REVIEW 3939
Development 140, 3939-3949 (2013) doi:10.1242/dev.080549
© 2013. Published by The Company of Biologists Ltd
The developmental origins of adipose tissue
Daniel C. Berry1, Drew Stenesen1, Daniel Zeve1 and Jonathan M. Graff1,2,*
Adipose tissue is formed at stereotypic times and locations in a
diverse array of organisms. Once formed, the tissue is dynamic,
responding to homeostatic and external cues and capable of a
15-fold expansion. The formation and maintenance of adipose
tissue is essential to many biological processes and when
perturbed leads to significant diseases. Despite this basic and
clinical significance, understanding of the developmental biology
of adipose tissue has languished. In this Review, we highlight
recent efforts to unveil adipose developmental cues, adipose
stem cell biology and the regulators of adipose tissue
homeostasis and dynamism.
Key words: Adipose, Adipocyte, Development, Stem cells, Niche,
Vascular niche
Introduction: fat facts
Adipose tissue plays a myriad of roles; it serves as a central nexus
of metabolic communication and control, an arbiter of
thermoregulation, a buffer against trauma and the cold, and a
regulator of reproduction and satiety. Fat is also associated with
emotionally charged issues, imparting various psychosocial
imprints that have changed over the centuries: from a sign of wealth
in the middle ages, to a Rubenesque celebration during the
Renaissance, to fear and loathing in modern Hollywood (unless of
course it is injected into the lips or other ‘cosmetically desirable’
locations). Despite the current social aversion to fat, the increase in
the percentage of people who are overweight has been marked and
is observed in nearly all regions of the world. The global prevalence
of obesity is ~15% and in the United States more than two thirds
of the population is overweight (Hossain et al., 2007). The health
care system is reeling not only from an obesity epidemic, but also
from a host of obesity-related negative sequelae including diabetes,
cardiovascular disease, cirrhosis and even cancer. The reasons for
this increase are well known – increased food consumption and
decreased exercise – but this is only because the body is
predisposed to fat accumulation.
What is also becoming clear is that a variety of evolutionary and
developmental forces have laid the foundation for the development
of obesity. Both invertebrates and vertebrates have fat-storing
tissues and many aspects of the mechanistic underpinnings are
conserved. These include the developmental programs,
transcriptional cascades and basic proteins that regulate fat
synthesis, storage and lipolysis (McKay et al., 2003; Suh et al.,
2006; Suh et al., 2008; Suh et al., 2007). It appears that fat tissues
evolved primarily as a safe harbor to store energy in times of plenty
and to provide fuel when food sources become insufficient.
However, in addition to serving as purely a storage depot, adipose
1
Department of Developmental Biology, University of Texas Southwestern Medical
Center, Dallas, TX 75390, USA. 2Department of Molecular Biology, University of
Texas Southwestern Medical Center, Dallas, TX 75390, USA.
*Author for correspondence ([email protected])
tissue is now recognized as the body’s largest endocrine organ,
controlling many aspects of systemic physiology by secreting
hormones (adipokines), lipids, cytokines and other factors (Gesta
et al., 2007; Nawrocki and Scherer, 2004; Spiegelman and Flier,
2001). Although many of the regulatory molecules are not yet
identified, they control a wide variety of biological actions,
including appetite, glucose homeostasis, insulin sensitivity, aging,
fertility and fecundity, and body temperature.
In mammals, adipose tissue forms in utero, in the peripartum
period and throughout life. Notably, even in adult humans new
adipocytes are generated continually and at substantial rates
(Spalding et al., 2008). Adipose tissue is composed of adipose stem
cells (the precursor cells that give rise to new adipocytes),
adipocytes (the fat-storing cells) and various other cell types, which
include mural, endothelial and neuronal cells. Adipose tissue
comprises various discrete depots, such as inguinal, interscapular,
perigonadal, retroperitoneal and mesenteric depots, which are
placed in defined positions throughout the body. These various
depots develop at specific and distinguishable pre- and postnatal
times and they have discrete and distinct morphologies (Gregoire
et al., 1998; Hossain et al., 2007; MacDougald and Mandrup,
2002). These observations support the notion that developmental
cues are pivotal for the proper formation, migration and
morphology of adipose tissue. Yet relatively little is known about
adipocyte development because adipocytes are spread throughout
the body, because of inherent difficulties in handling the enormous
and fragile adipose cells, and because of historic reliance on cell
culture models.
However, a new horizon is in sight; developmental tools are
beginning to be utilized and interest in adipose tissue development
is expanding. This recent tectonic shift towards a developmental
field has begun to improve our understanding of adipose
developmental biology and has positioned the field for extensive
advancement. This is timely, as the obesity crisis has increased the
demand for new therapeutic strategies for adipose-related illnesses.
Thus, there is an urgent need to understand better the mechanisms
and molecules that control the formation of adipocytes and the
expansion of adipose tissue due to caloric excess. Here, we review
the recent advancements in adipose tissue development, the
discovery and regulation of adipose stem cells, adipocyte turnover,
and potential therapeutic interventions for those suffering from
obesity and diabetes.
Brown and white fat: flavors of the month
In mammals, there are two main classes of adipose tissue that are
separated into major histological divisions, are molecularly distinct
and are functionally distinguishable. White adipose tissue (WAT)
is designed for energy storage whereas brown adipose tissue (BAT)
dissipates energy and generates heat. WAT is far more prevalent
and more biologically relevant, whereas BAT primarily functions
in the neonatal period, although recent reports have shown BAT to
be present and have functional roles in adults (Cypess et al., 2013;
Lidell et al., 2013). Adipose tissue is also present in a variety of
locations, such as palms, soles, scalp, periarticular regions and
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orbits, that are thought to have so called mechanical roles (Gesta et
al., 2007). In addition, adipocytes are positioned throughout the
skin and bone marrow, and recent evidence indicates that in those
locations adipocytes regulate stem cell biology, for example by
controlling epidermal and hematopoietic lineage decisions.
