Brown Remodeling of White Adipose
Tissue by SirT1-Dependent
Deacetylation of Pparg
Li Qiang,1Liheng Wang,1,6Ning Kon,2,6Wenhui Zhao,2Sangkyu Lee,4Yiying Zhang,3Michael Rosenbaum,3
Yingming Zhao,4Wei Gu,2Stephen R. Farmer,5and Domenico Accili1,*
1Naomi Berrie Diabetes Center, Department of Medicine
2Institute of Cancer Genetics, Department of Pathology
3Division of Molecular Genetics, Department of Pediatrics
College of Physicians and Surgeons of Columbia University, New York, NY 10032, USA
4Ben May Department for Cancer Research, University of Chicago, Chicago, IL 60637, USA
5Department of Biochemistry, Boston University School of Medicine, Boston, MA 02118, USA
6These authors contributed equally to this work
Brown adipose tissue (BAT) can disperse stored
energy as heat. Promoting BAT-like features in white
adipose (WAT) is an attractive, if elusive, therapeutic
approach to staunch the current obesity epidemic.
Here we report that gain of function of the NAD-
dependent deacetylase SirT1 or loss of function of
its endogenous inhibitor Deleted in breast cancer-1
(Dbc1) promote ‘‘browning’’ of WAT by deacetylating
peroxisome proliferator-activated receptor (Ppar)-g
on Lys268 and Lys293. SirT1-dependent deacety-
lation of Lys268 and Lys293 is required to recruit
the BAT program coactivator Prdm16 to Pparg,
leading to selective induction of BAT genes and
repression of visceral WAT genes associated with
insulin resistance. An acetylation-defective Pparg
mutant induces a brown phenotype in white adipo-
cytes, whereas an acetylated mimetic fails to induce
‘‘brown’’ genes but retains the ability to activate
‘‘white’’ genes. We propose that SirT1-dependent
Pparg deacetylation is a form of selective Pparg
modulation of potential therapeutic import.
Obesity and its comorbidities pose a growing therapeutic
challenge (Wang et al., 2011). White adipose tissue (WAT) is
the main ‘‘storage site’’ of excess energy, primarily in the form
of triglycerides. In addition, a functionally and morphologically
distinct adipocyte subset—whose dense mitochondrial, inner-
vation, and vascular content earned it the moniker of ‘‘brown’’
adipose tissue (BAT)—dissipates energy as heat (nonshivering
thermogenesis). Brown adipocytes uncouple mitochondrial
electron transport from ATP synthesis to a greater extent than
other cells by permeabilizing the inner mitochondrial membrane
to allow inter-membrane proton to leak back into the mitochon-
drial matrix, primarily through uncoupling protein-1 (Ucp1), but
also through other mitochondrial proteins (Ravussin and Gal-
gani, 2011). Promoting BAT function has therapeutic potential
to combat obesity (Farmer, 2009). But its limited amount and
activity in humans are unlikely to offset the positive energy
balance associated with excessive WAT deposition (Virtanen
and Nuutila, 2011).
As an alternative strategy to increase energy expenditure and
prevent weight gain, we investigated mechanisms that confer
BAT-like features onto WAT, thus remodeling the latter from an
energy storage into an energy-disposal site (Kozak, 2010). The
metabolic benefits of this conversion include prevention of
diet-induced obesity and increased insulin sensitivity (Seale
et al., 2011). Browning of rodent WAT can be brought about by
hormones and cytokines, such as Irisin (Bostro ¨m et al., 2012)
and Fgf21 (Fisher et al., 2012), as well as by transcriptional
modulation through Prdm16 (Seale et al., 2011), FoxC2 (Ceder-
berg et al., 2001), RIP140 (Powelka et al., 2006), 4E-BP1
(Tsukiyama-Kohara et al., 2001), TIF2 (Picard et al., 2002), pRb
and p107 (Scime ` et al., 2005). However, there is an unmet
need for strategies that would translate these mechanisms into
Activation of the nuclear receptor Pparg by thiazolidinediones
(TZDs) can also induce a brown-like phenotype in white adipo-
cytes by promoting expression of brown adipocyte-specific
genes (brown genes) and suppressing visceral WAT genes
(white genes) (Vernochet et al., 2009). The mechanism of this
‘‘browning’’ effect is unclear, and is unlikely to be clinically
applicable without further modulation, in view of the adverse
effects associated with TZD use (Kim-Muller and Accili, 2011).
Activation of the NAD+-dependent deacetylase SirT1 by
small molecules, calorie restriction or exercise promotes mito-
chondrial biogenesis and activities (Canto ´
Milne et al., 2007), raising the possibility that SirT1 regulates
BAT functions. Furthermore, SirT1 gain-of-function mimics the
et al., 2009;
620 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
insulin-sensitizing function of Pparg ligands in vivo (Banks
et al., 2008). In view of these facts, we asked whether the
browning activity of Pparg is mediated through its SirT1-depen-
dent deacetylation and whether SirT1 is also capable of
inducing browning of WAT, similar to TZDs. We report that
SirT1-dependent Pparg deacetylation promotes browning of
subcutaneous WAT by regulating ligand-dependent coactiva-
tor/corepressor exchange at the Pparg transcriptional complex.
We propose that SirT1-dependent Pparg deacetylation regu-
lates energy homeostasis, promoting energy expenditure over
SirT1 Deacetylates Pparg in a Ligand-Dependent
We investigated whether Pparg activation by TZD ligands
affects its acetylation. Indeed, treatment with rosiglitazone
dose-dependently decreased Pparg acetylation (Figure 1A).
Moreover, Pparg agonists facilitated, while the antagonist
GW9662 prevented the interaction between Pparg and SirT1
(Figures 1A and 1B). Pparg acetylation was augmented by ace-
tyltransferase Cbp (Figure 1C) or HDAC inhibitors trichostatin
A (TSA) and nicotinamide (Figure S1A available online).
Conversely, SirT1 overexpression or chemical activation with
resveratrol decreased Pparg acetylation levels (Figures 1C and
S1B). To determine whether Pparg is a SirT1 substrate, we per-
formed in vitro deacetylation assays with purified SirT1 and
acetylated Pparg. The data demonstrate that WT, but not cata-
lytically inactive mutant (H363Y) SirT1 deacetylates Pparg in
a NAD+-dependent manner. The SirT1 inhibitor nicotinamide
reversed the effect of SirT1 (Figures 1D and 1E). These findings
a Ligand-Dependent Manner
(A) Coimmunoprecipitation (coIP) of Flag-tagged
rosiglitazone treatment in 293 cells.
(B) CoIP of Flag-Pparg and SirT1 in 293 cells
treated with troglitazone (Trog) or GW9662 (GW)
(C) Pparg acetylation in 293 cells in response to
(D and E) In vitro Pparg deacetylation by SirT1.
Bovine serum albumin (BSA) is contained in the
(F) CoIP of Pparg with SirT1 in mouse adipose
tissues. We immunoprecipitated 2 mg of epidid-
ymal (eWAT) or inguinal fat lysates (iWAT) with
Pparg antibody H100, and blotted the immuno-
precipitates with SirT1 or Pparg (E8) antibodies.
(G) CoIP of SirT1 and Pparg with Flag M2 beads
using iWAT (5 mg) from SirBACO mice expressing
See also Figure S1.
1. SirT1 DeacetylatesPparg
reveal that SirT1 deacetylates Pparg.
Unlike other SirT1 substrates, Pparg
SirT1 in vivo.
