THE JOURNAL OF BIOLOGICAL CHEMISTRY© 2003 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 278, No. 28, Issue of July 11, pp. 25395–25400, 2003
Printed in U.S.A.

p53 Activation in Adipocytes of Obese Mice*
Received for publication, March 6, 2003, and in revised form, April 14, 2003
Published, JBC Papers in Press, May 6, 2003, DOI 10.1074/jbc.M302364200

Naoya Yahagi‡, Hitoshi Shimano§¶, Takashi Matsuzaka§, Yuho Najima‡, Motohiro Sekiya‡,
Yoshimi Nakagawa§, Tomohiro Ide§, Sachiko Tomita‡, Hiroaki Okazaki‡, Yoshiaki Tamura‡,
Yoko Iizuka‡, Ken Ohashi‡, Takanari Gotoda‡, Ryozo Nagai‡, Satoshi Kimura‡, Shun Ishibashi‡,
Jun-ichi Osuga‡, and Nobuhiro Yamada§From the ‡Department of Internal Medicine, Graduate School of Medicine, University of Tokyo, Tokyo 113-8655 andthe §Department of Internal Medicine, Institute of Clinical Medicine, University of Tsukuba, Ibaraki 305-8575, JapanThe tumor suppressor p53 is a transcription factor
that activates or represses its target genes after various
genotoxic stresses. We have previously shown that sterol regulatory element-binding protein-1 (SREBP-1), a
key transcriptional regulator of triglyceride synthesis,
and the lipogenic enzymes under its control are markedly suppressed in adipocytes from genetically obese
ob/ob mice. Here we demonstrate that p53 and its target
genes are highly induced in adipocytes of ob/ob mice ina fed state, leading to the negative regulation of
SREBP-1 and thereby lipogenic genes. In fact, disruption of p53 in ob/ob mice completely suppressed the p53regulated genes to wild-type levels and partially restored expression of lipogenic enzymes. Consistently,
reporter gene analysis showed that p53 overexpression
suppressed the promoter activity of the SREBP-1c geneand its downstream genes. Thus, the activation of p53
might constitute a negative feedback loop against excess fat accumulation in adipocytes. In conclusion, we
discovered a novel role of p53 in the pathophysiology ofobesity.

Obesity is a major health problem in industrialized societies,
affecting ⬃20 – 40% of adults (1). The genetically obese ob/ob
mice develop obesity, insulin resistance, and glucose intolerance owing to an inherited deficiency of the appetite-suppressing hormone, leptin (2– 6). The absence of leptin presents themost severe obesity known in both rodents and humans (7), andprovides a good model of obesity and its related syndromes,
including insulin resistance. Although the underlying mechanisms that link obesity and defective insulin signaling are as
yet undefined, hypertrophied adipocyte-derived cytokines suchas tumor necrosis factor (TNF)1-␣ have been reported to be
mediators of insulin resistance in obesity (8, 9).
In the insulin signaling pathways, a transcription factor
sterol regulatory element-binding protein-1 (SREBP-1) has recently been established to be a key molecule for the transcriptional regulation of triglyceride synthesis (10). SREBPs are* This work was supported by grants-in-aid from the Ministry of
Science, Education, Culture, and Technology of Japan. The costs ofpublication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked “advertisement”
in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
¶ To whom correspondence should be addressed: Dept. of Internal
Medicine, Institute of Clinical Medicine, University of Tsukuba, 1-1-1
Tennodai, Tsukuba, Ibaraki 305-8575, Japan. Fax: 81-298-63-2081;
E-mail: shimano-tky@umin.ac.jp.
1
The abbreviations used are: TNF, tumor necrosis factor; SREBP,
sterol regulatory element-binding protein-1; IGFBP-3, insulin-likegrowth factor-binding protein-3; GPDH, glycerol-3- phosphate
dehydrogenase.
This paper is available on line at http://www.jbc.org