White adipose tissue: the great white whale
White adipose tissues coordinate systemic metabolism. This key
role is highlighted by the metabolic dysfunctions (e.g.
hyperglycemia, hyperlipidemia, hypertension, diabetes, liver
disease, increased carcinogenesis, etc.) that are a consequence of
too many (obesity) or too few (lipodystrophy) white adipocytes.
There are two main divisions of white adipose tissue: subcutaneous
and visceral. Subcutaneous and visceral depots are simply
separated based upon gross anatomical location (Fig. 1A,B). Yet
this gross placement appears to define a variety of essential
features, including developmental timing, microscopic appearance,
molecular signature and biological function. In rodents,
subcutaneous depots, such as interscapular and inguinal, appear to
form prior to visceral depots, which include retroperitoneal,
perigonadal and mesenteric WAT (Fig. 1C,D). Each adipose depot
has a specific, distinct and reproducible morphology and texture.
Histologically, subcutaneous fat is heterogeneous and contains
A
A
Human
A
Vis
B
Vis
P
P
Lean
A
Mouse
Vis
SQ
Vis
P
SQ
SQ
Obese
A
SQ
C
Fig. 1. The anatomical distributions of fat.
(A,B) Coronal MRI of a lean (left) and an obese
(right) human (A) and mouse (B); yellow lines
delineate subcutaneous and visceral depots. A,
anterior; P, posterior; SQ, subcutaneous; Vis,
visceral. (C,D) Physical orientation, distribution
and location of subcutaneous and visceral
depots. Subcutaneous fat (C) is located in the
inguinal (IGW) and interscapular (ISCW) regions.
Visceral fat (D) is positioned at perigonadal
(PGW), retroperitoneal (RPW) and mesenteric
(MWAT) regions of the body. Yellow arrows
indicate the denoted depot location within the
body of the mouse.
Obese
Lean
SQ
mature unilocular adipocytes intercalated with small multilocular
adipocytes, whereas visceral fat is more uniform and appears to
consist primarily of large unilocular adipocytes (Fig. 2A,B)
(Tchernof et al., 2006; Tchkonia et al., 2007). Of note,
retroperitoneal adipose depots, which are placed within the visceral
class, have features that may be intermediate between subcutaneous
and visceral depots. Current thinking indicates that subcutaneous
and visceral depots are likely to have different contributions to
metabolism, with inherent health ramifications. Increased
subcutaneous fat deposition, sometimes called a pear-shaped or
female pattern of distribution, might protect against certain aspects
of metabolic dysfunction (Snijder et al., 2003a; Snijder et al.,
2003b). However, visceral depots, also known as an apple or male
pattern, are thought to be associated with metabolic complications
and appear to increase the risk of diabetes, hyperlipidemia and
cardiovascular disease (Grauer et al., 1984). Because of these still
ill-defined associations, it has become popular to term
subcutaneous adipose as ‘good fat’ and visceral as ‘bad fat’.
What might account for these associations? It is plausible that
the aforementioned histological differences may be at play.
Subcutaneous depots contain an abundance of small multilocular
adipocytes, which might be protective against metabolic
derangement (Salans et al., 1973; Weyer et al., 2000). In addition,
P
D
Visceral
IGW
PGW
RPW
MWAT
Development
ISCW
A
B
SQ
C
Visceral
Brown adipose
Fig. 2. Heterogeneity of fat. (A-C) Adipose depots are positioned at
stereotypic times and locations with defined sizes; shown are
subcutaneous (SQ) fat depots (A), perigonadal (visceral) fat depots (B) and
intrascapular brown adipose tissue (C) from three unrelated mice.
Histologically, white adipose depots differ in adipocyte size;
subcutaneous depots are more heterogeneous, containing both uni- and
multilocular adipocytes and interstitial tissue, whereas visceral depots are
more uniform and contain mostly unilocular adipocytes. Morphologically,
brown adipose tissue is very distinct from white adipose tissue with
uniformity of multilocular lipid droplets.
there is an increase in interstitial tissue in subcutaneous depots
(Fig. 2A). In conjunction with histological differences,
subcutaneous depots have increased rates of adipose turnover and
new adipocyte formation, and it is postulated that ‘younger’
adipocytes do not confer metabolic dysfunction (Björntorp et al.,
1971; Salans et al., 1973). Lipolytic rates also appear to be different
between the two depots (Fisher et al., 2002). Other factors that
might account for the differences include blood flow and proximity
to metabolically important visceral organs, such as the liver, which
produces glucose, lipids and cholesterols, and the intestine where
food is absorbed. It is also known that aspects of regional fat
deposition are inherited; for instance, it is widely appreciated that
the propensity to accumulate subcutaneous versus visceral fat runs
within families (Shi and Clegg, 2009). That is, one can only control
whether one becomes fat, not where one puts it.
Subcutaneous and visceral adipose depots also respond to
external stimuli in different ways. For example, in humans and
mice, steroid hormones confer depot-specific consequences (Shi
and Clegg, 2009); estrogen is thought to increase subcutaneous
depots, whereas adipose tissues within the neck, or within the
viscera, are more responsive to glucocorticoids. Furthermore,
subcutaneous and visceral depots display unique and
distinguishable responses to thiazolidinediones (TZDs), a class of
drugs widely prescribed for diabetes (Tontonoz and Spiegelman,
2008). Additional support for complexity within the lineages
derives from the clinical observation that lipodystrophy
(pathological absence of fat) preferentially affects different adipose
locations. For example, some patients with an inherited
lipodystrophy lack all metabolic fat but retain normal amounts of
mechanical fat (Herbst, 2012).
