroform was added and vortex mixed for 15 s. The phases were
separated by centrifugation (12,000 g,
10 min), and isopropyl alcohol was added to the aqueous phase to
precipitate total RNA. The resulting RNA was dissolved in diethyl
pyrocarbonate-treated water.
Ribonuclease (RNase) protection assay was performed as described
earlier (3). Briefly, the plasmids [pBluescript SK(
), Stratagene] containing rabbit PPAR
(109 bp of
Xma I fragment), PPAR
/
(337 bp
of PCR fragment), and PPAR
(316 bp of
EcoR I fragment) inserts were
linearized with appropriate restriction enzymes. Radioactive riboprobes
were synthesized in vitro from 1 µg of linearized plasmids containing
three different cDNA fragments of PPAR isoforms by using MAXIscript kit
(Ambion) for 1 h at 37°C in a total volume of 20 µl. The reaction
buffer contained 10 mM dithiothreitol (DTT), 0.5 mM each of ATP, CTP,
and GTP, 2.5 µM of UTP, and 5 µl of 800 Ci/mmol
[
-32P]UTP at 10 mCi/ml (DuPont, NEN, Boston, MA). Hybridization buffer included 80%
deionized formamide, 100 mM sodium citrate, pH 6.4, and 1 mM EDTA (RPA
II, Ambion). Twenty micrograms of total RNA were incubated at 45°C
for 12 h in hybridization buffer with 5 × 104 cpm labeled riboprobes. After
hybridization, RNase digestion was carried out at 37°C for 30 min,
and precipitated protected fragments were separated on 4%
polyacrylamide gel at 200 V for 3 h. The gel was exposed to Kodak XAR-5
film overnight at
80°C with intensifying screens.
Medullary interstitial cell and cortical collecting
duct cell culture. Rabbit medullary interstitial cells
(RMICs) were cultured as previously described (15). Briefly, the left
kidney of a female New Zealand White rabbit (1.5-2.0 kg) was
removed. The medulla was dissected and minced in 5 ml of sterile RPMI
1640 plus 20% (vol/vol) fetal bovine serum (FBS, Hyclone). The
homogenate was injected subcutaneously in the abdominal wall. Twenty to
thirty days postsurgery, the subcutaneous renal medullary nodules were minced into 1-mm fragments and explanted in culture plates. Cells were
cultured in RPMI-1640 tissue culture medium supplemented with 25 mM
N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic
acid buffer, L-glutamine (2 mM),
20% (vol/vol) FBS, streptomycin, and penicillin. Cultures were
incubated at 37°C in 95%
O2-5%
CO2. Cells in their third to
fourth passage were generally used for experimentation.
Cortical collecting ducts. Primary
cultures of rabbit cortical collecting ducts (CCDs) were grown on
semipermeable supports (Transwell; Costar, Cambridge, MA) as previously
described (6). Briefly, two rabbit kidneys were perfused with
Krebs-Ringer. The renal cortex was separated from the capsule and
medulla via gross dissection and passed through a tissue press. The
dispersed tissue was digested with collagenase (0.1%),
deoxyribonuclease (100 U/ml), and soybean trypsin inhibitor (1,000 U/ml, 37°C) in Krebs-Ringer. This suspension was then poured over
plates precoated with monoclonal antibody specific for rabbit CCD
(3G10) and incubated for 10 min. Nonadherent cells were removed by
gentle aspiration. The adherent CCD cells were resuspended by sharp
mechanical blow and plated onto collagen-coated semipermeable supports
(Costar). Cells were grown to confluence in Dulbecco's modified
Eagle's medium with 1 µM aldosterone, 1%
penicillin-streptomycin-neomycin, supplemented with 10%
FBS at 37°C in 95% O2-5%
CO2.
Preparation of freshly isolated inner medullary
collecting duct cells. Inner medullary collecting ducts
(IMCDs) were isolated by a modification of a method described by Zeidel
et al. (46). Rabbits were killed, and kidneys were perfused free of
blood with 30 ml of ice-cold nonbicarbonate Ringer solution diluted 1:1
with Joklik minimum essential medium containing 10% FBS. Kidneys were then perfused with 5 ml of Joklik medium containing 0.2% collagenase. Inner medullas were excised, finely minced, and incubated with 0.2%
collagenase in Joklik medium for 90 min at 37°C in a shaking water
bath. The resulting mixed inner medullary cell suspension was
fractionated over 16% Ficoll layer in nonbicarbonate Ringer solution
by centrifugation for 45 min at 2,300 g. IMCDs were located at the top of
the 16% Ficoll layer. The cells were collected and washed twice with
Joklik medium supplemented with 10% FBS. Cell viability was measured
by trypan blue exclusion and assessed by phase-contrast microscopy.
