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Institut de Biologie Animale, Université de Lausanne, CH-1015 Lausanne, Switzerland
Address all correspondence and requests for reprints to: Dr. Beatrice Desvergne or Dr. Walter Wahli, Institut de Biologie Animale, Université de Lausanne, CH-1015 Lausanne, Switzerland. E-mail: beatrice.desvergne@iba.unil.ch or walter.wahli{at}iba.unil.ch
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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, whose
activity is markedly up-regulated in the liver and small intestine
during fasting. Finally, PPAR
2 mRNA levels are decreased by 50%
during fasting in both white and brown adipose tissue. In conclusion,
fasting can strongly influence PPAR expression, but in only a few
selected tissues. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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(NR1C1);
PPARß (NR1C2), which is also called PPAR
, NUC-1, or FAAR; and
PPAR (NR1C3), have been reported in vertebrates as primitive as
cyclostoma (2), in lower vertebrates (3), and
in mammals (4). In mammals, alternative promoter usage and
differential splicing of the ppar gene transcripts yield
two isoforms, 1 and 2 (5). In human, a PPAR3 mRNA
is transcribed from a third promoter, resulting in a protein identical
to PPAR1 (6).
PPAR heterodimerizes with the 9-cis RXR to bind DR1
(direct repeat spaced by 1 bp) response elements, called PPAR response
element, identified in the promoters of target genes involved in
microsomal 
-hydroxylation, peroxisomal ß-oxidation,
mitochondrial ß-oxidation, ketogenesis, lipoprotein metabolism,
fatty acid binding, and fatty acid transport (7). Various
ligands have been described for PPARs: natural fatty acids, especially
polyunsaturated ones; arachidonate derivatives; hypolipidemic fibrates;
and insulin-sensitizing thiazolidinediones (8).
The nature of these PPAR ligands together with the function of
identified target genes suggest that PPARs play a key role in lipid and
glucose metabolism and homeostasis. PPARs are expressed in a broad
range of tissues in adult rodents, as previously shown by in
situ hybridization (9). PPAR is expressed in cells
with high catabolic rates of fatty acids and high peroxisomedependent
activities (hepatocytes, cardiomyocytes, proximal tubules of kidney,
brown adipocytes, and intestinal mucosa) (4, 9).
PPAR-null mutant mice allowed identification of this isotype as a
mediator of the pleiotropic effects of peroxisome proliferators in
liver (10). This isotype is the best studied to date
during fasting. Elevated glucocorticoid hormone levels, observed during
fasting and stress, up-regulate hepatic PPAR expression at the
transcriptional level through binding of GRs to response elements
located in the promoter of PPAR (11, 12, 13, 14, 15). The
critical role of PPAR in lipid catabolism as well as glucose
homeostasis is revealed unequivocally during fasting, where
PPAR-null mutant mice show abnormally elevated FFA fasting levels.
This indicates a dramatic inhibition of hepatic fatty acid oxidation
that results in hepatic steatosis, myocardial lipid accumulation,
hypoketonemia, hypoglycemia, and hypothermia (16, 17).
PPARß is expressed ubiquitously and is abundant in most tissues
(4, 9, 18). PPARß (also often called PPAR) function
has been recently linked to basal lipid metabolism, embryo
implantation, colon cancer, and inflammation (19, 20, 21, 22).
PPAR expression is restricted mainly to adipose tissue and large
intestine, with some expression in parts of the immune system (4, 9). Several loss of function experiments in mice have shown that
PPAR is important in placental development and is required for
adipogenesis in vivo (23, 24, 25).
Previous in situ hybridization studies in adult rat have demonstrated that PPARs are expressed in specific cell populations or restricted areas of a given tissue (9). However, information concerning the relative expression level of each PPAR isotype with respect to the two others is presently lacking. We thus quantitatively studied PPAR mRNA expression for each of the three PPAR isotypes by ribonuclease (RNase) protection assay in 22 different organs of the adult rat in the fed state, during an overnight fast, and after refeeding. This provides valuable physiological information about the particular and sometimes opposite effects for which the three PPAR isotypes are responsible.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and rPPARß and total white adipose tissue
(WAT) RNA for cloning rPPAR1 and rPPAR2, with the following
downstream primers in the reverse transcriptase reaction:
5'-ATCTCAGGATCCATCAGTACATGTCTCTGT-3' (rPPAR),
5'-CTGCGTGGATCCTTAGTACATGTCCTTGTA-3' (rPPARß), and
5'-GGTGGGGAATTCCTGCTAATACAAGTCCTT-3' (rPPAR).
The following upstream primers were added for subsequent PCR
amplification: 5'-CCACAAGGATCCAACATGGTGGACACAGAG-3' (rPPAR),
5'-CAAGTGGGATCCGTCATGGAACAGCCACAG-3' (rPPARß),
5'-TTACTGGAATTCCCATGGTTGACACAGAGA-3' (rPPAR1), and
5'-TCTGTTGAATTCTGTTATGGGTGAAACTCT-3' (rPPAR2).
The PCR products were subcloned by using the Rapid DNA Ligation Kit
(Roche Molecular Biochemicals) into pCRII vector
(Invitrogen, San Diego, CA) resulting in pCR-rPPAR,
pCR-rPPARß, pCR-rPPAR1, and pCR-rPPAR2 recombinant vectors. Two
independent clones of each isotype and isoform were sequenced manually
on both strands using the T7 Polymerase Sequencing Kit with
[-35S]deoxy-CTP [Amersham Pharmacia Biotech (Arlington Heights, IL) and Hartmann Analytik
(Brawnschweig, Germany)].