Transplantation methodologies have been employed in an
attempt to probe whether the functional differences between
visceral and subcutaneous depots might be intrinsic (autonomous)
or extrinsic (non-autonomous). However, these studies have not yet
produced a definitive answer, with some indicating inherent
differences and others suggesting that these functional attributes
alter when locations change. For example, Kahn and colleagues
performed an elegant series of murine reciprocal transplants
coupled with extensive metabolic analyses (Tran et al., 2008). The
studies indicated that transplantation of subcutaneous depots into
visceral locations improved metabolic parameters, whereas
transplantation of visceral depots into subcutaneous locations did
not (Tran et al., 2008). In addition to metabolic and morphological
changes, a striking rearrangement in gene expression towards a
subcutaneous signature occurred within three weeks of
transplantation (Satoor et al., 2011). Thus, there appear to be both
intrinsic and extrinsic forces that regulate adipose depot biology.
The factors and relative contributions of each are likely to have
fundamental and clinical significance and are areas of future
exploration.
A variety of studies support the notion that distinct and definable
molecular programs might underlie the functional differences
between subcutaneous and visceral adipose tissues (Gesta et al.,
2007). For example, gene expression profiling of adipose tissues in
rodents and humans demonstrated divergent and discrete molecular
signatures between the various depot locations. Notably, the pattern
of gene expression, even within a depot, changes depending on
adipocyte size (Jernås et al., 2006). Subcutaneous fat depots tend
to express higher levels of leptin, angiotensinogen and glycogen
synthase compared with visceral fat. Omental fat, part of the
visceral compartment, expresses increased levels of the insulin
receptor, 11β hydroxysteroid dehydrogenase (11β HSD) and
interleukin 6 (IL6) (Gesta et al., 2007; Masuzaki et al., 2001).
Large-scale gene expression analyses comparing rodent
subcutaneous and visceral adipose progenitor signatures also
identified a host of differentially expressed transcripts (Gesta et al.,
2006). These include a group of developmental regulators such as
homeobox (HOX) and forkhead box (FOX) family transcription
factors (Gesta et al., 2006; Macotela et al., 2012). Moreover, the
gene trends that were detected in mice were also observed in
human adipose samples (Gesta et al., 2006). Additional data
suggest that a subset of these genes correlate with body mass index
(BMI) and by extension relate to metabolic dysfunction and
cardiovascular disease (Gesta et al., 2006). The role of these genes
in fat formation and regulation remains unclear. However, many of
the identified genes are involved in developmental patterning, and
it is possible that they might regulate aspects of adipose tissue
formation. Studies on developmental signaling cascades have also
highlighted the notion that developmental forces are important to
adipose development and to regional distinctions. This can be
illustrated by the WNT pathway, which regulates adipose tissue
formation in a spatially significant manner (Longo et al., 2004;
Zeve et al., 2012). In relation to gene changes, fat-storing cells
derived from subcutaneous progenitors accumulate more lipid and
express higher levels of PPARγ and C/EBPα upon differentiation
compared with visceral progenitor cells (Baglioni et al., 2012;
Tchkonia et al., 2002). Subcutaneous progenitor cells also have
improved growth rate and different electrophysiological properties
(Baglioni et al., 2012; Macotela et al., 2012). Collectively, there
appear to be intrinsic genetic depot-specific differences in adipose
stem cells that result in different adipogenic potential, genetic
expression patterns, growth rates and biological properties. Thus,
these studies support the notion that distinct developmental cues
are likely to determine the complexity of the adipose lineage and
that all fat stem cells and adipocytes are not created equally.
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Brown adipose tissue: burning down the house
Brown adipocytes convert nutrients into chemical energy in the form
of heat (Kajimura et al., 2010). The primary role of BAT appears to
be in the neonatal period when temperature control is challenged
(Gregoire et al., 1998; MacDougald and Mandrup, 2002; Spiegelman
and Flier, 2001). Brown fat cells express a unique thermogenic and
mitochondrial genetic program that promotes mitochondrial
biogenesis, energy uncoupling and energy dissipation, which
provides essential heat to the organism (Kajimura et al., 2010).
Energy dissipation is accomplished by an abundance of
mitochondria, hence the eponymous color of BAT, as well as by
specialized proteins, including uncoupling protein 1 (UCP1). UCP1
collapses the electron gradient to generate heat rather than ATP
(Cannon and Nedergaard, 2004). Similar to WAT, BAT is present at
several stereotypic places and can be increased or decreased by
environmental cues (Fig. 2C). However, the brown adipocytes that
are present during neonatal life appear distinct from those that are
present, or can be induced, in adults (Wu et al., 2012).
In the neonatal period, the most prominent BAT depot is located
at the dorsal region between the scapulae (Cannon and Nedergaard,
2004). The cells in the interscapular region of the body are derived
from the dermomytome, an unusual tripotent engrailed 1 (EN1)positive cell lineage, which gives rise to brown fat bundles, the
dermis and muscle cells (Atit et al., 2006). Studies also indicate
that this specialized region of the body descends from a myogenic
factor 5 (MYF5)-positive source, but it is clear that only a subset
of brown adipocytes do so. Interestingly, recent data indicate that
a subset of white adipocytes also arises from MYF5-positive cells
(Sanchez-Gurmaches et al., 2012), and additional studies show that
brown and white fat progenitor cells share commonality.
Granneman and colleagues demonstrated that a platelet-derived
growth factor receptor alpha (PDGFRα)-positive lineage traces to
brown and white adipose tissue and that PDGFRα is present in
proliferating brown and white adipose progenitors (Lee et al.,
2012).