Immunoblotting. Confluent RMICs and
CCDs and freshly isolated IMCDs were harvested in sodium dodecyl
sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer
(120 mM Tris · HCl, pH 6.5, 4% SDS, 5 mM DTT, and
20% glycerol). This material was then heated in boiling water for 3 min, and protein concentration was determined. Ten milligrams of
protein extract were loaded onto a 10% SDS-PAGE minigel and run at 100 V. Proteins were transferred to nitrocellulose membrane at 14 V
overnight at 4°C. The membrane was washed three times with
phosphate-buffered saline (PBS) and incubated in blocking buffer
(Tris-buffered saline which contained 150 mM NaCl, 50 mM Tris, 0.05%
Tween 20 detergent, and 5% Carnation nonfat dry milk, pH
7.5) for 1 h at 24°C, followed by three washings with blocking
buffer at 5-min intervals. The nitrocellulose membrane was then
incubated in the anti-PPAR
antibody (rabbit anti-mouse PPAR
1,2 polyclonal antibody; Biomol, Plymouth Meeting, PA) diluted 1:2,000 in blocking buffer for 2 h at room temperature. Following three
additional washings, the membrane was incubated with biotinylated anti-rabbit immunoglobulin G antibody (1:2,000; Vector, Burlingame, CA)
for 1 h, followed by three 15-min washings. Antibody labeling was
visualized by addition of chemiluminescence reagent (DuPont NEN) and
exposing the membrane to Kodak XAR-5film.
In situ hybridization. In situ
hybridization was performed as previously described (4). The human
kidney and ureter was from a male patient who died of a gunshot wound
and was deemed unsuitable for renal transplantation because of multiple
renal arteries. Human bladder tissue was from specimens
removed for bladder cancer, using regions which were microscopically
uninvolved. Briefly, prior to hybridization, human or rabbit kidney,
ureter, and bladder sections were deparaffinized, refixed in
paraformaldehyde, treated with proteinase K (20 µg/ml), washed with
PBS, refixed in 4% paraformaldehyde, and treated with triethanolamine
plus acetic anhydride (0.25% vol/vol). Finally, sections were
dehydrated with 100% ethanol.
35S-labeled antisense and sense
riboprobes from rabbit PPAR
(524 bp), PPAR
/
(337 bp), and
PPAR
(758 bp) and human PPAR
(524 bp), PPAR
/
(471 bp), and
PPAR
(761 bp) were hybridized to the section at 55°C for 18 h.
After hybridization, the sections were washed at 65°C once in
5× SSC (1× SSC is 0.15 M NaCl and 0.015 M sodium citrate,
pH 7.0) plus 10 mM
-mercaptoethanol (BME), then once in
50% formamide, 2× SSC, and 100 mM BME for 30 min. After an
additional two washes in 10 mM Tris, 5 mM EDTA, 500 mM NaCl (TEN) at
37°C, the sections were treated with RNase A (10 µg/ml) at
37°C for 30 min, followed by another wash in TEN at 37°C.
Sections were then washed twice in 2× SSC and twice in 0.1× SSC at 65°C. Slides were dehydrated with graded ethanol containing 300 mM ammonium acetate. Slides were then dipped in emulsion (K5; Ilford, Knutsford, Cheshire, UK) diluted 1:1 with 2% glycerol and
exposed for 4-5 days at 4°C. After developing in Kodak D-19, slides were counterstained with hematoxylin. Photomicrographs were
taken using a Zeiss Axioskop microscope and either dark-field (Micro
Video Instruments, Avon, MA) or bright-field optics.
Immunostaining. To define the
PPAR-positive nephron segments, in situ hybridization was followed by
immunostaining of tissue sections using a goat anti-human Tamm-Horsfall
antibody (Organon-Technica), which specifically recognizes medullary
and cortical thick ascending limb as well as the early portion of the
distal tubule (15, 46). Tissue sections were incubated with serial
dilutions of the Tamm-Horsfall antibody (1:1,000, 1:1,500, 1:2,000,
1:3,000, 1:3,500, and 1:5,000) as previously described (5).