members of the basic helix-loop-helix leucine zipper family oftranscription factors that regulate fatty acid and cholesterolsynthesis (reviewed in Refs. 11–13). Whereas SREBP-2 plays acrucial role in regulation of cholesterol synthesis, SREBP-1
controls the transcription and expression of lipogenic enzymes
such as fatty acid synthase (FAS) (reviewed in Refs. 14 –17). In
fact, SREBP-1 and its downstream lipogenic enzymes are drastically induced when fasted animals are refed (18). These lipogenic genes belong to the group of genes that are induced moststrongly by glucose/insulin2 and can be regarded as indicatorsof insulin signaling.
We have recently reported that the refeeding responses of
SREBP-1 and its downstream lipogenic enzymes are markedly
suppressed in adipocytes of ob/ob mice, which is presumably
associated with impaired insulin signaling (19). Although theprecise role of this down-regulation is currently undefined, it
could be a negative feedback mechanism to prevent excess fat
accumulation in extremely obese animals.
The p53 gene was the first tumor suppressor gene to be
identified and has been found to be inactivated in most humancancers (20). The p53 protein is responsible for preventing
division of stressed cells and even causes programmed cell
death (apoptosis) through activation and/or suppression of thetranscription of target genes. For example, ␥-irradiation activates p53 to turn on the transcription of p21Waf1/CIP1, whichbinds to and inhibits cyclin-dependent kinases, thus blocking
the G1-S and G2-to-mitosis transitions. p53 not only activatestranscription of genes such as p21 through its response element, but also represses genes lacking the element by binding
to and sequestering essential transcription factors such as
TATA-binding protein (21, 22). The stresses that activate p53
are diverse, ranging from DNA damage to oxidative stress,
hypoxia, and heat shock (23). The cytostatic and cytotoxic effects of TNF␣ were also demonstrated to be mediated, at leastin part, by p53 activation (24 –26). Thus, p53 has been thought
to be a guardian angel against cellular stresses. Especially, it
has been extensively studied and well established as a tumor
suppressor. However, other roles of p53 beyond tumor suppression are still obscure.
Considering that TNF␣ is relevant to both cell growth andmetabolic events and that its effects are partly mediated by
p53, we speculated that p53 could be involved in situations ofmetabolic deterioration associated with insulin resistance. It is
possible that p53 as a general repressor of gene transcription
could prevent insulin-responsive genes from being activated.
Moreover, a previous report on the gene expression profile ofob/ob mouse adipose tissue examined by DNA microarray analysis has revealed that p21 and Bax ␣, both of which are well2

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N. Yahagi, unpublished data.

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p53 Activation in Adipocytes of Obese Miceknown p53 target genes, are increased from 2- to 3-fold in ob/ob
mice (27).
Based on these facts, we hypothesized that hypertrophied
adipocytes are under various stresses that induce p53, which inturn suppresses lipogenesis in a negative feedback regulation.
In our present study, we discovered that p53 is induced uponrefeeding in ob/ob adipocytes and activates its target genes
including p21. In addition, p53 is involved in the suppression of
SREBP-1 and the concomitant down-regulation of lipogenic
enzymes.
EXPERIMENTAL PROCEDURES

Animals—p53⫹/⫺-C57BL/6J (28) and Lep⫹/ob-C57BL/6J mice werepurchased from the Jackson Laboratories (Bar Harbor, Maine). These
mice were intercrossed to produce double homozygotes. Genotypes atthe p53 loci were determined using the PCR method with primer pairs
specific for the wild-type allele of p53 (primers directed against p53
exon 6 and 7; 5⬘-GGAAATTTGTATCCCGAGTATCTG-3⬘ and 5⬘-GTCTTCCAGTGTGATGATGGTAA-3⬘, respectively) and the mutant allele ofp53 (primers directed against neo; 5⬘-ATGATTGAACAAGATGGATTGC-3⬘ and 5⬘-TCGGTCTTGACAAAAAGAACC-3⬘). Genotypes at theleptin loci were determined by a PCR-based restricted fragment length
polymorphism analysis as previously described (19). Mice were housed
in a temperature-controlled environment with a 12-h light/dark cycleand free access to water and a standard chow diet (Oriental MF,
Oriental Yeast, Tokyo, Japan). All experiments were performed with
12-week-old male mice. For fasting and refeeding studies, mice were
fasted for 24 h or refed for 12 h after 24-h starvation. For the ob/ob⫻p53⫺/⫺ experiment, mice were refed for 12 h following a 24-h fastprior to sacrifice. All animals were sacrificed in an early phase of thelight cycle.
Nuclear Protein Extraction and Immunoblotting—Nuclear extract
protein from white adipose tissue was prepared as described previously(27). Briefly, fresh adipose tissue (⬃3 g pooled from 3–10 male mice)
was rinsed in ice-cold PBS, minced, and homogenized with 10 strokes ofa Teflon homogenizer in 15 ml of NDS buffer at 4 °C (10 mM Tris, pH
7.5, 10 mM NaCl, 60 mM KCl, 0.15 mM spermine, 0.5 mM spermidine, 14mM mercaptoethanol, 0.5 mM EGTA, 2 mM EDTA, 0.5% Nonidet P-40, 1
mM dithiothreitol) supplemented with protease inhibitors (12.5 ␮g/ml
N-Acetyl-Leu-Leu-norleucinal-CHO (ALLN, Calbiochem), 2.5 ␮g/ml
leupeptin, 2.5 ␮g/ml aprotinin, 2.5 ␮g/ml pepstatin A, 0.1 mM phenylmethylsulfonyl fluoride). Nonidet P-40 concentration was increased to
1% and nuclei were pelleted at 700 ⫻ g for 10 min, washed once with 25
ml NDS buffer (1% Nonidet P-40), filtered through 70 ␮m mesh, pelleted at 500 ⫻ g for 10 min, resuspended in 1 volume of 1% citric acid,lysed by the addition of 2.5 volumes of 0.1 M Tris, 2.5% SDS and 0.1 M
dithiothreitol, sonicated briefly, and heated to 90 °C for 5 min. Aliquotsof nuclear protein (15 ␮g) were subjected to SDS/PAGE. p53 was detected using a 1:500 dilution of anti-p53FL (sc-6243, Santa Cruz Biotechnology). The phosphorylation at Ser-15 of p53 was detected using a
1:1000 dilution of anti-p53Ser-15 (9284S, New England Biolabs). To
confirm equal loading of nuclear protein, a 1:500 dilution of anti-c-myc(sc-764, Santa Cruz Biotechnology) was also used. Bound antibodies
were detected using a horseradish peroxidase-coupled anti-rabbit IgG
secondary antibody (Amersham Biosciences) and visualized using ECLWestern blotting detection system kit (Amersham Biosciences).
Northern Blotting—Total RNA from epididymal fat pad was extracted using Trizol reagent (Invitrogen), and 10 ␮g RNA samples
equally pooled from each group (n ⫽ 3– 6) were run on a 1% agarose gel
containing formaldehyde and transferred to a nylon membrane. For
experiments with isolated adipocytes, total RNA from adipocytes isolated by collagenase digestion was prepared as previously described(29). In brief, fresh adipose tissue (⬃4 g) was minced in 8 ml buffer A (10mM HEPES at pH 7.4, 130 mM NaCl, 5.2 mM KCl, 1.3 mM KH2PO4, 2.7
mM CaCl2, 1.3 mM MgSO4, 25 mM NaHCO3, 3% BSA, 2 mM glucose, 200
nM adenosine) at 37 °C and mixed with equal volume of buffer A
containing 4 mg/ml collagenase (type II for adipocyte, Sigma). Following
1-hour digestion with gentle shaking at 37 °C, adipocytes were filteredthrough 250 ␮m nylon mesh, washed with 30 ml of buffer A, transferred
to another tube, and mixed with 4 volumes of Trizol reagent (Invitrogen). Total RNA sample (10 ␮g) from one mouse was applied on eachlane. The cDNA probes for SREBP-1, fatty acid synthase, ATP citratelyase, spot 14 and 36B4 (acidic ribosomal phosphoprotein P0) werecloned as described previously (30, 31). The cDNA probes for p53, p21,
p27, mdm-2, Bax ␣, insulin-like growth factor-binding protein-3 (IGFBP-3), glycerol-3-phosphate dehydrogenase (GPDH) and tumor necro-

