Obesity has become a serious worldwide health problem driving an
ever-growing interest for the study of adipose tissue development.
Obesity is the result of an imbalance between energy intake and
expenditure, and is often characterised by an increase in both
adipocyte size and numbers. In addition to storing and mobilizing
neutral lipids in response to various hormones, adipocytes are
endowed with intracrine, autocrine/paracrine and endocrine
properties, such as the secretion of leptin and adiponectin (Ailhaud
and Hauner, 2003). The adipocyte lineage originates from
mesenchymal progenitors, which by yet unknown mechanisms form
adipocyte precursor cells or preadipocytes, that then differentiate
into mature, lipid-containing adipocytes. This overall process is
termed adipogenesis and can occur during the whole lifespan of an
organism (Ailhaud et al., 1992; Hausman et al., 2001; Rosen and
The differentiation of preadipocytes into adipocytes has been
extensively studied in vitro (reviewed by Otto and Lane, 2005;
Rosen and MacDougald, 2006). This is mostly because of the
establishment of immortal, preadipocyte cell lines that were selected
from disaggregated mouse embryos or from adult adipose tissue for
their ability to accumulate cytoplasmic triacylglycerols (Green and
Kehinde, 1975; Green and Kehinde, 1976; Green and Meuth, 1974;
Negrel et al., 1978). These cell lines are believed to be faithful
models of preadipocyte differentiation and they have provided
important insights into the control of the late steps of adipogenesis.
Preadipocyte differentiation is governed by the sequential expression
of a set of key transcription factors, including members of the
CCAAT/enhancer binding protein (C/EBP) and the peroxisome
proliferator-activated receptor (PPAR) families (Mandrup and Lane,
1997; Tontonoz et al., 1995), as well as the adipocyte determination
and differentiation factor-1/sterol response element binding protein
1c (ADD1/SREBP1c) (Ailhaud et al., 1992; Rosen et al., 2000).
Terminal differentiation is accompanied by changes in the
expression of cytoskeletal and extracellular matrix proteins
(Spiegelman and Farmer, 1982) and by dramatic increases in the fat
cell-specific expression of PPAR?, adipocyte fatty acid binding
protein (FABP4) and several lipid-synthesizing enzymes such as
glycerophosphate deshydrogenase (GPDH) (MacDougald and Lane,
1995). However, established preadipocyte cell lines are limited for
studying early events of differentiation as they represent cells that
are already committed to the adipogenic lineage.
The initial commitment of mesenchymal progenitors to the
adipocyte lineage is much less understood, mostly because there are
no specific cell surface markers available so far to identify and isolate
mesenchymal progenitors or preadipocytes in vivo. Furthermore,
because adipose tissue cannot be detected macroscopically during
mammalian embryogenesis, minimal information is available
regarding the ontogeny of fat cells. In the trunk and limbs, where most
of the adipose tissues will finally form, mesenchyme is of mesodermal
origin and therefore adipocytes are thought to derive from mesoderm
only. However, it is worth noting that during development of higher
vertebrates, the mesoderm is not the only germ layer source of
mesenchymal cells. In the head, the facial bones, jaw and associated
connective tissues have been shown to derive from the neural crest
(NC). The NC is a vertebrate cell population that arises from the
neuroectoderm. After neural tube closure, NC cells undergo an
epithelio-mesenchyme transition and migrate to diverse regions in the
developing embryo. They then become widely distributed in
numerous sites, where they differentiate into diverse cell types. NC-
derivatives include pigment cells, neurons and glial cells of the
peripheral nervous system (PNS), as well as some endocrine cells. In
the head and neck, the NC also yields mesectodermal cells, which are
ectoderm-derived mesenchymal cells differentiating into connective
The generation of adipocytes by the neural crest
Nathalie Billon1,*, Palma Iannarelli2, Miguel Caetano Monteiro1, Corinne Glavieux-Pardanaud3,†,
William D. Richardson2, Nicoletta Kessaris2, Christian Dani1and Elisabeth Dupin3,‡
Fat cells (adipocytes) develop from adipocyte precursor cells (preadipocytes) that themselves derive from mesenchymal progenitors.