Probes for RNase protection assay
A BamHI/EcoRI fragment was excised from a
partial L27 (large ribosomal subunit, 27-kDa protein) cDNA [155–374
nucleotides (nt)] (11) and subcloned into the
pBluescript KS- vector (Stratagene,
La Jolla, CA). The resulting construct pBS-L27 (150 nt) was digested by
EcoRI, transcribed by T7 polymerase (Promega Corp.) yielding a 200-bp riboprobe and a digested fragment of
150 bp. We used a previously described 287-bp probe located in the E/F
domain to assess rPPAR mRNA expression (11). The
plasmid pCR-rPPARß was digested by XbaI and
BglII and the 171-bp fragment containing the 3'-end of the
E/F domain subcloned into the pBluescript SK+
vector (Stratagene) digested by XbaI and
BamHI to obtain the recombinant plasmid pBS-rPPARß (171
nt). A 215-bp antisense probe was synthesized by T7 RNA polymerase
(Promega Corp., Madison, WI) transcription of pBS-rPPARß
(171 nt) linearized by XbaI, resulting in a protected
fragment of 171 bp for rPPARß. The plasmid pCR-rPPAR2 was digested
by PvuII and EcoRI, and the 351-bp fragment
containing the 5' of the A/B domain was subcloned into the pBluescript
KS+ vector (Stratagene) digested by
SmaI and EcoRI to obtain the recombinant plasmid
pBS-rPPAR (351 nt). A 409-bp antisense riboprobe was synthesized by
T7 RNA polymerase (Promega Corp.) transcription of this
plasmid linearized by HindIII, resulting in a protected
fragment of 351 bp for rPPAR2 and 257 bp for rPPAR1. The G+C
content of the probes is 51.2% for rPPAR, 60.2% for rPPARß,
51.4% for rPPAR1, and 50.4% for rPPAR2, with calculated
Tm of 73.0, 71.7, 72.8, and 73.4 C,
respectively.
To assess mRNA expression of mouse PPARs, we constructed the following
probes. A BamHI/KpnI fragment (nt 160–503 of
GenBank accession no. X57638) encoding the A/B domain was excised off a
pSG5-mouse (m)PPAR vector and subcloned into the
BamHI/KpnI sites of pBluescript
SK+ (Stratagene). The recombinant
plasmid pBSmPPAR (333 nt) was linearized by XbaI, and a
369-bp antisense riboprobe was transcribed by T7 RNA polymerase
(Promega Corp.), yielding a protected fragment of 333 bp.
A pSG5-mPPARß vector was digested by SmaI/XhoI,
and a fragment coding for partial A/B and C domains (nt 31–315 of
GenBank accession no. U10375) was subcloned into pBluescript
KS+ (Stratagene). The recombinant
plasmid pBSmPPARß (288 nt) was linearized by XbaI, and a
328-bp antisense riboprobe was transcribed by T3 RNA polymerase
(Promega Corp.), resulting in a protected fragment of 288
bp. A fragment spanning nt 910-1093 of GenBank accession no. U01841 and
encoding the partial D and E/F domain of mPPAR was cut off of a
pCMV-mPPAR vector and subcloned into the
HindIII/SmaI sites of pBluescript
KS+ vector (Stratagene). An
antisense riboprobe of 237 bp was transcribed by T7 RNA polymerase
(Promega Corp.) from the recombinant plasmid pBSmPPAR
(185 nt) linearized by HindIII, yielding a protected
fragment of 185 bp. To assess mouse L27 mRNA expression we used the rat
pBS-L27 (150 nt) probe described above.
For all PPAR probes, a 1:1 ratio of
[-32P]UTP (800 Ci/mmol) to cold UTP was
used, whereas a 1:20 ratio was used for the L27 probe. Labeled probes
were precipitated by trichloroacetic acid on GM filters
(Whatman, Clifton, NJ) to determine radioisotope
incorporation, which was typically between 60–70%. Filters were
counted using an automatic liquid scintillation system (Kontron Instruments Ltd., Zurich, Switzerland), and specific activities
were calculated. In pilot experiments, each probe was hybridized with
increasing amounts of sample RNA (5–40 µg), ensuring that 1 ng
specific PPAR probes and 10 ng L27 probe provided a molar excess of
probe vs. target RNA up to the maximal amount of RNA
tested.