sis factor ␣ (TNF␣) were prepared by cloning RT-PCR products frommouse liver or adipose tissue RNA into pGEM-T easy vectors (Promega). The primers used for PCR were as follows: for p53, 5⬘ primer was
5⬘-GGAAATTTGTATCCCGAGTATCTG-3⬘ and 3⬘ primer was 5⬘-GTCTTCCAGTGTGATGATGGTAA-3⬘; for p21, 5⬘ primer was 5⬘-TGTCCAATCCTGGTGATGTC-3⬘ and 3⬘ primer was 5⬘-TCTCTTGCAGAAGACCAATCTG-3⬘; for p27, 5⬘ primer was 5⬘-GTCAAACGTGAGAGTGTCTAACG-3⬘ and 3⬘ primer was 5⬘-GCGAAGAAGAATCTTCTGCA-3⬘; formdm-2, 5⬘ primer was 5⬘-CCAGGCCAATGTGCAATAC-3⬘ and 3⬘
primer was 5⬘-GTGAGCAGGTCAGCTAGTTGAA-3⬘; for Bax ␣, 5⬘primer was 5⬘-GCTCTGAACAGATCATGAAGACA-3⬘ and 3⬘ primer
was 5⬘-CATGATGGTTCTGATCAGCTC-3⬘; for IGFBP-3, 5⬘ primer was
5⬘-CATGCCAAGATGGATGTCATC-3⬘ and 3⬘ primer was 5⬘-GAGGCAATGTACGTCGTCTTTC-3⬘; for GPDH, 5⬘ primer was 5⬘-CCATGGCTGGCAAGAAAGT-3⬘ and 3⬘ primer was 5⬘-AATCACTTCAGAAATGAGCTTCAG-3⬘; for TNF␣, 5⬘ primer was 5⬘-GGTTCTGTCCCTTTCACTCACTG-3⬘ and 3⬘ primer was 5⬘-TTGACCTCAGCGCTGAGTTG-3⬘. The
probes were labeled with [␣-32P]dCTP using Megaprime DNA labelingsystem kit (Amersham Biosciences). The membranes were hybridized
with the radiolabeled probe in Rapid-hyb Buffer (Amersham Biosciences) at 65 °C with the exception of p53 and TNF␣, for which ULTRAhyb hybridization buffer (Ambion) was used at 42 °C. The membranes were washed in 0.1⫻ SSC, 0.1% SDS at 65 °C. Blots were exposed
to Kodak XAR-5 film.
Plasmid Constructions—Luciferase gene constructs containing
2.6-kb fragments of the mouse SREBP-1c promoter (pBP1c-Luc) and
0.24-kb fragments of the rat fatty acid synthase promoter (pFAS-Luc)
were prepared as described previously (32, 33). Plasmid pGPDH-Luc
was constructed by PCR amplification of ⫺489 bp to ⫹23 bp of themouse glycerol-3-phosphate dehydrogenase (GPDH) promoter region(34) and insertion of the PCR products into pGL2-basic vector (Promega). Plasmid p53-Luc, a luciferase reporter plasmid containing 14
tandem copies of p53-binding motif (TGCCTGGACTTGCCTGG) andthe luciferase gene was purchased from Stratagene. Mouse p53 expression plasmid driven by the CMV promoter (pCMV-p53) was constructedby inserting a DNA fragment amplified with PCR using first strand
cDNA from mouse adipose tissue as a template, two primers (5⬘ primer:
5⬘-TGGCTGTAGGTAGCGACTACAGTTA-3⬘ and 3⬘ primer: 5⬘-AGGCAGTCAGTCTGAGTCAGG-3⬘) and Platinum Pfx DNA polymerase(Invitrogen) into pCMV7 vector (35). All DNA fragments generated with
PCR were verified by sequencing.
Transfections and Luciferase Assays—p53-null human osteogenic
sarcoma cell line SaOS-2 was obtained from the American Type Culture
Collection (ATCC). SaOS-2 cells were cultured in Dulbecco’s modified
Eagle’s medium containing 25 mM glucose, 100 units/ml penicillin, and100 ␮g/ml streptomycin sulfate supplemented with 10% fetal calf serum
under 5% CO2 at 37 °C. On day 0, cells were plated on a 12 well plate
at 5 ⫻ 104 cells/well. On day 3, pCMV-p53 plasmid (1.0 ␮g), eachluciferase reporter plasmid (0.25 ␮g), and a CMV-␤-galactosidase reference plasmid (pCMV-␤-gal, Promega, 0.1 ␮g) were transfected into
SaOS-2 cells using FuGENE 6 transfection reagent (Roche Applied
Science) according to the manufacturer’s protocol. As a control, thesame amount of empty vector pCMV7 was transfected instead of pCMVp53 plasmid. After 48-h incubation, cells were harvested, and theamount of luciferase activity in transfectants was measured by standard kits (Promega).
RESULTS