Although the events controlling preadipocyte differentiation into mature adipocytes have been largely explored, the mechanisms
that direct mesenchymal progenitors down the adipocyte pathway remain unknown. Similarly, although adipocytes are generally
thought to derive from mesoderm, key information is lacking regarding the origin and the development of the adipose tissue
during embryogenesis. The aim of this study was to gain insight into the ontogeny of fat cells, both in mouse embryonic stem (mES)
cell-derived cultures and during normal development. We first used genetically engineered mES cells to produce and select ES cell-
derived neuroepithelial progenitors and showed that neuroectoderm, rather than mesoderm, may be a source of adipocytes in mES
cell-derived cultures. We then used primary and secondary cultures of developing quail neural crest (NC) cells to demonstrate that
NC cells are able, upon stimulation with defined factors, to differentiate into adipocytes, thus providing a powerful system to study
the earliest stages of adipocyte differentiation. Finally, we mapped NC derivatives in vivo using Cre-mediated recombination in
transgenic mice and demonstrated that a subset of adipocytes originates from the NC during normal development.
KEY WORDS: Adipocyte, Differentiation, Development, Origin, Neural crest, Mouse, Quail
Development 134, 2283-2292 (2007) doi:10.1242/dev.002642
1Institut de Recherche, Signalisation, Biologie du Développement et Cancer, CNRS
UMR 6543, Centre de Biochimie, Faculté des Sciences, Université Nice Sophia-
Antipolis, Nice, France. 2Wolfson Institute for Biomedical Research and Department
of Biology, University College London, Gower Street, London WC1E 6BT, UK.
3Laboratoire d’Embryologie Cellulaire et Moléculaire, CNRS UMR 7128, Nogent sur
*Author for correspondence (e-mail: firstname.lastname@example.org)
†Present address: Hôpital Pitié-Salpétrière, LGN/CNRS UMR7091, Paris, France
‡Present address: Institut de Neurobiologie Alfred Fessard, CNRS UPR2197,
Laboratoire DEPSN, Gif-sur-Yvette, France
Accepted 3 April 2007
tissue cells, vascular smooth muscle cells, tendons, dermis,
odontoblasts, cartilages and bones (reviewed by Dupin et al., 2006; Le
Douarin and Kalcheim, 1999; Le Douarin et al., 2004). In contrast to
other mesenchymal cells, lineage relationships between the NC and
adipocytes have not been carefully explored in the past. Seminal
grafting experiments performed in the 1970s and 1980s in the avian
embryo indicated, however, that the NC might generate adipocyte-like
cells in some areas of the face and the neck (Le Lievre and Le Douarin,
Mouse embryonic stem (mES) cells might provide an alternative
system for studying the early steps of adipocyte development. mES
cells are proliferating, pluripotent stem cells that have been isolated
from blastocyst-stage mouse embryos (Bradley et al., 1984; Brook
and Gardner, 1997; Evans and Kaufman, 1981; Martin, 1981). They
can be propagated indefinitely in an undifferentiated state in vitro,
and they can be easily genetically modified (Smith et al., 1988;
Williams et al., 1988). When ES cells are cultured without
leukaemia inhibitory factor (LIF) on a non-adherent surface, they
aggregate to form embryoid bodies (EBs) in which the cells form
ectodermal, mesodermal and endodermal derivatives, thus offering
a unique cell culture model to study early steps of development
(Keller, 1995). Extra-embryonic endoderm, cardiac, endothelial and
haematopoietic cell types normally predominate during
differentiation of mES cells in EBs (Doetschman et al., 1985;
Fehling et al., 2003). However, exposure of developing EBs to all-
trans retinoic acid (RA) induces alternative lineages (Rohwedel et
al., 1999). At high doses, RA promotes neural differentiation (Bain
et al., 1995; Okabe et al., 1996). By contrast, early and transient
treatment with intermediate levels of RA seems to favour the
emergence of mesenchymal progenitors capable, when subsequently
exposed to appropriate signal molecules, of generating adipocytes,
osteoblasts or chondrocytes (Dani et al., 1997; Kawaguchi et al.,
2005; Phillips et al., 2003). It is unknown whether these different
mesenchymal cell types develop independently or from a common
multipotent precursor. In addition, the cellular and molecular
mechanisms underlying formation of mesenchymal precursors from
ES cells remain obscure. Recent data by Kawaguchi et al. suggest
that either a mesodermal subset or NC-like cells, or both, may be the
source(s) of mesenchymal precursors in mES cell-derived cultures
(Kawaguchi et al., 2005).