RNA preparation and RNase protection assay
Eight- to 9-wk-old male Sprague Dawley (
200 g) rats were
purchased from RCC BRL (Basel, Switzerland). Eight- to 12-wk-old male
pure-bred wild-type or PPAR null mice on an Sv129 background were
obtained from Dr. Frank Gonzalez (10). Rats and mice were
housed with a 12-h light, 12-h dark cycle beginning at 0700 h and
free access to water and standard laboratory chow. For the fasting
experiment, food was removed at 1900 h. Access to water remained
free, and animals were killed the following day at 0700 h. In the
refeeding experiment, food was reintroduced after 7.5 h of
fasting. Dexamethasone was injected into rats at 40 µg/kg BW, and
rats were killed 4 h later. Animals were anesthetized with Forene
(Abbott Laboratories, Chicago, IL), killed by
decapitation, and rapidly dissected. Tissues were directly frozen in
TRIzol (Life Technologies, Inc., Grand Island, NY) by
liquid nitrogen and kept at -70 C until RNA preparation. Tissues were
homogenized with a Polytron homogenizer (Kinematica, Lucerne,
Switzerland), and total RNA was prepared with TRIzol following the
instructions of the provider. The RNase protection assay was carried
out as previously described (11). Approximately 15 µg
total RNA were hybridized to 1 ng specific PPAR probes each (2 x
109 cpm/µg) and 10 ng L27 probe (1 x
108 cpm/µg). RNase digestion [RNase A,
Sigma (St. Louis, MO); RNase T1, Life Technologies, Inc.] was carried out for all the probes at 30 C. Gels were
exposed on phosphor screens of a StormImager 840 (Molecular Dynamics, Inc., Sunnyvale, CA). IQuant 2.5 software
(Molecular Dynamics, Inc.) was used for quantification by
drawing a line per lane and removing background. PPAR mRNA expression
was normalized to the previously calculated specific activity of the
probe and to L27 mRNA expression. The PPAR/L27 ratio was further
normalized to the UTP content of each PPAR probe. SD values
were calculated with Excel software (Microsoft Corp., Redmond, VA) and Cricket Graph software (Computer
Associates, Islandia, NY) used for diagrams.
All animal experimentation was approved by the commission of the Etat de Vaud (Switzerland) for authorization of live animal experimentation.
Western blot analysis
Nuclear extracts of rat liver were prepared as follows. One gram
of rat liver was frozen in liquid nitrogen and crushed with a hammer.
Tissues were further homogenized in 5 ml of a 0.5 M
sucrose, 50 mM Tris-HCl (pH 7.5), 1 mM EDTA,
and 25 mM KCl solution in a glass-Teflon homogenizer. Cells
were then lysed with 0.5% Triton X-100 for 30 min at 4 C. The
homogenate was layered on a 0.9 M sucrose, 50
mM Tris-HCl (pH 7.5), 1 mM EDTA, and 25
mM KCl solution. After centrifugation at 1800 x
g for 20 min, pelleted nuclei were resuspended in 40%
glycerol, 50 mM Tris-HCl (pH 8), 5
mM MgCl2, and 0.1
mM EDTA and stored at -70 C. The protein
concentration was determined by measuring absorbance at 230 and 260 nm.
Thirty-five micrograms of proteins were separated by SDS-PAGE and
blotted on a BA85 Protrane membrane (Schleicher & Schuell, Inc., Keene, NH). PPARß antibody was raised against the first
15 N-terminal amino acids of mPPARß (peptide sequence:
MEQPQEETPEAREEEC). Blots were blocked for 1 h at 37 C with 5%
nonfat dry milk in 25 mM Tris-HCl (pH 8.0), 140
mM NaCl, 2 mM KCl, and
0.05% Tween 20 (NFDM TBS-Tween) and washed twice for 10 min each time
in TBS-Tween. Primary antibody was incubated for 4 h at a final
concentration of approximately 2 µg/ml in 0.5% NFDM TBS-Tween, and
blots were washed twice for 10 min each time in 0.5% NFDM TBS-Tween. A
secondary goat antirabbit IgG peroxidase-conjugated antibody was
incubated for 1 h at a dilution of 1:3000 in 0.5% NFDM TBS-Tween,
and blots were washed twice for 10 min each time in 0.5% NFDM
TBS-Tween and twice for 30 min each time in 5% NFDM TBS-Tween.
Immunodetection and chemiluminescence were performed according to the
manufacturer’s protocol (ECL, Amersham Pharmacia Biotech), and membranes were exposed on Kodak
films. Signals were quantified using an Elscript 400-AT/SM densitometer
(Hirschmann, Neuried, Germany).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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2 isoform, to have access to adequate
probes for further mRNA expression studies. The sequence of rPPAR is
identical to that reported previously (26). For rPPARß
we found an A at position 1028, yielding Asp343
as in all known other mammal PPARs, instead of the previously reported
T yielding Val343 (18). In the
rPPAR2 sequence we read CG at positions 330 and 331 yielding
Arg111 instead of GC yielding the reported
Ala111 (27, 28). As our sequence is
based on two independent RT-PCR reactions for both rPPAR1 and
rPPAR2, a polymorphism at this site in exon 2 of the
ppar gene may therefore exist.
PPAR expression in rat tissues
PPAR mRNA expression for each of the three PPAR isotypes was
quantified by RNase protection assay in 22 different organs of the
adult rat. All experiments were performed so as to allow a comparison
of the relative levels between the different tissues and between the
different isotypes (see Materials and Methods).
In nearly all tissues analyzed, PPARß is the most abundant isotype,
with its highest expression levels in the digestive tract, kidney,
heart, diaphragm, and esophagus (Figs. 1
and 2B). In comparison to the highest levels
in the digestive tract, PPARß expression is about 4–5 times lower in
all of the other organs analyzed. PPAR is highly expressed in brown
adipose tissue (BAT), at about 4 times the levels observed in liver,
heart, diaphragm, esophagus, and kidney (Figs. 1 and 2A). It is indeed
the highest PPAR level regardless of the isotype or tissue considered
(Fig. 2). With respect to its expression in liver, PPAR is expressed
at about 3-fold lower levels in small intestine, bladder (data not
shown), and skeletal muscle. In all other tissues, including WAT,
PPAR mRNA remains detectable, but at low levels. In the digestive
tract, PPAR mRNA expression decreases from the duodenum and ileum
toward the rectum. PPAR1 is highly expressed in both WAT and BAT at
levels comparable to those of PPAR mRNA in BAT and PPARß mRNA in
the digestive tract (Figs. 1 and 2C). PPAR1 is expressed in colon
and cecum at about 3-fold lower levels than in adipose tissue. PPAR1
expression in spleen, stomach, rectum, lung, and heart is about 10
times lower than that in adipose tissue and at least 25 times lower in
small intestine, testis, kidney, and liver. The presence of both
PPAR isoforms has been consistently found in lung, WAT, BAT, and
skeletal muscle, and the relative amount of PPAR2 mRNA
vs. total PPAR mRNA is, respectively, 33%, 30%, 18%,
and 32% (Figs. 1 and 2D).