p53 Activation upon Refeeding in ob/ob Adipocytes at themRNA Level—In our first series of experiments, we demonstrated that ob/ob mouse adipose tissue had higher levels of p53
mRNA and protein than did wild type (Fig. 1). p53 is known tobe regulated at different levels including through transcription, protein phosphorylation and degradation, and we foundthat the primary step of p53 activation in ob/ob mouse adipose
tissue was at mRNA level, although this does not rule out
additional post-transcriptional regulations. Consistent with
the induced expression, an active form of p53, phosphorylated
at Ser-15, was also up-regulated in ob/ob adipose tissue (Fig.
1b).
Next we evaluated the mRNA abundance of p53 under various nutritional conditions. To our surprise, the increased expression of p53 in ob/ob mouse adipose tissue was limited to a
fed state and no difference was observed in a fasted state (Fig.
2a). The mRNA elevation of p53 appeared to be fully induced

p53 Activation in Adipocytes of Obese Mice

FIG. 1. p53 is increased in ob/ob mouse adipose tissue at themRNA level. a, Northern blot and b, Western blot analysis of p53 inwild type (WT) and ob/ob (OB) mouse adipose tissue. Mice were sacrificed in refed state. a, total RNA (10 ␮g) was subjected to Northernblotting, followed by hybridization with the indicated cDNA probes. A
cDNA probe for 36B4 (acidic ribosomal phosphoprotein P0) was used toconfirm equal loading. b, immunoblotting using antibody against fulllength p53 protein (left upper panel) and against phosphorylated Ser-15(right upper panel) is shown. Immunoblotting using antibody against
c-Myc (lower panel) is shown to indicate equal loading.

FIG. 2. p53 induction upon refeeding in ob/ob mouse adipocytes. a, Northern blot analysis of p53 and its downstream genes inadipose tissue from wild type (WT) and ob/ob (OB) mice (six male miceeach) is shown. Mice were sacrificed in a fasted (left two lanes) or refedstate. b, time course of p53 induction after refeeding in ob/ob mouseadipose tissue. Northern blot analysis of p53 in ob/ob mice (three malemice each) refed for the indicated time is shown. c, Northern blot
analysis of isolated adipocytes from fasted (left three lanes) or refed(right three lanes) ob/ob mice. Total RNA (10 ␮g) was subjected to
Northern blotting, followed by hybridization with the indicated cDNA
probes. A cDNA probe for 36B4 (acidic ribosomal phosphoprotein P0)
was used to confirm equal loading.