Immunolabeling was detected using a biotinylated rabbit anti-goat
antibody followed by visualization with an avidin-biotin horseradish
peroxidase labeling kit (Vectastain ABC kit) and diaminobenzidine
staining.
 |
RESULTS |
Cloning and sequencing of rabbit
PPAR
, -
/
, and
-
cDNAs.
Sequencing of a 524-bp rabbit PPAR
fragment amplified by RT-PCR
revealed a predicted amino acid sequence that was 96.0% and 97.1%
identical to the human and mouse PPAR
, respectively (Fig. 1). The predicted amino acid sequences of a
337-bp rabbit PPAR
/
fragment demonstrated 92.9% and 87.5% amino
acid identity to human and mouse homologs. Finally, a 758-bp rabbit
PPAR
fragment was 96.4% and 98.0% identical to human and mouse
homologs at the amino acid level (Fig. 1). At the nucleotide level,
there was less than 42% identity between the cloned cDNA fragments of
rabbit PPAR
,
/
, and
(although these cDNAs fragments
represent different regions of the full-length PPAR).

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Fig. 1.
Schematic depiction of reverse transcription-polymerase chain reaction
(RT-PCR) amplified rabbit peroxisome proliferator-activated receptor
(PPAR) fragments mapped against functional domains (A-F) of the
PPAR. Functional domains (A-F) include C-DNA binding domain (DBD)
and E/F-ligand binding domain (LBD). Rabbit cDNA probes are indicated
by the black lines, and their size is given in base pairs (bp).
Identity of the rabbit PPAR , - / , and - with human homologs
is given on right. (These sequences
have been submitted to GenBank with accession numbers AF013264,
AF013265, and AF013266, respectively).
|
|
PPAR
,
-
/
, and -
are
differentially expressed in rabbit tissues.
The distribution of rabbit PPAR
, -
/
, and -
mRNA was
determined by nuclease protection (Fig. 2).
Comparable mRNA loading was confirmed by simultaneous nuclease
protection with a rabbit glyceraldehyde-3-phosphate dehydrogenase
(GAPDH) riboprobe. PPAR
expression was highest in liver, kidney, and
heart, followed by ileum, adrenal, and urinary bladder. Lower but
significant PPAR
message expression was observed in lung, stomach,
and brain. PPAR
was the only PPAR detected by nuclease protection in
white adipose tissue (data not shown). No PPAR
or -
/
were
observed in mRNA harvested from white adipose tissue. PPAR
mRNA expression in white adipose tissue was greater than levels
observed in any other tissue. Nuclease protection also demonstrated
high PPAR
expression in bladder and ileum (Fig. 2). Lower levels
were expressed in kidney, spleen, adrenal, heart, liver, lung, and
brain. Although PPAR
/
mRNA was widely distributed, expression
levels were generally much lower than for PPAR
and -
. PPAR
/
mRNA was not detected in significant amounts in the liver. Thus the
PPAR isoforms are differentially expressed in adult rabbit tissues.

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Fig. 2.
Left: nuclease protection showing
tissue distribution of rabbit PPAR , PPAR / , and PPAR mRNAs.
RNAs (30-50 µg) from tissue of 3-mo-old rabbits were analyzed
and quantitated as described under MATERIAL AND
METHODS. Right:
nuclease protection for the differential expression of rabbit PPAR ,
PPAR / , and PPAR in renal cortex versus medullary tissue. For
each assay, the equal RNA loading was assessed by simultaneous use a
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) riboprobe.
|
|
Intrarenal localization of PPAR
,
-
/
, and -
.
In situ hybridization was used to examine the distribution of the three
PPAR isoforms in the kidney, ureter, and bladder. A PPAR
antisense
riboprobe predominantly hybridized to tubules in the renal cortex and
outer medulla of both rabbit and human kidney (Fig.
3). Photomicrographs demonstrated intense
labeling of proximal tubules and distal nephron segments including
thick ascending limb. In contrast no labeling of glomeruli or
collecting duct was noted (Fig. 3). In renal outer medulla, PPAR
hybridized to thick ascending limb and the S3 segment of the proximal
straight tubule. No labeling of cortical thick ascending limbs,
collecting ducts, or any structures in the inner medulla was detected.