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PPAR mRNA expression during fasting
We compared the mRNA expression of all three PPAR isotypes in
overnight-fasted male adult rats with that of their fed counterparts.
In an attempt to synchronize the feeding status, food was removed at
the end of the daylight hours, when rats do not feed extensively. The
rats were killed after a 12-h fast at the end of the dark cycle.
Strikingly, we observed a dramatic decrease in PPARß mRNA in the
liver and kidney by, respectively, 75% and 80% (Fig. 2B
). In none of
the other analyzed tissues did we observe any statistically significant
variations in PPARß mRNA levels (Fig. 2B). PPAR mRNA levels
increase in the liver during fasting as previously reported
(13) (Fig. 2A). We also observed a statistically
significant increase in PPAR expression in duodenum, jejunum, and
colon by 40%, 57%, and 20%, respectively, and a marked increase in
the ileum by 38%. A significant increase in PPAR expression was
also found in the thymus, where absolute PPAR expression is very low
(Fig. 2A). Interestingly, PPAR2 mRNA levels decreased in WAT and BAT
by up to 50% during fasting (Fig. 2D), whereas PPAR1 showed only a
weak regulation during fasting, with a decreased mRNA expression level
that was only significant in the ileum (Fig. 2C).
Nutritional regulation of PPARß mRNA expression in liver and
kidney
We then performed a refeeding experiment to study the potential
regulation of hepatic and renal PPARß mRNA expression by food intake.
The experiment started at the end of the daylight hours. The group of
refed rats was first fasted for 7.5 h and then had free access to
food for 4.5 h before mRNA levels were measured. Control groups in
this experiment comprised fed rats and rats fasted for 7.5 and 12
h (7.5 plus 4.5 h), respectively. We assessed PPAR and PPARß
mRNA expression by RNase protection assay in liver, kidney, heart, and
WAT (Fig. 3A). Again, we observed in both
liver and kidney a dramatic decrease in PPARß mRNA expression after
both 7.5 and 12 h of fasting by up to 70% (P <
0.05; Fig. 3B). Refeeding for 4.5 h after a 7.5-h fast was
sufficient to restore PPARß mRNA expression to the level observed in
fed animals (Fig. 3B). Concomitant to the decrease in mRNA levels,
PPARß protein abundance also decreased by about 30% during fasting,
as assessed by a semiquantitative Western blot on liver nuclear
extracts (Fig. 4).
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mRNA expression in liver, kidney, and heart (Fig. 3A).
We again observed a significant more than 2.5-fold increase in PPAR
mRNA gene expression during fasting in the liver (P <
0.05; Fig. 3C). Renal and cardiac PPAR mRNA expression levels were
not increased in a significant manner (P > 0.05; Fig. 3C).
Nutritional hepatic regulation of PPARß is independent of
PPAR
To further understand the opposite regulation of PPAR and
PPARß mRNA expression in liver during fasting, we used PPAR-null
mutant mice to investigate whether the observed tight nutritional
regulation of hepatic PPARß gene expression is dependent on PPAR
activity. The design of specific mouse PPAR probes allowed us to assess
PPARß mRNA expression in wild-type and mutant mice by RNase
protection assay (Fig. 5A). After an
overnight fast, we observed a similar decrease in hepatic PPARß mRNA
expression in wild-type and PPAR null-mutant mice by up to 80%
(Fig. 5B). Thus, nutritional regulation of hepatic PPARß mRNA
expression by fasting is independent of the activity of PPAR. A
parallel analysis of PPAR mRNA expression by RNase protection showed
similar expression levels of this latter subtype in both wild-type and
mutant mice, thus excluding compensatory effects of the PPAR isotype
in the absence of PPAR (data not shown).
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(13), the regulation of
hepatic PPARß mRNA expression levels by fasting and refeeding is
independent of plasma glucocorticoid levels, as injection of the
synthetic glucocorticoid dexamethasone had no effect on hepatic PPARß
mRNA expression (data not shown). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The predominant expression of PPARß in intestine also suggests a
potential role for this isotype in processes linked to food intake and
lipid metabolism. The mucosa of the small intestine resorbs diet lipids
that enterocytes transport via the cytosolic fatty acid-binding
proteins toward the smooth endoplasmic reticulum to resynthesize lipids
and triglycerides (30). In stomach, where no resorption of
nutrients occurs, PPARß expression is about 2 times lower than that
in intestine. A recent report using the PPARß-selective ligand GW2433
in PPAR-null mutant mice shows that the high levels of PPARß
present in the small intestine are sufficient to
trans-activate the liver fabp gene, which is a
PPAR target gene in the liver (31). PPARß may
therefore regulate the intestinal expression of other PPAR target genes
implicated in lipid metabolism for which a PPAR-dependent
trans-activation has been previously observed in liver.