Next we asked whether SirT1 interacts with Pparg in physio-
logic conditions. Accordingly, we detected SirT1 in Pparg
immunoprecipitates from epididymal and inguinal WAT (eWAT
and iWAT) lysates (Figure 1F). Taking advantage of Flag-tagged
SirT1 in SirT1 Bacterial Artificial Chromosome Overexpressor
transgenic mice (SirBACO) (Banks et al., 2008), we demon-
strated that Pparg could be detected by in vivo coimmunopreci-
pitation of Flag-SirT1 in iWAT (Figure 1G).
SirT1 Mimics the Effects of Pparg Ligands on White
and Brown Genes
Since both TZDs and SirT1 decrease Pparg acetylation, we
hypothesized that SirT1 gain-of-function phenocopies aspects
of Pparg activation by TZDs. A known function of TZDs is to
repress visceral WAT geneswhose induction isgenerally associ-
ated with insulin resistance (Vernochet et al., 2009). Activation
of SirT1 by resveratrol in 3T3-L1 white adipocytes had a similar
effect (Figure 2A). Overexpression of SirT1 did not affect
adipogenesis (Figures S2A–S2C), but selectively decreased
representative white genes Angiotensinogen (Agt), Resistin,
Wdnm1L, Chemerin, and Pank3. The catalytically inactive SirT1
mutant, H363Y, was unable to repress most white genes (Fig-
ure 2B). The inhibitory effect of SirT1 on these white genes is
consistent with its lipid mobilizing function in white adipocyte
(Picard et al., 2004). Next we compared the effects of SirT1
with those of Pparg agonists on brown genes in HIB-1B cells
(Tontonoz et al., 1994). We chose these cells because their
expression levels of Ucp1, SirT1 and its native inhibitor Deleted
in breast cancer-1 (Dbc1) (Kim et al., 2008; Zhao et al., 2008)
are more similar to subcutaneous WAT (iWAT) than those of
3T3-L1 white adipocytes (Figure S2D). Activating SirT1 with
resveratrol (Lagouge et al., 2006) or Pparg with troglitazone
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 621
increased Ucp1 while the SirT1 inhibitor nicotinamide mimicked
the Pparg antagonist GW9662 to repress it (Figure 2C). Overex-
pression of WT or H363Y mutant SirT1 did not inhibit cellular
differentiation (Figure S2E), but WT SirT1 decreased Pparg
acetylation and increased BAT markers Ucp1 and Dio2, while
the H363Y mutant had the opposite effect (Figures 2D and 2E).
These data indicate that SirT1 deacetylase activity is required
to promote brown gene expression and repress white genes in
a cell-autonomous manner that mimics liganded Pparg.
SirT1 Induces ‘‘Browning’’ of Subcutaneous WAT
In vivo, we observed a positive correlation of SirT1 levels with
brown gene expression in different mouse adipose tissues;
conversely, we observed a negative correlation of the SirT1
inhibitor Dbc1 with brown genes (Figures 3A and S3A). Based
on these findings, we probed the browning function of SirT1
in vivo by using three mouse models of altered SirT1 activity:
Sirt1 knockout mice (Sirt1?/?) (McBurney et al., 2003); mice
with increased SirT1 activity as a result of genetic deletion of
Dbc1 (Dbc1?/?) (Escande et al., 2010) and mice overexpressing
SirT1 (SirBACO) (Banks et al., 2008). In rare Sirt1?/?mice that
survived to adulthood (McBurney et al., 2003), we saw normal
levels of brown markers Ucp1 and C/ebpb in BAT (Figure S3B),
but lower levels in iWAT (Figure 3B), indicating that SirT1 is
required to maximize the thermogenic capacity of subcuta-
To validate our finding, we surveyed adipose tissues in
Dbc1?/?mice after triggering thermogenesis by overnight
cold exposure (4?C). We saw increased numbers of Ucp1-
immunoreactive paucilocular iWAT adipocytes, which are
usually unilocular (Figure 3C), together with increased expres-
sion of brown genes Ucp1 and C/ebpb, and suppression of
white genes, Chemerin and Resistin (Figures 3D and 3F),
suggestive of a ‘‘browning’’ of subcutaneous white adipocytes.
In contrast, knockout of Dbc1 had no effect on Ucp1 expres-
sion in BAT (Figures S3C and S3D). We observed a similar
Figure 2. SirT1 Mimics Pparg Ligand in Regulating Adipocyte Gene Expression
(A) Expression of white genes in 3T3-L1 adipocytes treated with 50 mM resveratrol or vehicle (Veh) overnight on differentiation day 7. *p < 0.05, **p < 0.01 versus
vehicle (n = 3).
(B) White gene expression in 3T3-L1 adipocytes overexpressing GFP, SirT1 or Sirt1-H363Y (HY). Fully differentiated cells were harvested on day 8 of differ-
entiation. *p < 0.05, **p < 0.01 versus GFP (n = 7).
(C) Ucp1 expression in HIB-1B brown adipocytes with treatments overnight on day 6 of differentiation. **p < 0.01 versus vehicle-treated controls (n = 3–4).
(D and E) Protein (D) and gene expression analysis(E) in HIB-1B cells overexpressing GFP, SirT1 and Sirt1-H363Y (HY). IP, immunoprecipitation; IB, immunoblot.
*p < 0.05, **p < 0.01 versus GFP (n = 3–9). Values are presented as means ± SEM.
See also Figure S2.
622 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
induction of Ucp1 and C/ebpb in visceral eWAT of Dbc1?/?
mice (Figure S3E) but, unlike in subcutaneous iWAT, we saw
no paucilocular adipocytes (Figure 3C). Owing to the extremely
low levels of Ucp1 in eWAT (Figures 3A and S3A), it’s unlikely
that its induction in this tissue contributes significantly to overall
thermogenesis, consistent with the observation that iWAT is
Figure 3. SirT1 Gain-of-Function Promotes Browning of Subcutaneous WAT
(A) Western blots in visceral (eWAT), subcutaneous (iWAT), and brown (BAT) adipose tissues from cold-exposed 8-week-old male mice.
(B) Western blotting of iWAT from chow-fed SirT1?/?mice and control littermates after overnight cold exposure.
(C–G) Eight-week-old, chow-fed male Dbc1?/?and SirBACO mice with their control littermates (WT) were exposed to 4?C overnight. H&E and Ucp1 immu-
nohistochemical staining of adipose tissues (C), western blotting (D and E), and gene expression analysis (F and G) of iWAT. *p < 0.05, **p < 0.01 for Dbc1?/?
versus WT (n = 6) or for SirBACO versus WT (n = 5). Values are presented as means ± SEM.
See also Figure S3.
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 623
Figure 4. Metabolic Correlates of SirT1-Dependent Pparg Deacetylation
(A and B) Time course (A) and area under the curve (AUC) (B) of IPGTT in chow-fed, 18-week-old male Dbc1?/?and WT mice before and after chronic cold
for Dbc1?/?versus cold-exposed Dbc1?/?mice. In (B) *p < 0.05. (n = 6–7).
(C and D) Ucp1 immunohistochemistry, H&E staining (C), and gene expression (D) in iWAT of 9-week-old male SirBACO mice after chronic cold exposure.
*p < 0.05, **p < 0.01 versus WT (n = 6).