within 6 h after refeeding (Fig. 2b). p53 was revealed to be
induced mainly in adipocytes when they were isolated fromstromal cells by collagenase digestion (Fig. 2c), and the residual
stromal cells showed far lower levels of p53 expression which
did not rise upon refeeding (data not shown).
p53 Activation in ob/ob Adipocytes Is Associated with Induc-

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FIG. 3. Effects of p53 absence on the mRNA expression of p53regulated genes in ob/ob mouse adipose tissue. Northern blot
analysis of various genes including p53-regulated genes in adipose
tissue from refed mice. Lanes (left to right) show wild type, ob/ob,
ob/ob⫻p53⫹/⫺ and ob/ob⫻p53⫺/⫺. Total RNA (10 ␮g) pooled equallyfrom six male mice was subjected to Northern blotting, followed byhybridization with the indicated cDNA probes. A cDNA probe for 36B4
(acidic ribosomal phosphoprotein P0) was used to confirm equalloading.

FIG. 4. Up-regulation of p21 by fasting is independent of p53.
Northern blot analysis of various genes including p53-regulated genes
in adipose tissue from wild type and p53⫺/⫺ mice is shown. Mice weresacrificed in a fasted (left two lanes) or refed state. A cDNA probe for
36B4 (acidic ribosomal phosphoprotein P0) was used to confirm equalloading.

tion of p53-regulated Genes—To clarify the potential role of p53
in ob/ob adipocytes, we further examined the expression profile
of p53 downstream genes such as p21, mdm-2, Bax ␣, andinsulin-like growth factor binding protein-3 (IGFBP-3). As
shown in Fig. 2a, these p53-regulated genes were elevated inadipose tissue of refed ob/ob mice. These data indicate that p53
activation causes the up-regulation of its target genes in ob/ob
adipocytes.
Effects of p53 Absence on the mRNA Expression of p53-regulated Genes in ob/ob Mouse Adipose Tissue—To assess theeffects of p53 deficiency in ob/ob mice and validate that theelevation of p53 downstream genes are really caused by p53, weintercrossed ob/ob and p53-null mice, and obtained 6 male mice
deficient in both leptin and p53 (ob/ob⫻p53⫺/⫺) in the
C57BL/6J background. Doubly mutant ob/ob⫻p53⫺/⫺ mice

25398p53 Activation in Adipocytes of Obese Mice

FIG. 5. p53 overexpression suppresses lipogenic gene responses upon refeeding. a, defective refeeding responses of lipogenic genes in
ob/ob mouse adipose tissue. Northern blot analysis of various genes related to lipogenesis in adipose tissue from wild type and ob/ob mice is shown.
Mice were sacrificed in a fasted (left two lanes) or refed state. b, partial de-repression of lipogenic genes in ob/ob mice lacking p53. Northern blot
analysis of lipogenic genes in adipose tissue from refed mice. Lanes (left to right) show wild type, ob/ob and ob/ob⫻p53⫺/⫺. FAS, fatty acid synthase;
ACL, ATP citrate lyase; S14, Spot 14. c, lipogenic gene suppression by p53 overexpression. p53-null SaOS-2 cells were cotransfected with indicatedpromoter-reporter plasmid and an expression vector, either empty pCMV7 vector (䡺) or pCMV-p53 expression vector (f). Luciferase activities areindicated as relative values to control. The figure shows one representative data out of four independent experiments. Bars, S.E. BP1c, sterol
regulatory element-binding protein-1c.