Brown adipocytes were once thought to be absent in adult
humans; however, recent data indicate that some adults have
energy-burning adipocytes with ‘brown-like’ characteristics, and
promotion of their formation or maintenance has potential as an
anti-obesity avenue (Cypess et al., 2009; Cypess et al., 2013; Lidell
et al., 2013; Whittle et al., 2011). In small-scale studies, it appears
that obesity reduces the appearance of brown-like cells and,
because of the therapeutic possibilities, a variety of studies have
been directed towards elucidating the molecular cascades and
factors underlying brown fat formation, with the aim of triggering
cells to have BAT-like energy-consuming actions (Cannon and
Nedergaard, 2004; Kajimura et al., 2010; Seale et al., 2008; Seale
et al., 2009). Many initial studies examined possible conservation
of function between the white and brown lineages, and revealed
that aspects of the overlapping core transcriptional machinery play
related roles in the two lineages (Kajimura et al., 2010). In addition,
various enzymes involved in lipogenesis and lipolysis are
expressed in both types of adipocytes. However, it also appears that
brown fat formation has unique aspects. For instance, bone
morphogenetic protein 7 (BMP7) can induce cultured cells to
undergo a brown adipocyte differentiation response (Tseng et al.,
2008). In vivo studies also highlight the potential role of BMP7 in
the formation or maintenance of BAT; Bmp7 null embryos have
less brown fat compared with controls and virally overexpressing
BMP7 increases brown but not white adipose tissue (Tseng et al.,
2008). Thus, it appears that there are distinct signals that promote
brown fat formation over white adipocyte formation and vice versa.
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In vitro adipocyte differentiation: artificial
sweeteners
Pioneering studies on in vivo formation of adipose tissue were
conducted in the first half of the 20th century but these were
scattered and primarily observational (Clark and Clark, 1940;
Napolitano, 1963; Napolitano and Gagne, 1963). Over the last few
decades, the major focus of the field has been cell culture modeling
primarily studying in vitro adipogenesis. The term adipogenesis
first described the transition of cultured and confluent 3T3-L1
fibroblast cells to individual lipid-laden cells upon incubation in a
powerful cocktail composed of artificial ‘inducers’: cAMP
inducers, glucocorticoid agonists and insulin or insulin-like growth
factor (IGF) (Green and Kehinde, 1975; Gregoire et al., 1998).
These fat-storing cells appeared to have several characteristics of
adipocytes and have served as the primary model. This wellstudied system has led to the elucidation of the cellular and
biological events that occur and the transcriptional hierarchy that
exists during the conversion of these cells from fibroblasts to
adipocyte-like cells (Rosen and Spiegelman, 2000; Tontonoz and
Spiegelman, 2008). It appears that aspects of the 3T3-L1
adipogenic transition also occur in other in vitro systems, such as
mouse embryonic fibroblasts (MEFs) and stromal vascular (SV)
cells (see below), when induced with the adipogenic induction mix
(Rosen and Spiegelman, 2000).
A significant advance derived from the 3T3-L1 cellular model
was the identification of relevant molecules expressed during fat
accumulation, most notably a transcriptional hierarchy. PPARγ, a
nuclear hormone receptor, is the centerpiece of this transcriptional
cascade and is necessary and sufficient for in vitro adipogenesis
(Chawla et al., 1994; Tontonoz et al., 1994). PPARγ controls genes
involved in lipid storage, lipid synthesis and glucose sensing
(Barak et al., 1999; He et al., 2003; Miles et al., 2000). Multiple
studies using conditional and hypomorphic alleles, all solidify the
role of PPARγ in differentiation of mature adipocytes. However, a
variety of data indicate that PPARγ might have significant roles in
addition to those in adipocytes and indeed may play central roles
in early adipose lineage development. For example, PPARγ is
expressed by embryonic day (E)14.5, much earlier than adipocyte
tissue development begins, and appears to mark the location of
subcutaneous fat depots that are present perinatally (Barak et al.,
1999; Brun et al., 1996). Furthermore, PPARγ is expressed in
adipose stem cells and appears to have key functions in the adipose
stem cell compartment, including roles in stem cell proliferation,
self-renewal, and cell determination (Tang et al., 2011; Tang et al.,
2008). Collectively, a variety of studies suggest that PPARγ is
crucial both for adipocyte terminal differentiation and as a
determinant that controls adipose tissue lineage development and
location.
Elegant in vitro studies have also identified many additional genes
that may be expressed in adipocytes or might regulate adipocyte
differentiation and function. Multiple signaling molecules can initiate
the cell model adipogenic cascade, including insulin, thyroid
hormone, glucocorticoids and TGFβ superfamily members
(BMP2/BMP4/GDF3) (Chapman et al., 1985; Flores-Delgado et al.,
1987; Zamani and Brown, 2011). During the initiation of
differentiation, a transcriptional cascade occurs with activation of
multiple factors, such as KLF4, KLF5, SMAD1/5/8, CREB,
KROX20 (EGR2), glucocorticoid receptor, STAT5A, STAT5B,
C/EBPβ and C/EBPδ. In turn, these factors can alter PPARγ
expression (Cristancho and Lazar, 2011; Siersbæk et al., 2012). Other
transcription factors, such as thyroid receptor, C/EBPα, SREBP-1c
(SREBF1), KLF15 and LXR (NR1H), are involved later in
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adipocyte maturation and regulate the expression of PPARγ as well
as that of other genes involved in fat storage (Siersbæk et al., 2012).
Terminal differentiation is characterized by the appearance of lipid
droplets and the expression of lipid/carbohydrate-storage proteins
[e.g. AP2 (FABP4), CD36, LPL, perilipin and GLUT4 (SLC2A4)]
and adipokines (leptin) (Cawthorn et al., 2012). Multiple
developmental signaling pathways, hormones and environmental
cues also inhibit in vitro adipogenesis, such as canonical WNT
signaling, retinoic acid, TNFα, Hedgehog (Hh) signaling,
SMAD/TGFβ and HIF1α (Berry et al., 2012; Cawthorn et al., 2007;
Ross et al., 2000; Suh et al., 2006; Sul, 2009; Wang et al., 2006). In
summary, studies in cell culture models indicate that the formation
of fat-storing cells is controlled by complex interactions of
stimulatory and inhibitory signals. A key challenge for the field,
however, is to determine whether these molecules have roles in
adipose tissue formation in vivo.