624 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
the WAT depot most prone to browning (Kozak, 2010). If the
browning effect associated with Dbc1 knockout was mediated
by increased SirT1 activity, it should be phenocopied by SirT1
gain-of-function in SirBACO mice. Indeed, cold-exposed
SirBACO mice had an increased number of Ucp1-immunoreac-
tive paucilocular adipocytes in iWAT, as did Dbc1?/?mice (Fig-
ure 3C). The expression of brown markers, including Ucp1,
Dio2, and C/ebpb was increased in iWAT of SirBACO mice
(Figures 3E and 3G). Similar to Sirt1?/?and Dbc1?/?mice,
SirBACO mice had only marginally enhanced expression of
Ucp1 and C/ebpb in BAT (Figures S3F and S3G). Interestingly,
Cbp-deficient mice—which are known to be insulin-sensitive—
show similar browning phenotypes (Yamauchi et al., 2002).
These data indicate a browning function of SirT1 in iWAT,
rather than BAT.
SirT1 Gain-of-Function and Pparg Ligands Have
Overlapping Insulin-Sensitizing Effects
We next investigated the metabolic correlates of our findings.
Aging Dbc1?/?mice become glucose intolerant without body
weight changes (Figures 4A and S4A). We exploited this feature
to critically test the hypothesis that promoting BAT-like features
in WAT benefits insulin sensitivity. We exposed insulin-resistant
Dbc1?/?mice and controls to 12?C, a moderately low tempera-
ture. Unlike acute cold exposure to 4?C, this treatment did not
result in loss of body weight (Figure S4A). The 4-week treatment
had no effect in controls, but restored glucose tolerance in
Dbc1?/?mice (Figures 4A and 4B), with increased expression
of brown genes in iWAT (data not shown). In SirBACO mice,
despite increasedfood intake
increased energy expenditure. Similar to acute cold exposure,
we observed changes in iWAT, but not in BAT and eWAT (Fig-
ures S4D and S4E), associated with the appearance of
smaller, paucilocular or multilocular adipocytes with more
intense Ucp1 immunoreactivity (Figure 4C). Gene expression
studies demonstrated elevated levels of brown (Ucp1, Cox7a1,
and Cidea), angiogenic (Vegf), and lipolytic genes (Atgl) (Fig-
ure 4D)—all indicative of improved WAT function along with
WAT ‘‘insulin-resistance’’ genes were concordantly de-
creased in iWAT from SirBACO mice on high-fat diet (HFD)
nochet et al., 2009). As expected, Pparg acetylation was
decreased in SirBACO mice (Figure 4F), presumably mirroring
TZD’s in vitro effect. If TZDs and SirT1-dependent Pparg
deacetylation promoted browning through a shared molecular
mechanism, we would expect that the in vivo effects of TZDs
would be partly offset by SirT1 gain-of-function, to the extent
that they are both mediated by browning of adipose tissue.
This hypothesis can only be tested indirectly, as the systemic
effects of SirT1 and TZDs on insulin sensitivity are not limited
to browning of WAT, nor are they exclusively mediated by one
another (i.e., there are SirT1-independent effects of TZDs and
vice versa). Nonetheless, we investigated the ability of rosiglita-
zone to restore glucose tolerance in SirBACO and Dbc1?/?
mice rendered insulin-resistant with high-fat feeding. Indeed,
administration of rosiglitazone improved glucose tolerance in
high-fat-fed WT mice, but had only partial, statistically nonsignif-
icant effects in either SirBACO (Figures 4G, 4H, and S4F) or
Dbc1?/?mice (Figures 4I, 4J, and S4G). These findings provide
additional evidence for our hypothesis.
SirT1 Deacetylates Pparg on Lys268 and Lys293
We sought to identify SirT1-dependent deacetylation sites in
Pparg. We used HPLC/MS/MS analysis on trypsin- and chymo-
trypsin-digested peptides to identify five acetylated lysines at
residues 98, 107, 218, 268, and 293, with ?87% coverage of
Pparg sequence (Figures S5A and S5B). Among them, two evo-
lutionally conserved residues in the helix 2-helix 20region,
Lys268 and Lys293 (Figure S5C), were deacetylated following
rosiglitazone treatment (Figures 5A and 5B), suggesting that
they are SirT1 substrates. In the Pparg tertiary structure (Fig-
ure S5D), Lys268 juts out from the groove containing Ser273
(Lin et al., 2009; Nolte et al., 1998), a cyclin-dependent kinase
5 (CDK5) phosphorylation site (Choi et al., 2010), while Lys293
lies as a potential gatekeeper with its side chain facing
Ser273. The loop following the Pparg helix 20is highly flexible,
and responds with different conformational changes to Pparg
ligands with distinct transcriptional signatures (Waku et al.,
2009). Ligand binding induces a conformational change that
hinders access to the groove, burying Lys293 (Figures 5C and
S5E). It bears to reason that this event may require debulking
acetylated Lys293 through deacetylation. Unlike Lys293,
Lys291 anchors on the opposite side of helix 20, and upon ligand
binding its long side chain abuts on the outside of the groove
(Figures S5D and S5E). Consistently, mass spectroscopy anal-
ysis of Pparg peptides showed that it is not acetylated (data
Humans carrying the Pparg mutation P467L (Pro495 of murine
Pparg2) are severely insulin-resistant and diabetic (Barroso
et al., 1999). This amino acid change disrupts a conserved coac-
tivator-binding motif LxxLL (murine Pparg’s amino acids 493–
497) in helix 12 (Figure S5F). Interestingly, SirT1 interacts with
another transcription factor, FoxO1, through the latter’s cognate
LxxLL motif (Nakae et al., 2006). Thus, we asked whether the
P467L mutation interferes with Pparg binding to and deacetyla-
tion by SirT1. Protein interaction modeling localized binding of
(E and F) Expression of white genes (E) and Pparg acetylation (F) in iWAT of male SirBACO mice after 8 weeks on HFD. Mice were placed on HFD at 6 weeks of
age. *p < 0.05, **p < 0.01 versus WT (n = 5). IP, Immunoprecipitation; IB, immunoblot.
(G and H) IPGTT (G) and AUC (H) in male SirBACO mice fed with HFD for 6 weeks (starting at 12 weeks of age), and treated with rosiglitazone (rsg) or vehicle (veh)
for 1 week. Body weight information is in Figure S4F. *p < 0.05, **p < 0.01 WT-rsg versus WT-veh (n = 6–8).
(I and J) IPGTTs (I) and AUC (J) in male Dbc1?/?mice fed with HFD for 25 weeks before and after 1 week rosiglitazone (rsg) administration. Mice were placed on
HFD at 5 weeks of age. Body weight information is in Figure S4G. *p < 0.05, **p < 0.01 for WT versus WT-rsg (n = 7). Values are presented as means ± SEM.
See also Figure S4.
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 625
Figure 5. Identification of Pparg Lys268 and Lys293 as SirT1 Substrates
(A and B) Annotation of MS/MS spectra of acetylated peptides of Pparg after trypsin digestion at Lys268 (A) and at Lys293 (B).
(C) 3-D model of liganded and unliganded Pparg structure generated by NIH Cn3D software, localizing Lys268, Lys293, and Ser273 within helix 2-helix 20region.
(D) DecreasedSirT1 binding and increasedacetylation of PpargP467L mutant.CoIP ofFlag-tagged WT or P467L mutant Ppargwith SirT1 in293cells. Pparg(E8)
antibody fails to recognize P467L mutant.
(E) Mutation of the LxxLL motif on helix 12 disrupts SirT1 binding. CoIP of Flag-tagged WT or 2LA mutant Pparg with SirT1 in 293 cells.