showed no significant difference in body weight, epididymal fat
pad weight, plasma glucose or insulin concentration comparedwith ob/ob mice (data not shown). The Northern blot analysis
on these mice in a refed state (Fig. 3) exhibited that p53regulated genes such as p21, mdm-2, Bax ␣, and IGFBP-3 in
ob/ob mice lacking p53 were completely suppressed to the samelevels as in wild type. These results established that p53 induction after refeeding caused the elevated expression of its
target genes in ob/ob adipocytes. Heterozygotes of p53 genedisruption in ob/ob mice maintained the similar levels of p53
downstream genes to those of ob/ob mice conceivably by compensation by the intact allele.
Up-regulation of p21 and IGFBP-3 by Fasting Is Independentof p53—In our studies of fasted and refed mice, we also foundthat p21 and IGFBP-3 are elevated when wild-type mice are
fasted. To determine whether these changes are ascribable top53, we analyzed p53⫺/⫺ mice in fasted or refed conditions. As
shown in Fig. 4, the elevation of p21 and IGFBP-3 in a fasted
state was also observed in p53⫺/⫺ mice, and hence, entirely
independent of p53.
Defective Refeeding Responses of Lipogenic Genes in ob/ob
Mice Is Mediated by p53—Lipogenic enzymes such as fatty acidsynthase and ATP citrate lyase are known to be markedly
induced in adipose tissue and liver when animals are refedafter starvation. In contrast, we have previously reported thatthe adipose tissue of ob/ob mice shows lower levels and defective refeeding responses in the expression of lipogenic enzymes
as well as SREBP-1 that regulates their transcription (19). In
the current studies we found that the suppression of lipogenic
enzymes in ob/ob adipose tissue is confined to a subgroup of
enzymes such as fatty acid synthase and ATP citrate lyase,whose expression is primarily dominated by SREBP-1 (Fig. 5a).
In contrast, glycerol-3-phosphate dehydrogenase, a key enzyme
for glycerogenesis and also important for lipogenesis, was not
suppressed in refed ob/ob mouse adipose tissue. Based on thelack of change in its gene expression by the SREBP-1 overexpression, we assume that glycerol-3-phosphate dehydrogenase
is not an SREBP-1 target gene.2 These results suggest that p53
suppresses lipogenic gene expression in ob/ob adipocytes by the
inhibition of SREBP-1 expression.
To test this hypothesis, we evaluated the mRNA expressionof lipogenic genes in doubly mutant ob/ob⫻p53⫺/⫺ mice. Northern blot analysis displayed that lipogenic enzymes such as fatty
acid synthase and ATP citrate lyase along with SREBP-1 werep53 Activation in Adipocytes of Obese Micemoderately elevated in ob/ob mice lacking p53, showing thatthe absence of p53 partially de-suppresses lipogenic gene downregulation (Fig. 5b). These findings demonstrate that p53 isinvolved in the suppression of lipogenic genes in ob/ob adipocytes. However, the restoration was limited within relatively
minor range, suggesting that other factors than p53 are involved in this negative regulation.
Mechanism by Which p53 Suppresses Lipogenic Gene Expression—To explore the mechanism by which p53 suppresses lipogenic gene expression, we performed luciferase reporter assays in cultured cells. We used p53-null cell line SaOS-2 as thetransfectant. As shown in Fig. 5c, p53 overexpression suppressed the promoter activity of fatty acid synthase gene aswell as that of SREBP-1c gene. In contrast, the promoter activity of glycerol-3-phosphate dehydrogenase gene was not suppressed by p53, which corresponds to the in vivo results described above. These findings provide further evidence that p53
suppress lipogenic genes by reducing SREBP-1 expression.
DISCUSSION

Our present studies clearly demonstrate that p53 is induced
upon refeeding in ob/ob adipocytes, leading to the negative
regulation of lipogenic genes. The mechanisms of p53 induction
are currently not clear. However, the causes that lead to p53
activation are diverse, ranging from DNA damage to oxidative
stress and hypoxia (23). We speculate that ob/ob adipocytes
receive various stresses that could induce p53 activation. The
autocrine excretion of TNF␣, the expression of which is increased in ob/ob adipocytes (8), could contribute to it. The
enlargement of cell volume might cause intracellular hypoxia
that could activate p53. Furthermore, in a fed state ob/ob miceare markedly hyperglycemic, which also tends to result inoxidative stress through various mechanisms (36 –38). Thus,ob/ob adipocytes are presumably placed under stressful conditions where p53 can be activated.
Recently it was reported that the phosphatidylinositol 3-kinase-Akt pathway plays an important role in the regulation of
p53 via mdm-2 by its enhancement of mdm-2-mediated ubiquitination and degradation of p53 (39). Hence, it is possiblethat insulin could suppress p53 activation by this pathway.
Given that the insulin signaling via phosphatidylinositol 3-kinase and Akt pathway is attenuated in ob/ob adipocytes (40,41) (i.e. insulin resistance), this mechanism might be involvedin the elevation of p53.
Conversely, it is also possible that the p53 activation could
cause insulin resistance and that p53 could be positioned as amodifier of insulin signaling. In addition to our present findingthat p53 down-regulates SREBP-1 and thereby lipogenesis,
p53 is reported to up-regulate the phosphatase PTEN (42),which opposes phosphatidylinositol 3-kinase and has been implicated to be related to insulin resistance (43, 44). These datasuggest that p53 could negatively regulate insulin action in a
broad spectrum. The concept that p53 could compete withinsulin signaling in nutritional regulation is highly probableconsidering that insulin ancestrally belongs to a growth factor
family closely related to proto-oncogenes, whereas p53 is atumor suppressor gene.
Obesity is defined as an enlargement of adipose tissue mass
and, in case of ob/ob mice, is due to both hyperplasia andhypertrophy of adipocytes (4, 5). Therefore, the hyperplastic
adipose tissue in obesity could be regarded as a kind of benign
tumor. In fact, it has been recently reported that anti-angiogenic agents developed to treat tumors are also effective forobesity (45). It is well known that in many tumors, althoughmutated, p53 mRNA are expressed at elevated levels (46). The
elevation of p53 both in tumors and in ob/ob adipose tissue