Adipose stem cells: getting to the root of the stem
Although some aspects of in vitro models reflect in vivo events, it
has become increasingly clear that in vivo adipocyte differentiation
is distinct. Therefore, the identification and characterization of the
adipose stem cell is essential to understanding adipose tissue
development, formation and maintenance. However, our
understanding of the developmental biology of adipose tissue has
lagged and our understanding of adipose stem cells is primitive.
Stem cells are crucial components for tissue development and
maintenance, and the notion of adipose progenitors stems from the
1940s. In the 1940s, it was noted that fibroblast-like cells change
morphology and acquire unilocular lipid droplets (Clark and Clark,
1940). The developmental or embryonic origin of adipose stem
cells remains largely unexplored and a variety of basic insights,
such as germ layer derivation, remain murky. Conventional
wisdom holds that WAT originates from the mesoderm germ cell
layer despite a lack of direct evidence supporting this hypothesis
(Duan et al., 2007); however, not all adipose tissue originates from
the mesoderm (Monteiro et al., 2009). For instance, a variety of
studies, such as those using chick-quail chimeras and modern Credependent cell marking studies in the mouse, indicate that
craniofacial adipose depots originate from neural crest cells (Billon
et al., 2007; Le Lièvre and Le Douarin, 1975).
Identifying and characterizing adipose stem cells
Recent efforts have begun to tease apart the endogenous adipose
stem/progenitor compartment. Tissue dissociation studies have
A
B
implicated the adipose stromal vascular fraction (SVF) as a
possible site of origin for adipose stem cells. The SVF is a
heterogeneous mixture of cells that can be isolated from adipose
tissue after dissociation and centrifugation, in which adipocytes
float and the remaining cells pellet (Hollenberg and Vost, 1968).
The SVF is composed of a myriad of cells, which includes
fibroblasts, endothelial cells, hematopoietic cells and neural cells.
Yet within this multiplicity of cell types the SVF appears to contain
bone fide adipose stem cells that proliferate, self-renew and
differentiate (Rodeheffer et al., 2008; Tang et al., 2008; Yamamoto
et al., 2007). The isolation, identification and characterization of
such adipose stem cells has only recently been performed using
two different approaches: tissue dissociation coupled with
fluorescence-activated cell sorting (FACS) and lineage tracing.
Utilizing cell surface markers, Friedman and colleagues identified
a CD24+ cell population within the SVF that had high adipogenic
potential in vitro (Rodeheffer et al., 2008). When this cell
population was injected into wild-type mice it did not form a
functional fat pad; however, when transplanted into A-Zip mice (an
engineered lipodystrophic model that lacks fat) the cells were able
to form adipose-like tissue. Thus, it appears that the A-Zip mice,
and possibly lipodystrophic mice in general, harbor a proadipogenic environment presumably containing various signals that
induce adipocyte differentiation (Rodeheffer et al., 2008).
Contemporaneously, Tang et al. also developed a flow-based
method for isolating adipose stem cells that could form adipose
tissue after transplantation, even in wild-type mice (Tang et al.,
2008). More importantly, by exploiting the observation that PPARγ
is expressed at low levels in adipocyte stem cells in order to label
genetically and lineage trace adipocyte stem cells, they reported
that PPARγ-expressing cells are present prior to birth and long
before adipocytes are formed (Tang et al., 2008). Moreover, these
cells proliferate, self-renew, differentiate and are capable of
repopulating the stem cell pool and contributing to the formation
of all adipocytes (Tang et al., 2011; Tang et al., 2008; Zeve et al.,
2009).
Remarkably, a new notion arose from these studies, that these
adipose stem cells are present in a vascular niche, residing as a
subset of perivascular mural cells within adipose depots that
express a variety of mural cell markers, such as PDGFRβ, αsmooth muscle actin and NG2 (CSPG4) (Fig. 3). Transplantation
of PDGFRβ mural cells, isolated from adipose depots, reconstituted
adipose depots in the recipients; yet equivalent PDGFRβ-positive
mural cells from other organs did not give rise to adipocytes or a
C
D
Tissue section
SVP
Fig. 3. From the vascular niche to adipocyte. (A) Illustration conceptualizing the adipose vascular niche depicting blood vessels (red), adipose stem
cells (green with green nuclei), mural cells (green with black nuclei) and mature adipocytes (yellow with green nuclei). (B) Mouse subcutaneous adipose
tissue stained with Hematoxylin and Eosin (H&E) stain. (C) Immunofluorescence image of mouse subcutaneous adipose tissue section showing the
expression of GFP (marking adipose stem cells, green), PECAM (an endothelial marker, red) and smooth muscle actin (SMA) (a mural cell marker, blue).
(D) Stromal vascular particulates (SVPs) were isolated from mouse subcutaneous fat depots and immunostained for GFP (adipose stem cell, green),
PECAM (endothelial marker, red) and SMA (mural cell marker, blue). Yellow arrows indicate adipose stem cells (green) residing at the vascular interface.
Development
GFP/PECAM/SMA
H&E
fat pad (Tang et al., 2008). These data support the notion that
adipose stem cells reside at the vascular interface and resemble
mural cells that can proliferate and differentiate into mature
adipocytes.
Some recent observations raise the possibility that endothelial
cells and hematopoietic stem cells might give rise to a subset of
adipocytes by changing fate into mural cells and then converting
to a progenitor population (Crossno et al., 2006; Gupta et al., 2012;
Medici et al., 2010; Sera et al., 2009; Tran et al., 2012). Such
lineage changes do not have significant precedence in other
endothelial compartments but may turn out to be important in
adipose biology. However, endothelial cell deletion of PPARγ did
not alter fat formation, under normal chow conditions, as would be
expected if endothelial cells are major components of the adipose
fate (Kanda et al., 2009). Recently, data indicate that neither
endothelial nor hematopoietic lineage markers label adipocytes
even under high fat diet conditions (Berry and Rodeheffer, 2013).