(F) Mutations of Lys268 or Lys293 decrease acetylation of P467L mutant Pparg in 293 cells.
(G) Pparg acetylation in pooled iWAT from four to five male mice exposed to 4?C overnight or fed HFD for 16 weeks. We used 12 mg protein to immunoprecipitate
Pparg using antibody H100.
(H) Interaction of Pparg with SirT1 in iWAT. Pooled iWAT from three to five male SirBACO mice exposed to 4?C overnight or fed HFD for 16 weeks. We used 8 mg
protein as in each lane to coIP Flag-tagged SirT1.
(I) Ppargacetylationinresponse toTZD following HFD. Male WT mice werefed HFD for18weeksand treatedwithrosiglitazone for 3days. Pooled iWATfrom four
mice was lysed, and 12 mg protein was used in each lane to immunoprecipitate with acetyl-Lysine (Ac-K) antibody.
626 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
SirT1 to ligand-bound Pparg within the LxxLL motif (Figure S5G).
Indeed, P467L Pparg was hyperacetylated, and showed
reduced binding to SirT1 that failed to be rescued by SirT1 over-
expression (Figure 5D). Alanine mutations of the LxxLL motif
(L496A/L497A) also abolished the interaction of Pparg and
SirT1 (Figure 5E). Mutating Lys293 or Lys268 in the hyperacety-
lated Pparg mutant P467L dramatically decreased its acetyla-
tion, providing further evidence that both residues are sites of
Pparg acetylation (Figure 5F). Of interest, Pro495 (Pro467 in
human) and Lys293 are juxtaposed by a bridge between Ile295
and His494 and the aromatic side chain of Phe315 in response
to ligand binding (Waku et al., 2009). This conformation should
facilitate SirT1 binding to the LxxLL motif, providing ready
access to its substrates, Lys293 and the nearby Lys268.
Physiologic Regulation of Pparg Acetylation
To evaluate the significance of Pparg acetylation, we investi-
gated its regulation by physiological cues. Pparg acetylation
decreased in iWAT when brown genes were activated by cold
exposure (Figure 5G), while it increased with insulin resistance
brought about by HFD (Figure 5G). The increase of Pparg
acetylation following HFD was associated with reduced inter-
action with SirT1 (Figure 5H) and was reversed by rosiglitazone
administration (Figure 5I), Dbc1 knockout (Figure 5J) or SirT1
overexpression (Figure 4F). Moreover, activation of SIRT1 by
resveratrol decreased PPARg acetylation in human subcuta-
neous adipose tissue (Figure 5K), in agreement with the
insulin-sensitizing effects of resveratrol in humans (Timmers
et al., 2011). Total Pparg acetylation changed relatively little
in vivo, consistent with the mass spectrometry demonstration
that only two of five lysine residues are responsive to rosiglita-
zone (Figure S5B).
BAT-like Functions of Pparg Acetylation Site Mutants
We investigated the effects of Pparg deacetylation in adipo-
cytes by generating stable clones of Swiss-3T3 fibroblasts
expressing WT Pparg, or acetylation-mimetic K293Q (KQ),
deacetylation-mimetic K293R (KR) and K268R/K293R (2KR)
Pparg mutants. WT Pparg induced differentiation of Swiss
3T3 fibroblasts into adipocytes, as did KR, whereas KQ delayed
it (Figures 6A and S6A). Strikingly, the 2KR mutant was more
potent than WT Pparg in promoting lipid accumulation,
increasing expression of the insulin-sensitizing adipokine,
adiponectin, and promoting degradation of anti-adipogenic
b-catenin (Liu et al., 2006) (Figures 6A and S6A). These data
indicate that deacetylation is required for full Pparg activation.
Mitochondrial activity, as assessed by membrane potential,
was inhibited by KQ, but enhanced by 2KR (Figure 6C). These
findings were unrelated to mitochondrial biogenesis, assessed
by measuring mitochondrial mass (Figure 6C), mitochondrial
DNA, or mitochondrial protein content (Figures S6C and S6D).
2KR was more potent than WT to activate BAT genes including
Ucp1, Elovl3, Cidea, Cox7a1, and Pgc1a (Figures 6D and S6E).
In contrast, KQ selectively repressed expression of BAT genes
and induced expression of white genes without affecting
canonical Pparg targets (Figures 6D, 6E, S6E, and S6F).
Consistent with the induction of BAT genes, basal mitochondrial
respiration and maximal mitochondrial respiratory capacity
increased in cells expressing 2KR, as did oligomycin-depen-
dent (i.e., ATP synthesis inhibition-dependent) proton leak (Fig-
ure 6B). Another TZD-responsive gene, Fgf21 (Wang et al.,
2008), was likewise repressed by KQ and activated by 2KR (Fig-
ure 6D). In summary, these data indicate that SirT1-dependent
deacetylation of Pparg Lys268 and Lys293 mediates TZD-
induced browning of white adipocytes, enhancing expression
of characteristic BAT genes and repressing white ‘‘insulin resis-
SirT1-Dependent Deacetylation Modulates Pparg
Ser273 phosphorylation regulates Pparg activity (Choi et al.,
2010). Interestingly, Ser273 phosphorylation parallels acetyla-
tion of Lys293, but not of Lys268 (Figures S7A and S7B). Thus,
we investigated whether the transcriptional selectivity and meta-
bolic functions of Pparg deacetylation are mediated through
ligand-dependent Ser273 dephosphorylation. Dephosphoryla-
tion of Ser273 by TZD is required to induce adiponectin (Choi
et al., 2010). In Swiss 3T3 cells, stable overexpression of Pparg
S273A (SA) or S273A/2KR (AR) mutants resulted in overlapping
effects on adiponectin (Figure S7C). In contrast, the AR mutant
trumped the effect of the SA mutant by activating brown genes
(Figure S7D). These data indicate that Pparg acetylation and
Ser273 phosphorylation coordinately promote adipokine pro-
duction, while deacetylation is the main signal affecting brown
Ligand binding induces Pparg/Rxr dimerization and cofactor
exchange. We hypothesized that Pparg deacetylation might
dial in specificity of brown gene expression through coactiva-
tor/corepressor recruitment. Prdm16 is a determinant of the
to induce browning of subcutaneous WAT (Seale et al., 2011).
We observed that Prdm16 binding to Pparg was dependent on
rosiglitazone (Figure 7A, lanes 1–2). Pparg acetylation by Cbp
prevented Prdm16 binding, but rosiglitazone was able to partly
relieve the inhibition to an extent that was commensurate with
Pparg deacetylation (Figure 7A, lanes 3–4). Accordingly acetyla-
tion-defective Pparg 2KR showed constitutive, ligand-indepen-
dent binding to Prdm16 (Figure 7B). Likewise, Cbp blocked the
Pparg/Prdm16 interaction while SirT1 overexpression was able
to rescue it (Figure 7C), supporting a key role of deacetylation
in this process. SirT1 gain-of-function mimicked the effect of
Pparg ligand to promote Prdm16 binding while the Pparg
(J) Pparg acetylation in response to TZD and deletion of Dbc1. Chow-fed male WT mice were treated with rosiglitazone for 3 days. Twelve milligrams of protein
from pooled iWAT of four mice was used for IP with Pparg antibody (H100).
(K) PPARg acetylation in human adipose tissue in response to resveratrol. Human subcutaneous adipose fragments were treated with resveratrol (50 mM) for
12 hr; 1.2 mg protein was immunoprecipitated with PPARg antibody (H100).