The potential role of PPARß in metabolism is further underscored by
our observation that tissues with high metabolic rate, such as liver,
kidney, and oxidative muscles, also express PPARß mRNA at high
levels. Among muscular tissues, we observe a concomitant decrease in
PPAR and PPARß expression with decreasing importance of fatty acid
oxidation, i.e. skeletal muscle and bladder show lower
expression levels of both PPAR and PPARß compared to heart and
diaphragm. Taken together, these observations suggest that in tissues
where fatty acid oxidation is used for energy production, a link
between PPARß and the necessary high lipid metabolism may exist.
Fasting specifically up-regulates hepatic and intestinal PPAR
expression
Among all tissues analyzed, only the liver and small intestine
show increased PPAR mRNA expression during fasting. The high fasting
PPAR mRNA levels observed in the liver underline the prominent role
of this isotype in hepatic lipid metabolism. Up-regulation of hepatic
PPAR expression and subsequent activation of the PPAR target
genes implicated in hepatic fatty acid oxidation (i.e.
mitochondrial ß-oxidation, microsomal -hydroxylation, and
peroxisomal ß-oxidation) are critical events in the adaptive response
to fasting (16, 17, 32). In addition to the liver, PPAR
induces cardiac fatty acid oxidation during fasting (17).
Interestingly, we observed no increase in PPAR mRNA expression
during fasting in this oxidative muscle. In kidney also, PPAR mRNA
expression is not markedly increased, although fatty acids are
efficiently oxidized in the renal cortex. These results are in line
with the observation that clofibrate-fed rats show an increase in
cortical expression of mitochondrial and peroxisomal PPAR target
genes, whereas no significant change in PPAR mRNA itself is observed
(33).
Interestingly, GR mRNA levels are comparable in liver, kidney, and
heart (34), but corticosteroids do not modulate its
expression in the two latter tissues (35). Additional
experiments will be necessary to determine whether the GRs present in
the intestinal epithelium (36) mediate the observed
increase in intestinal PPAR mRNA expression.
Nutritional regulation of PPARß mRNA expression
An important finding of this study is the dramatic decrease in
hepatic and renal PPARß mRNA expression during fasting, which is
rapidly restored to control levels by refeeding. In agreement with a
regulation of PPARß by food intake and in addition to our results, a
recent report shows that PPARß mRNA expression levels remain low
after a 72-h fast (37). We observe a general tendency
toward decreased PPARß mRNA levels during fasting in all analyzed
tissues, but the tight regulation by feeding status in liver and kidney
suggests a tissue-specific mechanism of PPARß regulation. Our results
show that PPARß regulation during fasting is independent of plasma
glucocorticoid hormone levels and PPAR activity. Also, the
abnormally elevated plasma FFA concentrations observed in the
PPAR-null mutant mice (16) do not alter the expression
of PPARß compared with that in wild-type mice. Thus, serum lipid
levels alone are not sufficient for the nutritional regulation of
PPARß. The observation that PPARß is down-regulated in both the
liver and kidney, but PPAR is up-regulated only in the liver,
further suggests different regulatory mechanisms involved in the
expression of these two PPAR isotypes during fasting.
The similar regulation of PPARß expression in the liver and kidney may be related to metabolic fates common to these two tissues, e.g. gluconeogenesis and lipogenesis. As gluconeogenesis is increased during fasting, PPARß would be a negative regulator of this pathway. Interestingly, sterol regulatory element-binding protein-1c/adipocyte determination differentiation-dependent factor-1 shows a similar expression pattern as PPARß during fasting and refeeding (38, 39). This transcription factor is induced by food intake and mediates the expression of lipogenic genes induced by glucose and insulin (40). Further studies will be necessary to determine whether PPARß is implicated in sensing serum carbohydrate levels and is targeted by hormonal signaling leading to lipogenesis.
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Acknowledgments |
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null mutant mice; Daniel Bachmann,
Mai Perroud, and Nathalie Deriaz for expert technical help; and Michael
Man for excellent advice in statistical analysis. pSG5-mPPAR was a
kind gift of S. Green, pSG5-mPPARß of P. Grimaldi and pCMV-mPPAR
of R. Evans. |
Footnotes |
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2 and AJ306400 for rat
PPARß.
1 Present address: Pfizer Global Research and Development, Ann Arbor Laboratories, Molecular Sciences, 2800
Plymouth Road, Ann Arbor, Michigan 48105. 
2 Present address: Laboratoire de Chimie Clinique, Center Hospitalier
Universitaire Vaudois, 1011 Lausanne, Switzerland.
3 Present address: Department of Pharmacy and Pharmacology,
University of Bath, Claverton Down, Bath, United Kingdom BA2 7AY.
Abbreviations: BAT, Brown adipose tissue; NFDM TBS-Tween, 5% nonfat dry milk in 25 mM Tris-HCl (pH 8.0), 140 mM NaCl, 2 mM KCl, and 0.05% Tween 20; nt, nucleotides; RNase, ribonuclease; WAT, white adipose tissue.
Received February 26, 2001.
Accepted for publication July 6, 2001.