Factors that regulate adipose stem cells
The observation that PPARγ marks adipocyte stem cells that reside
in the perivascular niche raises several important questions about
the functional role of PPARγ within these cells. TZDs, a group of
glucose-lowering drugs prescribed for people with type 2 diabetes,
are PPARγ agonists that promote insulin sensitivity and lower
blood glucose. However, a common and distressing side effect for
patients treated with TZDs is significant weight gain and increased
fat mass (de Souza et al., 2001; Hiragun et al., 1988; Sandouk et
al., 1993a; Sandouk et al., 1993b). This process is multifaceted in
that TZDs stimulate new adipocyte formation and apoptosis of
large adipocytes (Okuno et al., 1998), cause fat deposition
remodeling from visceral to subcutaneous (Akazawa et al., 2000;
Kelly et al., 1999), increase fluid retention (Muto et al., 2001) and
augment appetite (Shimizu et al., 1998). This clinical observation
coupled with the finding that PPARγ marks the adipose stem cell
raised the possibility that TZDs might alter adipocyte stem cells,
for example by inducing them to differentiate thereby increasing
the number of adipocytes and leading to weight gain (Yki-Järvinen,
2004). Consistent with this hypothesis, in vivo administration of
TZDs to mice enhanced differentiation of the adipose stem cell
population and increased formation of additional mature adipocytes
(Tang et al., 2011). Furthermore, rosiglitazone (a TZD and
clinically prescribed medicine) enhanced adipose stem cell
proliferation and reduced the fraction of adipose stem cells in the
quiescent state. Intriguingly, chronic rosiglitazone administration
reduced the number of adipose stem cells, reduced the adipogenic
potential of the extant stem cell compartment and appeared to
exhaust the adipocyte stem population (Tang et al., 2011). These
studies indirectly implicated PPARγ in adipose stem cell
proliferation, self-renewal, lineage decisions, stem and niche
identity and adipocyte differentiation (Tang et al., 2011). Studies
examining these notions are a major area of future experimentation.
The adipose stem cell niche: blood brothers
The niche, the microenvironment in which stem cells reside, is
highly specialized and controls stem cell behaviors, such as
proliferation, quiescence and differentiation. In the 1940s, Clark
and Clark examined fat formation in rabbit ears using a chambered
system to displace the tissue and watch capillary growth and fat
formation into the chamber (Clark and Clark, 1940). The chamber
methodology allowed ‘real-time’ visualization of fat formation in
a quasi in vivo setting. These innovative experiments also indicated
that numerous minute lipid droplets appeared first and coalesced to
Development 140 (19)
form a large unilocular droplet as the cell morphology changed
(Clark and Clark, 1940). Electron micrographs from the 1960s
provided further evidence that adipocyte progenitor cells reside at
the blood vessel and ‘peel’ away as they begin the transition to
mature adipocytes (Napolitano, 1963; Napolitano and Gagne,
1963). This notion was further supported by Hausman and
colleagues who observed the appearance of vascular structures that
slightly protrude before the formation of adipocytes (Crandall et
al., 1997). Their microscopic studies also indicated that
angiogenesis recruits adipose stem cells and stimulates these stem
cells to differentiate (Crandall et al., 1997). These observations
were supported by studies of Han et al., whose data suggested that
angiogenesis precedes visceral epididymal adipose tissue depot
development (Han et al., 2011). This relationship between vessels
and adipose lineage cells appears to be reciprocal, as adipocyte
stem cells stimulate blood vessel formation. For instance, in vivo
mixing of adipose stem cells with endothelial cells showed that
adipose stem cells stimulate vasculogenesis and indicate that
adipocyte differentiation and angiogenesis occur in concert
(Traktuev et al., 2009). Furthermore, adipose stem cells secrete
multiple angiogenic regulators, some of which are transcriptionally
regulated by PPARγ, including vascular endothelial growth factor
(VEGF), angiopoietin-like 4, fibroblast growth factor 2 and matrix
metalloproteinases (MMPs) (Cao, 2007; Fukumura et al., 2003;
Gealekman et al., 2008; Kersten et al., 2000). Moreover, treating
adipose tissue fragments with TZDs increased angiogenic sprout
formation and similar results were obtained in vivo in mouse
(Gealekman et al., 2008). Inhibiting or activating angiogenic
factors controls adiposity; in the epididymal fat depot, blunting
VEGF/VEGFR2 signaling inhibits fat pad formation (Han et al.,
2011), and transgenically overexpressing VEGF in mature
adipocytes enhances vasculogenesis and reduces adipocyte size
(Nishimura et al., 2007; Sun et al., 2012), demonstrating the
orchestration between blood vessels and adipocytes.
Although the niche is known to be a major element of stem cell
control, examination of the niche has been hindered by difficulties
in assessing it as an intact structural unit. The ability to study the
adipose stem cell vascular niche has been aided by procedures that
maintain the native structure of the microenvironment, thus
allowing the isolation of stromal vascular particulates (SVPs) as an
organotypic culture system (Tang et al., 2011; Tang et al., 2008).
Isolation of SVPs showed that these vessels do not contain lipidfilled adipocytes but rather adipose stem cells that wrap around the
blood vessel and express a bevy of mural cell markers (Fig. 3C,D).
Upon adipogenic signals, such as TZDs, the stem cells on the SVP
transition to lipid-filled adipocytes loosely associated with the
particulate (Tang et al., 2008). Taken together, it seems likely that
the vascular network is the niche for adipocyte stem cells, and that
this microenvironment provides adipogenic cues to inhibit or
initiate adipocyte differentiation. The identification of such cues is
an area that may attract significant attention.
Adipocyte turnover: churning the fat
Stem cells play important roles in tissue development as well as in
homeostatic maintenance and response to environmental stimuli.