We estimated relative levels of PPARg acetylation (Ac-Pparg) using densitometry with NIH ImageJ software.
See also Figure S5.
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 627
antagonist GW9662 blocked the effect of SirT1 (Figure 7D). In
matin immunoprecipitates from the Ucp1 enhancer (Chen et al.,
not its HY mutant (Figure 7E). In contrast, neither in vivo nor
in vitro did we detect any interaction between Prdm16 and
SirT1 (data not shown).
Site-directed mutagenesis of individual lysine residues
demonstrated that Lys293, but not Lys268, was critical to recruit
Prdm16 to Pparg (Figure 7F). Transcriptional corepressor NCoR
is an important component of the Pparg complex. Thus, we
investigated whether acetylation of these two lysines affected
NCoR binding. Intriguingly, mutating either lysine to arginine
Required for Its Browning Function
(A) Western blots analyses of Swiss 3T3 fibro-
blasts expressing WT, K293R (KR), K293Q (KQ),
and K268R/K293R (2KR) Pparg on day 6 of
(B) Oxygen consumption rates (OCR) in fully
differentiated cells (Day 8). *p < 0.05, **p < 0.01 for
2KR versus WT cells; #p < 0.05, ##p < 0.01 for KQ
versus WT, n = 6–7. The higher basal OCR in cells
expressing the KQ mutant is likely due to
increased cell number, owing to their shorter
doubling time (Figure S6B).
(C) Mitochondrial mass (green) and membrane
potential (red) on day 8 of differentiation.
(D and E) Brown gene (D) and white gene
expression(E) on day 8 in cells treated with vehicle
(Veh) or rosiglitazone (Rsg) overnight. *p < 0.05,
**p < 0.01 versus WT cells; #p < 0.05, ##p < 0.01
versus untreated cells (n = 3). Values are pre-
sented as means ± SEM.
See also Figure S6.
abolished the interaction of NCoR with
Pparg, suggesting that acetylation of
both residues is required for this interac-
tion (Figure 7F). Consistent with these
data, Cbp increased NCoR binding to
Pparg, while SirT1 prevented it (Fig-
ure 7G). Therefore, deacetylation of
Pparg on Lys293 is required to recruit co-
activator Prdm16, while deacetylation on
Lys268 and Lys293 is required to clear
corepressor NCoR. These data indicate
that the acetylation state of Lys268 and
Lys293 is critical for corepressor/coacti-
This study illustrates that SirT1 gain-of-
function induces BAT-like remodeling
of white adipocytes in vivo and in vitro
by deacetylating Pparg on Lys293 and
Lys268. We identify SirT1 as a Pparg
deacetylase, whose effects mimic biochemical, cell biological,
and physiological outcomes of ligand-dependent Pparg activa-
tion. Mechanistically, this process is associated with recruitment
of the brown cofactor Prdm16 and clearance of the corepressor
NCoR from the Pparg complex. We propose a model in which
the metabolic benefits of SirT1, the browning function of Pparg
ligands and their ability to selectively recruit coactivator
Prdm16 are subsumed under the unifying mechanism of Pparg
(Cohen et al., 2004) and deacetylates Pparg at Lys268 and
Lys293. Thus, we suggest that Pparg acetylation on Lys268
and Lys293 is a signal of plenty, as it favors lipid storage and
cell proliferation (Figure S6B). Conversely, deacetylation of
628 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
Figure 7. Deacetylation of Pparg Modulates the Coactivator/Corepressor Exchange
(A) Pparg interacts with Prdm16 in a ligand- and deacetylation-dependent manner. CoIP of Flag-tagged Pparg with Prdm16 in 293 cells, following Cbp trans-
fection or rosiglitazone treatment.
(B) Acetylation-defective Pparg increase binding of Prdm16. CoIP of Flag-tagged Pparg (WT) or 2KR mutant with Prdm16 in 293 cells.
(C) SirT1 promotes Pparg interaction with Prdm16. CoIP of Flag-tagged Pparg with Prdm16 following transfection of Cbp and/or SirT1 in 293 cells.
(D) SirT1 mimics TZD to increase Pparg interaction with Prdm16. CoIP of Flag-tagged Pparg with Prdm16 following overexpression of SirT1 and/or overnight
treatments in 293 cells.
(E) ChIP analysis of Pparg and Prdm16 binding to the Ucp1 promoter in HIB-1B adipocytes expressing GFP, WT, or H363Y (HY) mutant SirT1.
(F) Deacetylation of Pparg Lys268 or Lys293 inhibits binding of NCoR. CoIP of Flag-tagged Pparg (WT), K268R, or K293R mutants with exogenous Prdm16 and
endogenous NCoR in 293 cells.
(G) Acetylation of Pparg promotes binding of NCoR. CoIP of Flag-tagged Pparg with NCoR following transfection with SirT1 and Cbp in 293 cells.
(H) Model of SirT1-dependent Pparg deacetylation in energy homeostasis and insulin sensitivity. When nutrients are available, SirT1 is inactive and Pparg is
acetylated on Lys268 and Lys293. This condition favors lipid storage. During energy deprivation, SirT1 becomes active and is recruited to Pparg, possibly as
a result of ligand-induced conformational changes, to deacetylate Lys268 and Lys293. In white adipocytes, the deacetylated Pparg interacts with Prdm16 to
promote thermogenesis (energy expenditure) and improve insulin sensitivity.
See also Figure S7.
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 629
Pparg tilts the balance from energy storage to energy expendi-
ture (thermogenesis) and promotes insulin sensitivity.
Pparg Deacetylation in Browning WAT
Promoting brown features in white adipocytes can prevent
obesity and diabetes (Farmer, 2009). In the present study, we
propose a mechanism by which SirT1-mediated Pparg deacety-
lation might contribute to this process. The browning function
we did not detect any effect of SirT1 gain- or loss-of-function in
BAT in our mouse models. From a therapeutic standpoint, this is
consistent with the idea that BAT itself plays a modest role in
protecting from diet-induced weight gain, whereas converting
WAT into a less efficient energy storage site might. It also
suggests that browning of WAT occurs in a mechanistically
distinct fashion from BAT activation. Indeed, several factors
that promote WAT browning, such as Irisin (Bostro ¨m et al.,
2012), Fgf21 (Fisher et al., 2012), and Prdm16 (Seale et al.,
2011) do so independently of BAT.
Unlike the browning factors mentioned above, SirT1 does not
constitutively induce brown genes in iWAT at ambient tempera-
ture (data not shown), suggesting that hormonal or environ-
mental cues—such as cold exposure—are required to activate
tion on Lys268 and Lys293 could therefore be exploited as
readout to identify factors that modulate the browning process.
The ability to regulate browning of WAT should be viewed as
an essential component of any therapeutic approach that lever-
ages this mechanism, since ‘‘browned’’ cells are energy ineffi-
cient, and might have undesired effects if left unchecked.
The SirT1-dependent browning process occurs in a heteroge-
neous fashion within morphologically and anatomically homoge-
neous adipose depots. This might be related to sympathetic
innervation, a key modulator of BAT and Ucp1 induction in
WAT (Bartness et al., 2010). Or it is possible that the ability to
activate the brown program in response to Pparg ligand or
SirT1 gain-of-function is limited to a specialized subset of adipo-
cytes. Alternatively, since vascular endothelial cells have plurip-
otent potential and can differentiate into pre-adipocytes (Tran
et al., 2012), activation of SirT1 and Pparg deacetylation could
favor pre-adipocyte differentiation into brown (or ‘‘brite’’) adipo-
cytes. This possibility is unlikely to account for brown gene
induction in WAT following acute cold exposure, but might
underlie the chronic effect of cold exposure.