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Fig. 3.
In situ hybridization showing intrarenal distribution of PPAR in
rabbit and human.
A-D:
photomicrographs of PPAR distribution in rabbit kidney.
E and
F: human kidney. White grains show
areas of specific hybridization of PPAR .
A: ×50 rabbit renal cortex. Note
absence of labeling of glomeruli (g, and arrow heads).
B: ×100 rabbit cortex plus
counterstain with anti-Tamm Horsfall antibody (immunoreactivity shown
as brown reaction product) glomerulus (g) is unlabeled.
C: ×50 junction between rabbit
renal inner medulla (im) and outer medulla (om).
D: ×400 rabbit outer medulla
plus counterstain with anti-Tamm Horsfall antibody showing grains over
Tamm Horsfall-positive tubules. E:
×50 photomicrograph of PPAR riboprobe hybridization to human
kidney cortex. F: human renal medulla.
|
|
In contrast to PPAR
, no significant expression of PPAR
/
or
PPAR
was detected in the renal cortex; however, significant expression of PPAR
was detected in the IMCDs of both rabbit and human (Fig. 4). PPAR
mRNA was also
detected in the urothelium lining the renal papillae. PPAR
expression in the kidney was specific for the medullary collecting
duct, without significant detection in other portions in the kidney.
Furthermore, neither PPAR
nor -
/
were detected in the
medullary collecting duct. No hybridization of sense riboprobes for
PPAR
or either of the other two PPARs (PPAR
/
or PPAR
) was
observed (data not shown).

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Fig. 4.
In situ hybridization of PPAR riboprobe to rabbit and human kidney.
A: rabbit kidney papilla ×200
dark-field, note labeling of inner medullary collecting ducts (IMCDs; d
and arrowheads) and urothelium (u) lining the renal papillae.
B: human renal medulla dark-field
×50. White grains indicate areas of PPAR mRNA expression.
C: rabbit renal medulla ×400
bright-field, black grains indicate hybridization over papillary
collecting duct. D: human renal
medulla ×200 bright-field, dark-field. Lumen of a labeled
collecting duct (d). White grains show PPAR hybridization over
medullary collecting duct.
|
|
PPAR
protein is highly expressed in
IMCDs.
Immunoblots of protein extracts from RMICs, CCDs, and IMCDs using a
polyclonal anti-PPAR
antibody demonstrated that IMCDs highly
expressed PPAR
, with lower expression in cultured CCDs and RMICs
(Fig. 5). The presence of PPAR
protein
in IMCDs corresponds with in situ hybridization data.

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Fig. 5.
Detection of PPAR protein in rabbit medullary interstitial cells
(RMICs), cortical collecting duct (CCDs), and IMCDs. Immunoblotting was
performed as described in METHODS AND
MATERIALS and demonstrates abundant PPAR in freshly
isolated IMCDs (left lane) with less
immunoreactivity in cultured CCDs
(middle lane) and the least in
cultured renal medullary interstitial cells
(right lane).
|
|
PPAR expression in bladder and ureter.
PPAR
mRNA labeling was particularly intense in the transitional
urothelium of rabbit and human ureter and bladder (Figs.
6 and 7). The
expression of PPAR
was restricted to the epithelium, with no
expression detected in surrounding smooth muscle. PPAR
and
PPAR
/
were also detected in the urothelium of the ureter and
bladder of both species, albeit less intensely. Uroepithelial
expression of PPAR
was significantly lower than PPAR
, with
PPAR
/
expression appearing to be intermediate.

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Fig. 6.
In situ hybridization of PPAR
(A), PPAR /
(B), and PPAR
(C) showing mRNA distribution in
rabbit urinary bladder. (photomicrographs ×50). Dark-field
illumination shows expression of PPAR /
(D) and -
(E) isoforms in the human bladder.
F: ×400 bright-field
illumination of human bladder showing brown grains depicting
hybridization over transitional epithelium of human bladder .
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Fig. 7.
Ureteral expression of mRNAs for PPARs.
A: in human ureter, in situ
hybridization of PPAR / (dark-field ×50).
B: in human ureter, PPAR
(bright-field/dark-field, ×100).
C: in rabbit ureter in situ
hybridization of PPAR (×100 bright-field
illumination). White grains indicate hybridization.