There are two distinguishable responses to adipose tissue
expansion: hypertrophy and hyperplasia. However, the level of
contributions of the two responses varies depending upon genetic
background, modifier effects, diet, biological and hormonal milieu,
and depot preference for fat storage. Adipose tissue can expand
from 2-3% to 60-70% of body weight in response to positive
energy balance (Fig. 1A,B; Fig. 4D) (Hossain et al., 2007). A
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Development 140 (19)
A
B
SQ
Visceral
SQ
Visceral
GFP
SQ
P2
C
P30
–Dox (P2-P30)
SQ
P30 P90
D
Visceral
P180
P30
P90
+Dox (P2-P30)
P180
SQ
Chow
Visceral
HFD
Chow
Fig. 4. Fat expansion. (A) Isolated postnatal
day (P)2 and P30 subcutaneous (inguinal) fat
depots, demonstrating the extensive
expansion that occurs within this short
window of time. (B) Adipo-trak mice (Tang
et al., 2008) were treated with placebo
(–Dox) or doxycycline (+Dox) at denoted
times. In the absence of doxycycline the GFP
marker is expressed and the adipose depots
are green. In the presence of doxycycline, all
GFP signal is lost, indicating that there was a
massive recruitment of new stem cells and
adipocyte replacement. (C) White adipose
depots enlarge with age; subcutaneous and
visceral fat depots were isolated from 30-,
90- and 180-day-old mice. Depot weight
nearly quintupled between 30 and 180 days.
(D) Subcutaneous inguinal (SQ) and visceral
perigonadal depots obtained from mice on
a standard diet (chow) or a high fat diet
(HFD), demonstrating the dramatic
expansion in response to HFD.
HFD
hypertrophic response is characterized by pre-existing adipocytes
enhancing triglyceride storage; estimates indicate that a single
adipocyte can increase its volume two- to threefold (Hirsch and
Batchelor, 1976; Salans et al., 1973). A prolonged hypertrophic
response appears to be a causal link between adipose expansion
and metabolic dysfunction such as reducing adipocyte insulin
sensitivity (Hossain et al., 2007; MacDougald and Mandrup, 2002).
During obesogenesis, there is significantly higher adipocyte
turnover, shortened adipocyte lifespan and a markedly increased
rate of apoptosis (Strissel et al., 2007). Hypertrophy also promotes
local inflammation, for example by recruiting ‘unhealthy’
macrophages to adipose tissue thereby facilitating adipocyte cell
death and other events that trigger maladaptive metabolic
consequences (Osborn and Olefsky, 2012). Furthermore,
hypertrophy provokes increases in several adipocyte extracellular
signals, such as IGF1, IGFBP and TNFα, that are linked to negative
outcomes. Of note, these factors appear able to alter various aspects
of adipose stem cell biology, such as proliferation, quiescence and
differentiation (Cao, 2007).
Adipose tissue can also expand by forming new adipocytes,
and newly formed, small adipocytes might be protective against
metabolic dysregulation (de Souza et al., 2001; Okuno et al.,
1998; Strissel et al., 2007). In the 1960s, it was shown in rodents
that cells within adipose depots proliferate, that adipocytes are
renewed and that adipocytes have a finite lifespan (Hellman and
Hellerstrom, 1961; Hollenberg and Vost, 1968). Indeed, caloric
excess stimulates the adipose stem cell compartment to proliferate
and differentiate. For example, Joe et al. reported a significant
increase in bromodeoxyuridine (BrdU) incorporation in the SVF
of subcutaneous adipose depots in response to high-fat diet,
whereas visceral fat increased as a result of hypertrophy (Joe et
al., 2009). Other studies have begun to examine the source(s) of
high-fat diet-induced increases in proliferation observed in the
adipose lineage. Several studies have since aimed to gain a better
understanding of adipocyte turnover but have yielded mixed
findings. In humans, it was found that 8.4% of adipocytes
turnover every year (Spalding et al., 2008). However, estimates
indicate that young adult mice generate ~15% of adipocytes each
month. The stem cell compartment also appears to be active with
some estimates suggesting that 4.8% of mouse adipose stem cells
are replicating at a given time (Rigamonti et al., 2011). These
findings suggest that adipose stem cell proliferation not only
contributes to adipocyte formation but also replenishes the stem
cell pool. Support for this notion comes from the findings that
during postnatal development [specifically, the first 30 days of
life (P0-P30 in mice)], there is rapid turnover and replenishing
events in the adipose depot (Fig. 4A,B) (Tang et al., 2008).
Furthermore, adipose tissue maintains its ability to expand as the
organism ages, and aging is associated with increased percentage
of body fat, which cannot only be attributed to hypertrophy
(Fig. 4C). Further studies to examine adipose stem cell
proliferation are needed to define the mechanisms that control
these events, how the stem cell divisions regulate adipose biology
and whether interventions directed towards cycle cell
manipulation are therapeutically tenable.
Adipose stem cells presumably confer the ability of adipose
depots to turnover, and they may also underlie the ability of
adipose tissue to regenerate. In humans, lipectomy and liposuction
are commonly performed medical procedures that remove excess
adipose tissue. However, removal of adipose tissue causes other fat
depots to remodel with both hypertrophic and hyperplastic
responses (Mauer et al., 2001; Reyne et al., 1983), and full
regeneration of the removed fat pad also occurs leading to repeat
operations. Experiments using rodent model systems have aimed
to characterize this regenerative phenomenon. When inguinal fat
(subcutaneous) depots were removed from rats the fat pad was
regenerated within ~13 weeks (Larson and Anderson, 1978).
Lipectomies in obese mice stimulated the rate of local and global
lipogenesis and stimulated adipocyte differentiation (Bueno et al.,
2011). This indicates that stem cells are potential sources of the
renewal, respond to environmental cues and provide experimental
paradigms for further exploration.
Development
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Development 140 (19)
the cause of traditional transplant failure (Han et al., 2011; Satoor
et al., 2011; Tran et al., 2008).