25399

might result from a common abnormality characteristic of hyperplastic tissue, for instance, low vascularity.
With respect to the consequences of p53 activation in ob/ob
adipocytes, we are speculating that, besides insulin signaling,
p53 in ob/ob adipocytes might influence cellular turnover processes. Adipocytes in ob/ob mice are reported to be prone toapoptosis (47). Moreover, some markers of immature adipocytes such as adipocyte differentiation-related protein are increased (48). These facts might perhaps imply that the turnover rate of adipocytes is higher and immature cells are
relatively increased in obese animals. Thus, although the exact
life cycle of adipocytes is not known and precise measurementof their life span needs to be analyzed, p53 might shorten thelife span of adipocytes. In keeping with this, it has been recently reported that a transgenic mouse with chronic p53 activation exhibits accelerated aging of organs including adipose
tissue (49).
In conclusion, we discovered that p53 is induced upon refeeding in ob/ob adipocytes, a mechanism by which lipogenic genes
are negatively regulated. This finding might open up a new
aspect of this diverse regulator relating cell growth and nutritional regulation.
Acknowledgment—We thank Alyssa H. Hasty for critical reading of
the manuscript.
REFERENCES
1. Kuczmarski, R. J., Flegal, K. M., Campbell, S. M., and Johnson, C. L. (1994)
J. Am. Med. Assoc. 272, 205–211
2. Ingalls, A. M., Dickie, M. M., and Snell, G. D. (1950) J. Hered. 41, 317–318
3. Mayer, J., Bates, M. W., and Dickie, M. M. (1951) Science 113, 746 –747
4. Herberg, L., and Coleman, D. L. (1977) Metabolism 26, 59 –99
5. Bray, G. A., and York, D. A. (1979) Physiol. Rev. 59, 719 – 809
6. Zhang, Y., Proenca, R., Maffei, M., Barone, M., Leopold, L., and Friedman,
J. M. (1994) Nature 372, 425– 432
7. Montague, C. T., Farooqi, I. S., Whitehead, J. P., Soos, M. A., Rau, H.,
Wareham, N. J., Sewter, C. P., Digby, J. E., Mohammed, S. N., Hurst, J. A.,
Cheetham, C. H., Earley, A. R., Barnett, A. H., Prins, J. B., and O’Rahilly,
S. (1997) Nature 387, 903–908
8. Hotamisligil, G. S., Shargill, N. S., and Spiegelman, B. M. (1993) Science 259,
87–91
9. Uysal, K. T., Wiesbrock, S. M., Marino, M. W., and Hotamisligil, G. S. (1997)
Nature 389, 610 – 614
10. Foretz, M., Guichard, C., Ferre, P., and Foufelle, F. (1999) Proc. Natl. Acad.
Sci. U. S. A. 96, 12737–12742
11. Brown, M. S., and Goldstein, J. L. (1997) Cell 89, 331–340
12. Brown, M. S., and Goldstein, J. L. (1999) Proc. Natl. Acad. Sci. U. S. A. 96,
11041–11048
13. Brown, M. S., Ye, J., Rawson, R. B., and Goldstein, J. L. (2000) Cell 100,391–398
14. Shimano, H. (2000) Trends Cardiovasc. Med. 10, 275–278
15. Shimano, H. (2001) Prog. Lipid Res. 40, 439 – 452
16. Shimano, H. (2002) Vitam. Horm. 65, 167–194
17. Horton, J. D., Goldstein, J. L., and Brown, M. S. (2002) J. Clin. Invest. 109,
1125–1131
18. Horton, J. D., Bashmakov, Y., Shimomura, I., and Shimano, H. (1998) Proc.
Natl. Acad. Sci. U. S. A. 95, 5987–5992
19. Yahagi, N., Shimano, H., Hasty, A. H., Matsuzaka, T., Ide, T., Yoshikawa, T.,
Amemiya-Kudo, M., Tomita, S., Okazaki, H., Tamura, Y., Iizuka, Y.,
Ohashi, K., Osuga, J., Harada, K., Gotoda, T., Nagai, R., Ishibashi, S., andYamada, N. (2002) J. Biol. Chem. 277, 19353–19357
20. Vogelstein, B., Lane, D., and Levine, A. J. (2000) Nature 408, 307–310
21. Seto, E., Usheva, A., Zambetti, G. P., Momand, J., Horikoshi, N., Weinmann,
R., Levine, A. J., and Shenk, T. (1992) Proc. Natl. Acad. Sci. U. S. A. 89,
12028 –12032
22. Mack, D. H., Vartikar, J., Pipas, J. M., and Laimins, L. A. (1993) Nature 363,
281–283
23. Oren, M. (1999) J. Biol. Chem. 274, 36031–36034
24. Jeoung, D.-i., Tang, B., and Sonenberg, M. (1995) J. Biol. Chem. 270,
18367–18373
25. Donato, N. J., and Perez, M. (1998) J. Biol. Chem. 273, 5067–5072
26. Rokhlin, O. W., Gudkov, A. V., Kwek, S., Glover, R. A., Gewies, A. S., andCohen, M. B. (2000) Oncogene 19, 1959 –1968
27. Soukas, A., Cohen, P., Socci, N. D., and Friedman, J. M. (2000) Genes Dev. 14,
963–980
28. Livingstone, L. R., White, A., Sprouse, J., Livanos, E., Jacks, T., and Tlsty,
T. D. (1992) Cell 70, 923–935
29. Rodbell, M. (1964) J. Biol. Chem. 239, 375–380
30. Shimano, H., Horton, J. D., Hammer, R. E., Shimomura, I., Brown, M. S., andGoldstein, J. L. (1996) J. Clin. Invest. 98, 1575–1584
31. Shimano, H., Yahagi, N., Amemiya-Kudo, M., Hasty, A. H., Osuga, J., Tamura,Y., Shionoiri, F., Iizuka, Y., Ohashi, K., Harada, K., Gotoda, T., Ishibashi,S., and Yamada, N. (1999) J. Biol. Chem. 274, 35832–35839
32. Amemiya-Kudo, M., Shimano, H., Yoshikawa, T., Yahagi, N., Hasty, A. H.,