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 W. Shan, P. S. Palkar, I. A. Murray, E. I. McDevitt, M. J. Kennett, B. H. Kang, H. C. Isom, G. H. Perdew, F. J. Gonzalez, and J. M. Peters Ligand Activation of Peroxisome Proliferator-Activated Receptor {beta}/{delta} (PPAR{beta}/{delta}) Attenuates Carbon Tetrachloride Hepatotoxicity by Downregulating Proinflammatory Gene Expression Toxicol. Sci., October 1, 2008; 105(2): 418 - 428. [Abstract] [Full Text] [PDF]
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 B.-C. Yu, C.-K. Chang, H.-Y. Ou, K.-C. Cheng, and J.-T. Cheng Decrease of peroxisome proliferator-activated receptor delta expression in cardiomyopathy of streptozotocin-induced diabetic rats Cardiovasc Res, October 1, 2008; 80(1): 78 - 87. [Abstract] [Full Text] [PDF]
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 N. Petrovic, I. G. Shabalina, J. A. Timmons, B. Cannon, and J. Nedergaard Thermogenically competent nonadrenergic recruitment in brown preadipocytes by a PPAR{gamma} agonist Am J Physiol Endocrinol Metab, August 1, 2008; 295(2): E287 - E296. [Abstract] [Full Text] [PDF]
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 T.-L. Yue, S. S. Nerurkar, W. Bao, B. M. Jucker, L. Sarov-Blat, K. Steplewski, E. H. Ohlstein, and R. N. Willette In Vivo Activation of Peroxisome Proliferator-Activated Receptor-{delta} Protects the Heart from Ischemia/Reperfusion Injury in Zucker Fatty Rats J. Pharmacol. Exp. Ther., May 1, 2008; 325(2): 466 - 474. [Abstract] [Full Text] [PDF]
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 A. Koch, B. Konig, G. I. Stangl, and K. Eder PPAR{alpha} Mediates Transcriptional Upregulation of Novel Organic Cation Transporters-2 and -3 and Enzymes Involved in Hepatic Carnitine Synthesis Experimental Biology and Medicine, March 1, 2008; 233(3): 356 - 365. [Abstract] [Full Text] [PDF]
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 T. Mori, H. Kondo, T. Hase, I. Tokimitsu, and T. Murase Dietary Fish Oil Upregulates Intestinal Lipid Metabolism and Reduces Body Weight Gain in C57BL/6J Mice J. Nutr., December 1, 2007; 137(12): 2629 - 2634. [Abstract] [Full Text] [PDF]
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 Pieter de Lange, M. Moreno, E. Silvestri, A. Lombardi, F. Goglia, and A. Lanni Fuel economy in food-deprived skeletal muscle: signaling pathways and regulatory mechanisms FASEB J, November 1, 2007; 21(13): 3431 - 3441. [Abstract] [Full Text] [PDF]
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A. Koch, B. Konig, J. Spielmann, A. Leitner, G. I. Stangl, and K. Eder Thermally Oxidized Oil Increases the Expression of Insulin-Induced Genes and Inhibits Activation of Sterol Regulatory Element-Binding Protein-2 in Rat Liver J. Nutr., September 1, 2007; 137(9): 2018 - 2023. [Abstract] [Full Text] [PDF]
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C. R. Bush, J. M. Havens, B. M. Necela, W. Su, L. Chen, M. Yanagisawa, P. Z. Anastasiadis, R. Guerra, B. A. Luxon, and E. A. Thompson Functional Genomic Analysis Reveals Cross-talk between Peroxisome Proliferator-activated Receptor {gamma} and Calcium Signaling in Human Colorectal Cancer Cells J. Biol. Chem., August 10, 2007; 282(32): 23387 - 23401. [Abstract] [Full Text] [PDF]
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 I. G. Lunde, M. Ekmark, Z. A. Rana, A. Buonanno, and K. Gundersen PPAR{delta} expression is influenced by muscle activity and induces slow muscle properties in adult rat muscles after somatic gene transfer J. Physiol., August 1, 2007; 582(3): 1277 - 1287. [Abstract] [Full Text] [PDF]
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 M. Bunger, H. M. van den Bosch, J. van der Meijde, S. Kersten, G. J. E. J. Hooiveld, and M. Muller Genome-wide analysis of PPAR{alpha} activation in murine small intestine Physiol Genomics, July 18, 2007; 30(2): 192 - 204. [Abstract] [Full Text] [PDF]
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 M. J Leaver, M T. Ezaz, S. Fontagne, D. R Tocher, E. Boukouvala, and G. Krey Multiple peroxisome proliferator-activated receptor {beta} subtypes from Atlantic salmon (Salmo salar) J. Mol. Endocrinol., March 1, 2007; 38(3): 391 - 400. [Abstract] [Full Text] [PDF]
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 T. Saether, T. N Tran, H. Rootwelt, H. J Grav, B. O Christophersen, and T. B Haugen Essential fatty acid deficiency induces fatty acid desaturase expression in rat epididymis, but not in testis Reproduction, February 1, 2007; 133(2): 467 - 477. [Abstract] [Full Text] [PDF]
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M. A. Lomax, F. Sadiq, G. Karamanlidis, A. Karamitri, P. Trayhurn, and D. G. Hazlerigg Ontogenic Loss of Brown Adipose Tissue Sensitivity to {beta}-Adrenergic Stimulation in the Ovine Endocrinology, January 1, 2007; 148(1): 461 - 468. [Abstract] [Full Text] [PDF]