The ability to induce adipose stem cells into a variety of
lineages, such as skeletal myocytes for muscle degeneration,
cardiac myocytes for a failing heart, or neural fates for Alzheimer’s
disease, also has significant potential. Confirmation and
identification of the conditions necessary to promote lineage
variation might lead to production of cells or tissues amenable for
autologous transplant of modified tissue to treat a host of
degenerative diseases. Yet the full range of lineage
capacity/plasticity of isolated adipose progenitor cells remains
unknown. For instance, altering canonical WNT signaling (via a
conditional allele of β-catenin) in adipose stem cells disrupted
adipose development in mice (Fig. 5) (Zeve et al., 2009). These
mutant mice had a marked paucity of adipocytes, and a
lipodystrophic phenotype, with ensuing hypertriglyceridemia.
Notably, these mutants displayed essentially a complete loss of
subcutaneous adipose tissue accompanied by a lineage change in
the subcutaneous adipose stem cells. Of note, the secretome of
WNT mutant stem cells was altered and the cells produced high
levels of a glucose-lowering hormone, glucodyne, which had many
actions similar to insulin but functioned through distinct
mechanisms (Zeve et al., 2012).
The therapeutic adipose stem cell
The ability to isolate and track the responses of adipocyte stem
cells has provided a platform for the characterization and
manipulation of these cells, with the aim of providing some
beneficial therapeutic outcomes. Recent studies show that TZDs
improve symptoms of the metabolic syndrome in part by
mobilizing adipocyte progenitors to differentiate into new insulinsensitive adipocytes (Tang et al., 2011). This indicates that adipose
stem cells are pharmacologically accessible, that increased
formation of new adipocytes may be beneficial and that adipose
stem cells may be subject to additional types of manipulation.
Efforts to control differentiation or proliferation might thus serve
as weight loss intervention for overweight or pre-obese individuals.
Alternatively, other methods directed at inducing adipocyte
differentiation in order to maintain new small insulin-sensitive
adipocytes (healthy adipocytes) might provide metabolic relief to
those suffering with obesity and metabolic dysfunction. In addition
to pharmacological intervention, adipose stem cells are uniquely
poised for regenerative and genetic therapy. WAT provides
minimally invasive access to large numbers of progenitor cells
necessary for cell-based therapeutics. Furthermore, rejection is not
a concern because these cells can be harvested from and
administered to the same person. In principle, applications could
include reprogramming cells into other lineages or inducing cells
into the adipose lineage, for example to improve wound healing or
to use as a ‘filler’ to repair defects or other cosmetic concerns.
Plastic and reconstructive surgeons have long been plagued with
the inconsistencies of adipose grafting in wound healing and soft
tissue augmentation (Sterodimas et al., 2012). However, new
technologies enriching graft material for adipose progenitor cells
are being developed and show initial promise (Gir et al., 2012). An
issue in surgical applications is that transplants of adipose depots
tend to fail and necrose because of the lack of vascular supply.
However, transplanted adipose stem cells recruit vessel formation
and the formed adipose depots are well vascularized overcoming
A
Conclusions
In recent decades, attention to adipose tissue has increased in
parallel with a rising epidemic of obesity and its negative effects
on whole body metabolism and increased incidence of various
illnesses and conditions. Only recently have research efforts shifted
to understanding the developmental biology of this tissue (Han et
al., 2011; Rodeheffer et al., 2008; Tang et al., 2011; Tang et al.,
2008). The complexity of the various adipose lineages (white,
brown, induced brown, subcutaneous, visceral, etc.), the difficulty
in working with such a fragile cell type, and the non-contiguous
nature of this tissue has made it difficult to understand its
B
WNT-activated fibrotic tissue
Fig. 5. Manipulating the adipose stem cell: kill the stem, kill the tree. (A) Illustration conceptualizing the importance of adipose stem cells to the
formation of adipose tissue. This illustration demonstrates the extensive branching of the vascular network and shows that adipose stem cells (green
nuclei) and mural cells (black nuclei) reside on the vasculature. Adipose stem cells then transition to a multilocular progenitor giving rise to the
unilocular adipocyte. Inset: Histological image of subcutaneous fat from a wild-type mouse. (B) Disrupting adipose stem cells by activating WNT
signaling results in a fate change that promotes the formation of fibrotic tissue (purple). Inset: Histological image of subcutaneous fat from a mouse
with activated WNT signaling.
Development
Wild-type adipose tissue
SQ
SQ
WNT
W
NT activation
acttivattion
developmental origin, and multiple origins might exist. However,
with the generation of developmental tools, such as lineage tracing,
researchers are now poised to understand the developmental cues
and origin of adipose tissue and to answer a slew of interesting and
undefined questions. For instance, what are the cell types of origin
of the adipose lineage? What is the timing of adipose lineage
determination and specification? Do adipose stem cells arise in situ
on the blood vessel or do they migrate and arrive from elsewhere?
What are the signals derived from the blood vessel niche that
stimulate these stem cells to proliferate and differentiate or that
hold them in the quiescent state? What is the importance of adipose
stem cells to the homeostasis and maintenance of the fat pad under
both normal energy intake and excess nutrient load? Do other antidiabetes drugs alter adipose stem cell biology, similar to TZD
treatment? Do growth factors and development signaling pathways
alter stem cell behavior and adipocyte formation? In the midst of
the obesity epidemic, the recent discoveries and the answers to
these open-ended questions would provide hope that there is light
at the end of the tunnel.
Acknowledgements
We thank all current and former members of the Graff lab for their helpful
comments and discussion points leading to refinement of our concepts. We
especially thank Dr Ayoung Jo and Kia Huang for providing data images for
specific figures, and Dr Todd Soesbe for MRI imaging.We apologize for the
editorial constraints that limit the ability to cite a vast number of papers that
pioneered the way for the above studies and notions for future advances.
Funding
J.M.G. is supported by the National Institute of Diabetes and Digestive and
Kidney Diseases and the National Institutes of Health [R01-DK066556, R01DK064261 and R01-DK088220]. D.C.B. is supported by the National Heart,
Lung and Blood Institute [5T32HL007360-34]. Deposited in PMC for release
after 12 months.
Competing interests statement
J.M.G. is a co-founder and shareholder of Reata Pharmaceuticals.
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