25400p53 Activation in Adipocytes of Obese Mice

Okazaki, H., Tamura, Y., Shionoiri, F., Iizuka, Y., Ohashi, K., Osuga, J.,
Harada, K., Gotoda, T., Sato, R., Kimura, S., Ishibashi, S., and Yamada, N.
(2000) J. Biol. Chem. 275, 31078 –31085
33. Shimano, H., Horton, J. D., Shimomura, I., Hammer, R. E., Brown, M. S., andGoldstein, J. L. (1997) J. Clin. Invest. 99, 846 – 854
34. Phillips, M., Djian, P., and Green, H. (1986) J. Biol. Chem. 261, 10821–10827
35. Andersson, S., Davis, D. L., Dahlback, H., Jornvall, H., and Russell, D. W.
(1989) J. Biol. Chem. 264, 8222– 8229
36. Williamson, J. R., Chang, K., Frangos, M., Hasan, K. S., Ido, Y., Kawamura, T.,
Nyengaard, J. R., van den Enden, M., Kilo, C., and Tilton, R. G. (1993)
Diabetes 42, 801– 813
37. Inoguchi, T., Li, P., Umeda, F., Yu, H. Y., Kakimoto, M., Imamura, M., Aoki,
T., Etoh, T., Hashimoto, T., Naruse, M., Sano, H., Utsumi, H., and Nawata,
H. (2000) Diabetes 49, 1939 –1945
38. Kaneto, H., Xu, G., Song, K. H., Suzuma, K., Bonner-Weir, S., Sharma, A., andWeir, G. C. (2001) J. Biol. Chem. 276, 31099 –31104
39. Ogawara, Y., Kishishita, S., Obata, T., Isazawa, Y., Suzuki, T., Tanaka, K.,
Masuyama, N., and Gotoh, Y. (2002) J. Biol. Chem. 277, 21843–21850
40. Shao, J., Yamashita, H., Qiao, L., and Friedman, J. E. (2000) J. Endocrinol.
167, 107–115
41. Carvalho, E., Rondinone, C., and Smith, U. (2000) Mol. Cell Biochem. 206,

7–16
42. Stambolic, V., MacPherson, D., Sas, D., Lin, Y., Snow, B., Jang, Y., Benchimol,S., and Mak, T. W. (2001) Mol. Cell 8, 317–325
43. Nakashima, N., Sharma, P. M., Imamura, T., Bookstein, R., and Olefsky, J. M.
(2000) J. Biol. Chem. 275, 12889 –12895
44. Butler, M., McKay, R. A., Popoff, I. J., Gaarde, W. A., Witchell, D., Murray,
S. F., Dean, N. M., Bhanot, S., and Monia, B. P. (2002) Diabetes 51,
1028 –1034
45. Rupnick, M. A., Panigrahy, D., Zhang, C. Y., Dallabrida, S. M., Lowell, B. B.,
Langer, R., and Folkman, M. J. (2002) Proc. Natl. Acad. Sci. U. S. A. 99,
10730 –10735
46. Balint, E., and Reisman, D. (1996) Cancer Res. 56, 1648 –1653
47. Loftus, T. M., Kuhajda, F. P., and Lane, M. D. (1998) Proc. Natl. Acad. Sci.
U. S. A. 95, 14168 –14172
48. Nadler, S. T., Stoehr, J. P., Schueler, K. L., Tanimoto, G., Yandell, B. S., andAttie, A. D. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 11371–11376
49. Tyner, S. D., Venkatachalam, S., Choi, J., Jones, S., Ghebranious, N.,
Igelmann, H., Lu, X., Soron, G., Cooper, B., Brayton, C., Hee Park, S.,
Thompson, T., Karsenty, G., Bradley, A., and Donehower, L. A. (2002)
Nature 415, 45–53