Integrating Acetylation with Other Posttranslational
Modalities of Pparg Regulation
Lysine residues can be modified by sumoylation and ubiquitina-
tion. Fgf21 (Dutchak et al., 2012) and TZDs (Pascual et al., 2005)
are insulin-sensitizers with browning properties (Fisher et al.,
2012; Vernochet et al., 2009) that promote Pparg sumoylation.
The Fgf21-dependent sumoylation site on Pparg, Lys107
(Dutchak et al., 2012), is strongly acetylated in our mass spec-
trometry analysis, but its acetylation is unaffected by rosiglita-
zone. It will be of interest to determine whether there exists
a reciprocal regulation of acetylation and sumoylation at this
site during the browning process. Ligand activation of Pparg
increases its ubiquitin-mediated degradation (Hauser et al.,
2000), a process that could also be regulated by deacetylation
of Lys268 and Lys293, because we observed that mutation of
these two residues increased Pparg stability (data not shown).
TZDs inhibit CDK5-dependent phosphorylation of Pparg
Ser273 (Choi et al., 2010), a residue buried within the groove
lined by Lys268 and Lys293. Consistent with prior results in
different systems (Qiang et al., 2010, 2011), we show that
Ser273 phosphorylation correlates with Lys293 acetylation, sug-
gesting that the acetylation state of these two residues affects
access of related kinases or phosphatases, to the phosphoryla-
tion site. We also found that the effects of acetylation and phos-
phorylationof theseresiduesonbrowngeneexpression diverge,
while those on the regulation of adiponectin overlap. Therefore,
acetylation of Pparg regulates its phosphorylation-mediated
function. In summary, acetylation might integrate energy status
with ligand availability to regulate protein stability and the effects
of sumoylation and phosphorylation on Pparg activity.
Following the introduction of biguanides in the clinical manage-
ment of diabetes in the 1940s, TZDs represent the only addition
to the pharmacopeia of insulin-sensitizers. But their use has
been tainted by liver toxicity, adverse effects on fluid balance,
body weight, bone turnover, and certain types of cancer, as
well as lingering if elusive links to cardiovascular risk (Rosen,
2010). In addition, the TZD saga has negatively affected devel-
opment of transcription factor-based therapeutics, even as
recognition of their homeostatic role has grown exponentially
(Kim-Muller and Accili, 2011). Our data do not amount to a
rehabilitation of TZDs for treatment of metabolic disorders, but
raise the interesting possibility that, by judicious use of SirT1
agonists in combination with appropriate Pparg ligands, it is
possible to render WAT less efficient, thus warding off the
adverse effects of TZD and paving the way for a reevaluation
of Pparg as a therapeutic target (Qiang and Accili, 2012).
SirBACO mice (C57BL/6J background), Dbc1?/?and Sirt1?/?mice (129/J 3
C57BL/6J background) were housed at 23 ± 1?C in a 12 hr light/dark cycle
with free access to normal chow (PicoLab rodent diet 20) (LabDiet 5053).
The Columbia University Animal Care and Utilization Committee approved
all procedures. The high-fat diet contained 60% calories from fat, 20% from
protein, and 20% from carbohydrates (Research Diet, D12492). For acute
cold exposure, we placed animals at 4?C for 16 hr, and for chronic cold
exposure we placed animals at 12?C on a 12-hr light/diet cycle for 4 weeks.
We injected rosiglitazone intraperitoneally (i.p.) daily at a dose of 10 mg/kg
body weight, performed IPGTT as described (Banks et al., 2008), and deter-
mined body compositions by NMR (Bruker Optics).
Plasmids and Cell Culture
We obtained pcDNA-Flag-Pparg2 (# 8895), pBabe-puro-Flag-HA-Pparg2
(# 8859) and pcDNA-Prdm16 (#15503) from Addgene, and generated Pparg
mutants K293R, K293Q, K268R, K268Q, 2KR (K268R/K293R), P467L,
P467L/K293R and P467L/K268Q, S273A, S273A/K268R/K293R (AR), and
L496A/L497A (2LA) by site-directed mutagenesis. In this study, Pparg refers
to the longer isoform Pparg2. We subcloned WT and H363Y mutant SirT1
cDNA into the BamHI site of pEGFP-N1 to generate N-terminal FLAG- and
C-terminal GFP-tagged SirT1. We cloned these constructs into plasmid
pBabe-blast, modified by replacing the puromycin-resistance gene with
630 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.
a blasticidin-resistance gene, to engineer retroviral vectors for derivation of
stably transduced clones as described (Vernochet et al., 2009). The final
concentration of puromycin or blasticidin for selection is 2.5 mg/ml. HA-Pparg
(Kitamura et al., 2005) and SirT1 adenoviruses have been described (Nakae
et al., 2003). For transient expression in HEK293T (293) cells, we transfected
pcDNA-Pparg or pEGFP-SirT1 using TransIT-LT1 reagent (Mirus Bio), or
transduced Pparg or SirT1 adenoviruses. We differentiated and maintained
3T3-L1 preadipocytes, Swiss-3T3 fibroblasts (both from ATCC), and HIB-1B
brown preadipocytes as described (Vernochet et al., 2009). We treated cells
overnight with resveratrol (10 mM), troglitazone (10 mM), rosiglitazone (5 mM),
nicotinamide (20 mM), or GW9662 (10 mM).
We extracted proteins and performed western blotting as described (Qiang
et al., 2010). Sources of antibodies are Dbc1 (Bethyl Laboratories); Ucp1
(Abcam); Adiponectin (Affinity BioReagents); SirT1 and b-catenin (Millipore);
Flag M2 and HA (Sigma-Aldrich); Prdm16 (R&D systems); monoclonal or
polyclonal acetyl-Lysine, aP2, Perilipin and phospho-CDK substrate (Cell
Signaling); actin, tubulin, C/ebpb, Pparg (E8), Pparg (H100), NCoR, and Cox
III (Santa Cruz).
Immunoprecipitation and In Vivo Pparg Acetylation
We performed coimmunoprecipitation (coIP) in Flag IP buffer (50 mM Tris [pH
7.9], 150 mM NaCl, 10% glycerol supplemented with protease inhibitors). For
immunoprecipitation (IP), we lysed cells into high-salt Flag IP buffer. To detect
salt Flag IP buffer and delipidated them by ultracentrifugation (40,000 rpm
for 2 hr at 4?C). We incubated lysates with antibody-conjugated agarose
beads overnight at 4?C, washed the beads four times and eluted the pre-
cipitates by Flag/HA peptides or boiling in nonreducing sample buffer. The
antibody-conjugated agarose beads are Flag M2 beads and monoclonal
Anti-HA (Sigma-Aldrich), Pparg (H100)-conjugated (Santa Cruz), and anti-
acetyl-Lysine agarose (ImmuneChem).
Gene Expression Analysis
We isolated RNA with RNeasy Lipid Tissue kit (QIAGEN) and DNase I diges-
tion, synthesized cDNA with High-capacity cDNA Reverse Transcription kit
(Applied Biosystems), and performed quantitative real-time PCR (Q-PCR)
with goTaq qPCR Master Mix (Promega) in a Bio-Rad CFX96 Real-Time
PCR system. We calculated relative gene expression levels by DDCt method
using cyclophilin A as internal control. We list primer sequences in Table S1.