D: ×400 bright-field of in situ
hybridization for PPAR over rabbit ureter brown-black grains
indicate hybridization. As in bladder, expression of both isoforms was
predominantly in the transitional epithelium.
|
|
 |
DISCUSSION |
The kidney is a major site of fatty acid, ketone body, and
prostaglandin metabolism (8, 15, 26), as well as prostaglandin synthesis (9). One major mechanism regulating lipid metabolism is the
transcriptional control of enzymes involved in oxidation of fatty acids
(10). The PPARs are ligand-activated transcription factors intimately
involved in the expression of several of these enzymes. We cloned cDNA
fragments of rabbit PPAR
, -
/
, and -
and mapped their
distribution in normal rabbit tissues and the urinary tracts of both
rabbit and human. The rabbit PPAR fragments cloned for these studies
are highly homologous to their human and murine counterparts. PPAR mRNA
species were highly expressed in kidney as well as other tissues more
classically identified with lipid metabolism, such as liver and adipose
tissue. The mRNA for these PPAR isoforms display distinct distribution
in normal tissues and within the kidney.
Recent evidence suggests PPAR
, -
/
, and -
complex with and
are activated by distinct endogenous lipid ligands (24,
45). Among the ligands demonstrated to activate PPAR-mediated gene transcription are arachidonate metabolites including
LTB4,
PGA1, PGA2,
PGJ2,
PGD2,
PGI2, and
8-S-HETE (7, 12, 23, 45). PPAR
is
uniquely activated by LTB4 and
8-S-HETE, whereas prostaglandins PGA2,
PGD2, and
PGJ2 uniquely activate PPAR
(12, 21, 23, 45). Upon binding their respective ligands, PPARs form
heterodimers with the retinoid X receptor (RXR). This complex binds to
a specific response element [peroxisome proliferator response
element (PPRE)] in target genes (33, 35, 41, 43, 48). The genes
downstream of these PPREs include enzymes implicated in regulation of
fatty acid metabolism, cholesterol metabolism, and adipogenesis (38). The present studies suggest unique intrarenal localization of the
PPARs, implying differential control of their activation along the
urinary tract.
As demonstrated in a previous report examining PPAR expression in rat
(1), the present results show that PPAR
is abundantly expressed in
tissues with high mitochondrial and
-oxidation activity including
liver, renal cortex, intestinal mucosa, and heart. This may correspond
with the demonstrated role of PPAR
in regulating genes encoding
mitochondrial and peroxisomal activities involved in the metabolism of
fatty acids. In situ hybridization demonstrates that PPAR
mRNA is
highly expressed in proximal tubules, with little labeling of glomeruli
or collecting ducts. PPAR
induces the expression of a variety of
genes in the rabbit including cytochrome P-450 4A6 (CYP4A6) (27).
CYP4A6 is an
-hydroxylase for arachidonate, laurate, and other fatty
acids (36). CYP4A6 has been shown have an upstream PPRE, and enzyme
expression is induced by fibrates via PPAR
(27).
Although the intrarenal expression of CYP4A6 in rabbit has not yet been
mapped, the cytochrome P-450 4A family is highly expressed in rat proximal tubule (2) and suggests PPAR
may
regulate fatty acid catabolism via induction of these enzymes in the
proximal tubule. There is less data on candidate genes that may be
activated by PPAR
in the medullary thick ascending limb, the other
major site of PPAR
mRNA expression along the nephron.
Of the three PPAR isoforms, the biochemical and physiological role of
PPAR
/
is least clearly defined. Although PPAR
/
was detected
by nuclease protection in rabbit kidney, no region-specific labeling
was observed by in situ hybridization. In contrast, significant expression of PPAR
/
mRNA in transitional epithelium of the
urinary bladder and ureter was detected, suggesting this PPAR may play a role in regulating gene expression in these tissues. To date, no
specific ligand that activates PPAR
/
has been identified. A
recent reported (20) showed that PPAR
/
can competitively inhibit
the activity of other PPARs either at the level of the PPRE or by
titrating out a limiting factor required for the transcription activity
of PPAR
(e.g., RXR
). Since PPAR
is also highly expressed in
the urothelium, such a mechanism may play a role in PPAR-
/
action
this tissue.