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 M. A. Dominick, M. R. White, T. P. Sanderson, T. van Vleet, S. M. Cohen, L. E. Arnold, M. Cano, S. Tannehill-Gregg, J. D. Moehlenkamp, C. R. Waites, et al. Urothelial Carcinogenesis in the Urinary Bladder of Male Rats Treated with Muraglitazar, a PPAR{alpha}/{gamma} Agonist: Evidence for Urolithiasis as the Inciting Event in the Mode of Action Toxicol Pathol, December 1, 2006; 34(7): 903 - 920. [Abstract] [Full Text] [PDF]
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P. de Lange, P. Farina, M. Moreno, M. Ragni, A. Lombardi, E. Silvestri, L. Burrone, A. Lanni, and F. Goglia Sequential changes in the signal transduction responses of skeletal muscle following food deprivation FASEB J, December 1, 2006; 20(14): 2579 - 2581. [Abstract] [Full Text] [PDF]
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 J. Shimakura, T. Terada, H. Saito, T. Katsura, and K.-i. Inui Induction of intestinal peptide transporter 1 expression during fasting is mediated via peroxisome proliferator-activated receptor {alpha} Am J Physiol Gastrointest Liver Physiol, November 1, 2006; 291(5): G851 - G856. [Abstract] [Full Text] [PDF]
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 A. J. Gilde, J.-C. Fruchart, and B. Staels Peroxisome Proliferator-Activated Receptors at the Crossroads of Obesity, Diabetes, and Cardiovascular Disease J. Am. Coll. Cardiol., October 27, 2006; 48(9_Suppl_A): A24 - A32. [Abstract] [Full Text] [PDF]
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 H. Boulanger, R. Mansouri, J. F. Gautier, and D. Glotz Are peroxisome proliferator-activated receptors new therapeutic targets in diabetic and non-diabetic nephropathies? Nephrol. Dial. Transplant., October 1, 2006; 21(10): 2696 - 2702. [Full Text] [PDF]
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Q. Yang, R. Kurotani, A. Yamada, S. Kimura, and F. J. Gonzalez Peroxisome Proliferator-Activated Receptor {alpha} Activation during Pregnancy Severely Impairs Mammary Lobuloalveolar Development in Mice Endocrinology, October 1, 2006; 147(10): 4772 - 4780. [Abstract] [Full Text] [PDF]
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 M. J. Yoon, G. Y. Lee, J.-J. Chung, Y. H. Ahn, S. H. Hong, and J. B. Kim Adiponectin Increases Fatty Acid Oxidation in Skeletal Muscle Cells by Sequential Activation of AMP-Activated Protein Kinase, p38 Mitogen-Activated Protein Kinase, and Peroxisome Proliferator-Activated Receptor {alpha} Diabetes, September 1, 2006; 55(9): 2562 - 2570. [Abstract] [Full Text] [PDF]
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K. Tsintzas, K. Jewell, M. Kamran, D. Laithwaite, T. Boonsong, J. Littlewood, I. Macdonald, and A. Bennett Differential regulation of metabolic genes in skeletal muscle during starvation and refeeding in humans J. Physiol., August 15, 2006; 575(1): 291 - 303. [Abstract] [Full Text] [PDF]
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 M Adachi, R Kurotani, K Morimura, Y Shah, M Sanford, B B Madison, D L Gumucio, H E Marin, J M Peters, H A Young, et al. Peroxisome proliferator activated receptor {gamma} in colonic epithelial cells protects against experimental inflammatory bowel disease Gut, August 1, 2006; 55(8): 1104 - 1113. [Abstract] [Full Text] [PDF]
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 J. Lopez-Soriano, C. Chiellini, M. Maffei, P. A. Grimaldi, and J. M. Argiles Roles of Skeletal Muscle and Peroxisome Proliferator-Activated Receptors in the Development and Treatment of Obesity Endocr. Rev., May 1, 2006; 27(3): 318 - 329. [Abstract] [Full Text] [PDF]
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 B. Desvergne, L. Michalik, and W. Wahli Transcriptional Regulation of Metabolism Physiol Rev, April 1, 2006; 86(2): 465 - 514. [Abstract] [Full Text] [PDF]
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D. Patsouris, J. K. Reddy, M. Muller, and S. Kersten Peroxisome Proliferator-Activated Receptor {alpha} Mediates the Effects of High-Fat Diet on Hepatic Gene Expression Endocrinology, March 1, 2006; 147(3): 1508 - 1516. [Abstract] [Full Text] [PDF]
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 F. Blaschke, Y. Takata, E. Caglayan, R. E. Law, and W. A. Hsueh Obesity, Peroxisome Proliferator-Activated Receptor, and Atherosclerosis in Type 2 Diabetes Arterioscler. Thromb. Vasc. Biol., January 1, 2006; 26(1): 28 - 40. [Abstract] [Full Text] [PDF]
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 J. C. Corton and H. M. Brown-Borg Peroxisome Proliferator-Activated Receptor {gamma} Coactivator 1 in Caloric Restriction and Other Models of Longevity J. Gerontol. A Biol. Sci. Med. Sci., December 1, 2005; 60(12): 1494 - 1509. [Abstract] [Full Text] [PDF]
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M. J. Leaver, E. Boukouvala, E. Antonopoulou, A. Diez, L. Favre-Krey, M. T. Ezaz, J. M. Bautista, D. R. Tocher, and G. Krey Three Peroxisome Proliferator-Activated Receptor Isotypes from Each of Two Species of Marine Fish Endocrinology, July 1, 2005; 146(7): 3150 - 3162. [Abstract] [Full Text] [PDF]