We used paraffin-embedded sections for H&E, and Ucp1 immunohistochem-
istry(Abcam,1:100 dilution). Weassessed adipocyte lipid content by Oilred-O
We used unpaired 2-tailed Student’s t tests to evaluate statistical significance
and p < 0.05 to declare a statistically significant change. We present all values
as means ± standard error of means (SEM).
figures, and one table and can be found with this article online at http://dx.doi.
We thank K. Tsuchiya and L. Ozcan for help with different aspects of this work,
T. Kolar and A. Flete for technical support, Q. Zhang for help with structural
analysis, U. Pajvani and G. Heinrich for comments on the manuscript, and
members of the Accili laboratory for discussion of the data. We thank
B. Spiegelman for providing HIB-1B cells. This work was supported by NIH
grants HL087123, DK58282, DK64773, DK063608 (Columbia Diabetes
Research Center), and RR024156 (Columbia University CTSA).
Received: September 19, 2011
Revised: February 19, 2012
Accepted: June 7, 2012
Published: August 2, 2012
Banks, A.S., Kon, N., Knight, C., Matsumoto, M., Gutie ´rrez-Jua ´rez, R., Ros-
setti, L., Gu, W., and Accili, D. (2008). SirT1 gain of function increases energy
efficiency and prevents diabetes in mice. Cell Metab. 8, 333–341.
Barroso, I., Gurnell, M., Crowley, V.E., Agostini, M., Schwabe, J.W., Soos,
M.A., Maslen, G.L., Williams, T.D., Lewis, H., Schafer, A.J., et al. (1999). Domi-
nant negative mutations inhuman PPARgamma associated withsevere insulin
resistance, diabetes mellitus and hypertension. Nature 402, 880–883.
Bartness, T.J., Vaughan, C.H., and Song, C.K. (2010). Sympathetic and
Bostro ¨m, P., Wu, J., Jedrychowski, M.P., Korde, A., Ye, L., Lo, J.C., Rasbach,
K.A., Bostro ¨m, E.A., Choi, J.H., Long, J.Z., et al. (2012). A PGC1-a-dependent
myokine that drives brown-fat-like development of white fat and thermogene-
sis. Nature 481, 463–468.
Canto ´,C.,Gerhart-Hines, Z.,Feige, J.N.,Lagouge,M.,Noriega,L.,Milne,J.C.,
Elliott, P.J., Puigserver, P., and Auwerx, J. (2009). AMPK regulates energy
expenditure by modulating NAD+ metabolism and SIRT1 activity. Nature
Cederberg, A., Grønning, L.M., Ahre ´n, B., Taske ´n, K., Carlsson, P., and
Enerba ¨ck, S. (2001). FOXC2 is a winged helix gene that counteracts obesity,
hypertriglyceridemia, and diet-induced insulin resistance. Cell 106, 563–573.
Chen, W., Yang, Q., and Roeder, R.G. (2009). Dynamic interactions and coop-
erative functions of PGC-1alpha and MED1 in TRalpha-mediated activation of
the brown-fat-specific UCP-1 gene. Mol. Cell 35, 755–768.
Choi,J.H.,Banks,A.S.,Estall,J.L.,Kajimura, S.,Bostro ¨m,P.,Laznik,D.,Ruas,
J.L., Chalmers, M.J., Kamenecka, T.M., Blu ¨her, M., et al. (2010). Anti-diabetic
drugs inhibit obesity-linked phosphorylation of PPARgamma by Cdk5. Nature
Cohen, H.Y., Miller, C., Bitterman, K.J., Wall, N.R., Hekking, B., Kessler, B.,
Howitz, K.T., Gorospe, M., de Cabo, R., and Sinclair, D.A. (2004). Calorie
restriction promotes mammalian cell survival by inducing the SIRT1 deacety-
lase. Science 305, 390–392.
Dutchak, P.A., Katafuchi, T., Bookout, A.L., Choi, J.H., Yu, R.T., Mangelsdorf,
D.J., and Kliewer, S.A. (2012). Fibroblast growth factor-21 regulates PPARg
activity and the antidiabetic actions of thiazolidinediones. Cell 148, 556–567.
Escande, C., Chini, C.C., Nin, V., Dykhouse, K.M., Novak, C.M., Levine, J., van
Deursen, J., Gores, G.J., Chen, J., Lou, Z., and Chini, E.N. (2010). Deleted
in breast cancer-1 regulates SIRT1 activity and contributes to high-fat diet-
induced liver steatosis in mice. J. Clin. Invest. 120, 545–558.
Farmer, S.R. (2009). Obesity: Be cool, lose weight. Nature 458, 839–840.
Fisher, F.M.,Kleiner,S.,Douris,N., Fox, E.C.,Mepani,R.J., Verdeguer, F.,Wu,
J., Kharitonenkov, A., Flier, J.S., Maratos-Flier, E., and Spiegelman, B.M.
(2012). FGF21 regulates PGC-1a and browning of white adipose tissues in
adaptive thermogenesis. Genes Dev. 26, 271–281.
Hauser, S., Adelmant, G., Sarraf, P., Wright, H.M., Mueller, E., and Spiegel-
man, B.M. (2000). Degradation of the peroxisome proliferator-activated
receptor gamma is linked to ligand-dependent activation. J. Biol. Chem.
Kim, J.E., Chen, J., and Lou, Z. (2008). DBC1 is a negative regulator of SIRT1.
Nature 451, 583–586.
Kim-Muller, J.Y., and Accili, D. (2011). Cell biology. Selective insulin sensi-
tizers. Science 331, 1529–1531.
Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc. 631
Kitamura, Y.I., Kitamura, T., Kruse, J.P., Raum, J.C., Stein, R., Gu, W., and Download full-text
Accili, D. (2005). FoxO1 protects against pancreatic beta cell failure through
NeuroD and MafA induction. Cell Metab. 2, 153–163.
Kozak, L.P. (2010). Brown fat and the myth of diet-induced thermogenesis.
Cell Metab. 11, 263–267.
Lagouge, M., Argmann, C., Gerhart-Hines, Z., Meziane, H., Lerin, C., Daussin,
F., Messadeq, N., Milne, J., Lambert, P., Elliott, P., et al. (2006). Resveratrol
improves mitochondrial function and protects against metabolic disease by
activating SIRT1 and PGC-1alpha. Cell 127, 1109–1122.
Lin, Y.Y., Lu, J.Y., Zhang, J., Walter, W., Dang, W., Wan, J., Tao, S.C., Qian, J.,
Zhao, Y., Boeke, J.D., et al. (2009). Protein acetylation microarray reveals that
NuA4 controls key metabolic target regulating gluconeogenesis. Cell 136,
Liu, J., Wang, H., Zuo, Y., and Farmer, S.R. (2006). Functional interaction
between peroxisome proliferator-activated receptor gamma and beta-
catenin. Mol. Cell. Biol. 26, 5827–5837.
McBurney, M.W., Yang, X., Jardine, K., Hixon, M., Boekelheide, K., Webb,
J.R., Lansdorp, P.M., and Lemieux, M. (2003). The mammalian SIR2alpha
protein has a role in embryogenesis and gametogenesis. Mol. Cell. Biol. 23,
Milne, J.C., Lambert, P.D., Schenk, S., Carney, D.P., Smith, J.J., Gagne, D.J.,
Jin, L., Boss, O., Perni, R.B., Vu, C.B., et al. (2007). Small molecule activators
of SIRT1 as therapeutics for the treatment of type 2 diabetes. Nature 450,
Nakae, J., Kitamura, T., Kitamura, Y., Biggs, W.H., III, Arden, K.C., and Accili,
D. (2003). The forkhead transcription factor Foxo1 regulates adipocyte differ-
entiation. Dev. Cell 4, 119–129.