PPAR
is highly expressed in adipose tissue, but lower expression
levels have been previously reported in other tissues (9). After
binding a peroxisome proliferator, such as clofibrate, WY-14,643, 15-deoxy
12,14-PGJ2,
or thiazolidinediones, PPAR
activates adipogenesis, transforming fibroblasts into adipocytes (12, 23, 38). Importantly, the thiazolidinediones have recently been approved for use in the treatment
of diabetes mellitus (29) and have been shown to be particularly
high-affinity ligands for PPAR
(12, 23). The present studies
demonstrate for the first time that PPAR
is not only expressed in
adipose tissue but is also highly expressed in the distal urinary tract
both at the mRNA and protein level. In both rabbit and human kidney,
PPAR
mRNA was predominantly detected in IMCD. It is also highly
expressed in the transitional urothelium of ureter and bladder. This
contrasts with previous studies in rat where expression of PPAR
in
the renal medulla was not reported (1). PPAR
expression in urinary
bladder was not examined in those studies. Although the localization of
the PPAR mRNAs in rabbit and human kidney are consistent with each other, one must be cautious in interpreting their distribution in
humans, given the limited number of patients studied.
Little is known about the biological roles of PPAR
in collecting
duct, ureter, or bladder epithelium; however, its expression in these
tissues could have implications for renal effects of the antidiabetic
thiazolidinediones as well as in bladder carcinogenesis (32). It is
relevant to note that PPAR
is not only activated by prostaglandins
but that the medullary collecting is also the major site of
prostaglandin synthesis in the kidney (11, 37). Although the urothelium
has not been demonstrated to be a major site of prostaglandin
biosynthesis, prostanoid concentrations in the urine are in the
nanomolar range, well above those in plasma (9, 31). Importantly,
PGJ2 metabolites, proposed ligands for PPAR
, have been reported in human urine (16). Thus high endogenous prostaglandin concentrations in the urine could activate PPAR
in the medullary collecting duct, ureter, and bladder.
Diverse biological effects of the prostanoid ligands for PPAR
,
including PGA2,
PGD2, and
PGJ2, have been
described. These effects range from inhibition of cell cycle
progression and induction of apoptosis to suppression of viral
replication (13, 17, 22, 34). Although many of the biological effects
of the prostaglandins are mediated by G protein coupled,
membrane-spanning receptors (4, 39), the effects of prostaglandins on
cell proliferation appear to be mediated by nuclear binding
proteins (13, 26). PPARs have not been directly implicated as the
nuclear receptors mediating these effects of
PGA2,
PGD2, or
PGJ2 on cell growth and death;
however, these or other, similar, prostanoid-activated nuclear
transcription factors may be involved (22, 26). Roles for PPARs in
processes not directly involved in lipid metabolism remain to be
established. Even so, PPAR-mediated regulation of prostaglandin
metabolism might modify local concentrations of prostaglandins and
their effects on urinary epithelia.
In summary, we have cloned fragments of rabbit PPAR
, -
/
, and
-
and described their tissue distribution. Within the kidney and in
the lower urinary tract, there is distinct distribution of these PPAR
isoforms. In both human and rabbit kidney, PPAR
is predominantly
expressed in proximal tubules, with lower expression in medullary thick
ascending thick limbs. In contrast, PPAR
is predominantly located in
IMCDs. We have also demonstrated that PPAR
protein is expressed in
the IMCD. No distinct intrarenal localization of PPAR
/
was
observed. High levels of expression of PPAR
mRNA were detected in
the urothelium of ureter and bladder of both rabbit and human. Lower
but significant expression of PPAR
/
was also detected in the
urothelium. The physiological roles of these lipid-activated
transcription factors in urinary epithelia remain to be determined.
 |
ACKNOWLEDGEMENTS |
We thank Reyadh Redha for expert technical support. We also thank
Dr. Mitchell Lazar for providing us with a sample of the anti-PPAR
antibody.
 |
FOOTNOTES |
M. D. Breyer is a recipient of a Veterans Affairs Clinical Investigator
Award. Additional support for this project was provided by the George
M. O'Brien Kidney Center through National Institute of Diabetes and
Digestive and Kidney Diseases (NIDDK) Grant
2P50-DK-39261, a Veterans Affairs Merit Award, and NIDDK
Grants 2P01-DK-38226 and DK-37097 (to M. D. Breyer).
Address for reprint requests: M. D. Breyer, F-427 ACRE Bldg., Veterans
Affairs Medical Center, Nashville, TN 37212.
Received 21 February 1997; accepted in final form 28 August 1997.
 |
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