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G. Medina-Gomez, S. Virtue, C. Lelliott, R. Boiani, M. Campbell, C. Christodoulides, C. Perrin, M. Jimenez-Linan, M. Blount, J. Dixon, et al. The Link Between Nutritional Status and Insulin Sensitivity Is Dependent on the Adipocyte-Specific Peroxisome Proliferator-Activated Receptor-{gamma}2 Isoform Diabetes, June 1, 2005; 54(6): 1706 - 1716. [Abstract] [Full Text] [PDF]
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T. C. McCarthy, P. T. Pollak, E. A. Hanniman, and C. J. Sinal Disruption of Hepatic Lipid Homeostasis in Mice after Amiodarone Treatment Is Associated with Peroxisome Proliferator-Activated Receptor-{alpha}Target Gene Activation J. Pharmacol. Exp. Ther., December 1, 2004; 311(3): 864 - 873. [Abstract] [Full Text] [PDF]
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 J. M. Huss and D. P. Kelly Nuclear Receptor Signaling and Cardiac Energetics Circ. Res., September 17, 2004; 95(6): 568 - 578. [Abstract] [Full Text] [PDF]
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 J. Zhang, M. Fu, T. Cui, C. Xiong, K. Xu, W. Zhong, Y. Xiao, D. Floyd, J. Liang, E. Li, et al. Selective disruption of PPAR{gamma}2 impairs the development of adipose tissue and insulin sensitivity PNAS, July 20, 2004; 101(29): 10703 - 10708. [Abstract] [Full Text] [PDF]
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 L. L. Spriet, R. J. Tunstall, M. J. Watt, K. A. Mehan, M. Hargreaves, and D. Cameron-Smith Pyruvate dehydrogenase activation and kinase expression in human skeletal muscle during fasting J Appl Physiol, June 1, 2004; 96(6): 2082 - 2087. [Abstract] [Full Text] [PDF]
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 B. Desvergne, L. Michalik, and W. Wahli Be Fit or Be Sick: Peroxisome Proliferator-Activated Receptors Are Down the Road Mol. Endocrinol., June 1, 2004; 18(6): 1321 - 1332. [Abstract] [Full Text] [PDF]
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D. J. Kim, T. E. Akiyama, F. S. Harman, A. M. Burns, W. Shan, J. M. Ward, M. J. Kennett, F. J. Gonzalez, and J. M. Peters Peroxisome Proliferator-activated Receptor {beta} ({delta})-dependent Regulation of Ubiquitin C Expression Contributes to Attenuation of Skin Carcinogenesis J. Biol. Chem., May 28, 2004; 279(22): 23719 - 23727. [Abstract] [Full Text] [PDF]
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M. van Bilsen, P. J.H Smeets, A. J Gilde, and G. J van der Vusse Metabolic remodelling of the failing heart: the cardiac burn-out syndrome? Cardiovasc Res, February 1, 2004; 61(2): 218 - 226. [Abstract] [Full Text] [PDF]
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B. CANNON and J. NEDERGAARD Brown Adipose Tissue: Function and Physiological Significance Physiol Rev, January 1, 2004; 84(1): 277 - 359. [Abstract] [Full Text] [PDF]
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F. Yadetie, A. Laegreid, I. Bakke, W. Kusnierczyk, J. Komorowski, H. L. Waldum, and A. K. Sandvik Liver gene expression in rats in response to the peroxisome proliferator-activated receptor-{alpha} agonist ciprofibrate Physiol Genomics, September 29, 2003; 15(1): 9 - 19. [Abstract] [Full Text] [PDF]
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 C. M. Komar and T. E. Curry Jr Inverse Relationship Between the Expression of Messenger Ribonucleic Acid for Peroxisome Proliferator-Activated Receptor {gamma} and P450 Side Chain Cleavage in the Rat Ovary Biol Reprod, August 1, 2003; 69(2): 549 - 555. [Abstract] [Full Text] [PDF]
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M. C. Sugden and M. J. Holness Recent advances in mechanisms regulating glucose oxidation at the level of the pyruvate dehydrogenase complex by PDKs Am J Physiol Endocrinol Metab, May 1, 2003; 284(5): E855 - E862. [Abstract] [Full Text] [PDF]
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A. J. Gilde, K. A.J.M. van der Lee, P. H.M. Willemsen, G. Chinetti, F. R. van der Leij, G. J. van der Vusse, B. Staels, and M. van Bilsen Peroxisome Proliferator-Activated Receptor (PPAR) {alpha} and PPAR{beta}/{delta}, but not PPAR{gamma}, Modulate the Expression of Genes Involved in Cardiac Lipid Metabolism Circ. Res., March 21, 2003; 92(5): 518 - 524. [Abstract] [Full Text] [PDF]
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M. Jalouli, L. Carlsson, C. Ameen, D. Linden, A. Ljungberg, L. Michalik, S. Eden, W. Wahli, and J. Oscarsson Sex Difference in Hepatic Peroxisome Proliferator-Activated Receptor {alpha} Expression: Influence of Pituitary and Gonadal Hormones Endocrinology, January 1, 2003; 144(1): 101 - 109. [Abstract] [Full Text] [PDF]
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D. M. Muoio, P. S. MacLean, D. B. Lang, S. Li, J. A. Houmard, J. M. Way, D. A. Winegar, J. C. Corton, G. L. Dohm, and W. E. Kraus Fatty Acid Homeostasis and Induction of Lipid Regulatory Genes in Skeletal Muscles of Peroxisome Proliferator-activated Receptor (PPAR) alpha Knock-out Mice. EVIDENCE FOR COMPENSATORY REGULATION BY PPARdelta J. Biol. Chem., July 12, 2002; 277(29): 26089 - 26097. [Abstract] [Full Text] [PDF]
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