Nakae, J., Cao, Y., Daitoku, H., Fukamizu, A., Ogawa, W., Yano, Y., and
Hayashi, Y. (2006). The LXXLL motif of murine forkhead transcription factor
FoxO1 mediates Sirt1-dependent transcriptional activity. J. Clin. Invest. 116,
Nolte, R.T., Wisely, G.B., Westin, S., Cobb, J.E., Lambert, M.H., Kurokawa, R.,
Rosenfeld, M.G., Willson, T.M., Glass, C.K., and Milburn, M.V. (1998). Ligand
binding and co-activator assembly of the peroxisome proliferator-activated
receptor-gamma. Nature 395, 137–143.
Pascual, G., Fong, A.L., Ogawa, S., Gamliel, A., Li, A.C., Perissi, V., Rose,
D.W., Willson, T.M., Rosenfeld, M.G., and Glass, C.K. (2005). A SUMOyla-
tion-dependent pathway mediates transrepression of inflammatory response
genes by PPAR-gamma. Nature 437, 759–763.
Picard, F., Ge ´hin, M., Annicotte, J., Rocchi, S., Champy, M.F., O’Malley, B.W.,
Chambon, P., and Auwerx, J. (2002). SRC-1 and TIF2 control energy balance
between white and brown adipose tissues. Cell 111, 931–941.
Picard, F., Kurtev, M., Chung, N., Topark-Ngarm, A., Senawong, T., Machado
De Oliveira, R., Leid, M., McBurney, M.W., and Guarente, L. (2004). Sirt1
promotes fat mobilization in white adipocytes by repressing PPAR-gamma.
Nature 429, 771–776.
A., Tang, X., Straubhaar, J., Cherniack, A.D., Parker, M.G., and Czech, M.P.
(2006). Suppression of oxidative metabolism and mitochondrial biogenesis
by the transcriptional corepressor RIP140 in mouse adipocytes.J. Clin. Invest.
Qiang, L., and Accili, D. (2012). FGF21 and the second coming of PPARg. Cell
Qiang, L., Banks, A.S., and Accili, D. (2010). Uncoupling of acetylation from
phosphorylation regulatesFoxO1 function independent of itssubcellularlocal-
ization. J. Biol. Chem. 285, 27396–27401.
Qiang, L., Lin, H.V., Kim-Muller, J.Y., Welch, C.L., Gu, W., and Accili, D. (2011).
Proatherogenic abnormalities of lipid metabolism in SirT1 transgenic mice are
mediated through Creb deacetylation. Cell Metab. 14, 758–767.
Ravussin, E., and Galgani, J.E. (2011). The implication of brown adipose tissue
for humans. Annu. Rev. Nutr. 31, 33–47.
Rosen, C.J. (2010). Revisiting the rosiglitazone story—lessons learned. N.
Engl. J. Med. 363, 803–806.
Scime `,A.,Grenier, G.,Huh,M.S.,Gillespie,M.A.,Bevilacqua,L.,Harper, M.E.,
and Rudnicki, M.A. (2005). Rb and p107 regulate preadipocyte differentiation
into white versus brown fat through repression of PGC-1alpha. Cell Metab. 2,
Seale, P., Bjork, B., Yang, W., Kajimura, S., Chin, S., Kuang, S., Scime `, A.,
Devarakonda, S., Conroe, H.M., Erdjument-Bromage, H., et al. (2008).
PRDM16 controls a brown fat/skeletal muscle switch. Nature 454, 961–967.
Seale, P., Conroe, H.M., Estall, J., Kajimura, S., Frontini, A., Ishibashi, J.,
Cohen, P., Cinti, S., and Spiegelman, B.M. (2011). Prdm16 determines the
thermogenic program of subcutaneous white adipose tissue in mice. J. Clin.
Invest. 121, 96–105.
Timmers, S., Konings, E., Bilet, L., Houtkooper, R.H., van de Weijer, T., Goos-
sens, G.H., Hoeks, J., van der Krieken, S., Ryu, D., Kersten, S., et al. (2011).
Calorie restriction-like effects of 30 days of resveratrol supplementation on
energy metabolism and metabolic profile in obese humans. Cell Metab. 14,
Tontonoz, P., Graves, R.A., Budavari, A.I., Erdjument-Bromage, H., Lui, M.,
Hu, E., Tempst, P., and Spiegelman, B.M. (1994). Adipocyte-specific tran-
scription factor ARF6 is a heterodimeric complex of two nuclear hormone
receptors, PPAR gamma and RXR alpha. Nucleic Acids Res. 22, 5628–5634.
Tran, K.V., Gealekman, O., Frontini, A., Zingaretti, M.C., Morroni, M., Gior-
dano, A., Smorlesi, A., Perugini, J., De Matteis, R., Sbarbati, A., et al. (2012).
The vascular endothelium of the adipose tissue gives rise to both white and
brown fat cells. Cell Metab. 15, 222–229.
Tsukiyama-Kohara, K., Poulin, F., Kohara, M., DeMaria, C.T., Cheng, A., Wu,
Z., Gingras, A.C., Katsume, A., Elchebly, M., Spiegelman, B.M., et al. (2001).
Adipose tissue reduction in mice lacking the translational inhibitor 4E-BP1.
Nat. Med. 7, 1128–1132.
Vernochet, C., Peres, S.B., Davis, K.E., McDonald, M.E., Qiang, L., Wang, H.,
Scherer, P.E., and Farmer, S.R. (2009). C/EBPalpha and the corepressors
CtBP1 and CtBP2 regulate repression of select visceral white adipose genes
during induction of the brown phenotype in white adipocytes by peroxisome
proliferator-activated receptor gamma agonists. Mol. Cell. Biol. 29, 4714–
Virtanen, K.A., and Nuutila, P. (2011). Brown adipose tissue in humans. Curr.
Opin. Lipidol. 22, 49–54.
Waku, T., Shiraki, T., Oyama, T., Fujimoto, Y., Maebara, K., Kamiya, N., Jin-
gami, H., and Morikawa, K. (2009). Structural insight into PPARgamma
activation through covalent modification with endogenous fatty acids. J.
Mol. Biol. 385, 188–199.
Wang, H., Qiang, L., and Farmer, S.R. (2008). Identification of a domain within
peroxisome proliferator-activated receptor gamma regulating expression of
a group of genes containing fibroblast growth factor 21 that are selectively
repressed by SIRT1 in adipocytes. Mol. Cell. Biol. 28, 188–200.
Wang, Y.C., McPherson, K.,Marsh,T.,Gortmaker,S.L.,and Brown, M.(2011).
Health and economic burden of the projected obesity trends in the USA and
the UK. Lancet 378, 815–825.
Yamauchi, T., Oike, Y., Kamon, J., Waki, H., Komeda, K., Tsuchida, A., Date,
Y., Li, M.X., Miki, H., Akanuma, Y., et al. (2002). Increased insulin sensitivity
despite lipodystrophy in Crebbp heterozygous mice. Nat. Genet. 30, 221–226.
regulation of the deacetylase SIRT1 by DBC1. Nature 451, 587–590.
632 Cell 150, 620–632, August 3, 2012 ª2012 Elsevier Inc.