The aim of this study was to gain insight into the ontogeny of fat
cells, both in ES cell-derived cultures and during normal
development. We first used genetically engineered mES cells to
produce and select mES cell-derived neuroepithelial progenitors and
we show that the neuroectoderm/NC, rather than the mesoderm, may
be a source of adipocytes in mES cell-derived cultures. Most
importantly, we then isolated primary NC cells from developing
quail embryos and we demonstrated that these cells can be induced
to differentiate into mature adipocytes upon stimulation with defined
growth factors and hormones. Finally, we used Cre-lox fate mapping
in Sox10-Cre transgenic mice to demonstrate that a subset of
adipocytes originate from the NC during normal development.
MATERIALS AND METHODS
ES cell culture and adipocyte differentiation
All reagents were from Sigma unless otherwise indicated. All cells were
incubated at 37°C in a humidified 5% CO295% air atmosphere. CGR8
(Mountford et al., 1994) and OSG (Billon et al., 2002) mES cells were
maintained without feeders in Glasgow modification of Eagle’s medium
(GMEM, Invitrogen) supplemented with foetal calf serum (FCS; 10%,
Dutscher), non-essential amino acids (Invitrogen), glutamine (10 mM,
Invitrogen), sodium pyruvate (100 mM, Invitrogen), 2-mercaptoethanol (1
mM) and LIF, as described (Wdziekonski et al., 2006).
EB formation and adipocyte differentiation were performed according to
the protocol described by Dani et al. (Dani et al., 1997) with minor
modifications. ES cell culture medium without LIF was used throughout and
changed every day during suspension culture and every 2 days after plating.
The hanging drop method (Hole, 1999) was employed to aggregate 103cells
per 20 ?l of medium for 3 days. Cell aggregates were then pooled and
cultured in suspension with RA (10–7M) for 3 days. For subsequent
differentiation, EBs were plated on gelatin-coated dishes. After 24 hours,
adipocyte differentiation was induced by the addition of insulin (170 nM),
triiodothyronine (T3, 2 nM), and roziglitazone (0.5 ?M), a treatment
referred to herein as DIF1.
Neural differentiation protocol on genetically engineered ES cells
Genetically engineered, selectable sox2-?geo/oct4-tk ES cells (OSG) have
been described elsewhere (Billon et al., 2002). Basically, these cells have a
?geogene inserted into the Sox2locus and a hygromycin-thymidine-kinase
(tk) fusion gene inserted into the Oct4 locus. As Sox2 is specifically
expressed in neuroepithelial cells and Oct4 is expressed in undifferentiated
ES cells, treatment of these doubly targeted ES cells with both Ganciclovir
and G-418 allows selection of neuroepithelial cells, while eliminating
residual undifferentiated ES cells. Neural differentiation and selection of
neuroepithelial precursors were performed as described (Billon et al., 2002):
on day 0, differentiation was induced by growing the cells in suspension
without LIF in order to induce formation of EBs, and RA (10–6M) was
added on day 4 to promote neural development. After 2 days, the medium
was changed to a 50:50 mixture of Dulbecco’s modified Eagle’s medium
(DMEM)-F12 containing N2 supplement and Neurobasal medium
containing B27 supplement (Invitrogen). FGF-2 (PreproTech, Rocky Hill,
NJ, USA) was added at 20 ng/ml together with G-418 (100 ?g/ml) and
Ganciclovir (2.5 ?M) to select for neuroepithelial cells and against
undifferentiated ES cells, respectively. At day 8, EBs were dissociated with
trypsin and replated in the same medium on poly-D-lysine (PDL; 10 ?g/ml)-
and laminin (10 ?g/ml)-coated tissue culture flasks. After 2 more days (day
10; 4 days of selection), adipocyte development was promoted by the
addition of DIF1 treatment (see above) in ES cell medium without LIF.
Quail NC cell cultures
Fertile quail eggs, obtained from commercial sources, were incubated at
38°C and staged according to Hamburger and Hamilton (HH) (Hamburger
and Hamilton, 1951) or according to the number of pairs of somites formed.
Cephalic neural crest cells (CNCCs) were obtained from explants of
mesencephalic-rhombencephalic neural tubes that were microsurgically
removed from quail embryos at the 6-8 somite stage (stage 9 HH). Trunk
neural crest cells (TNCCs) were isolated similarly from neural tubes
dissected at the 18-25 somite stage from the level of the last 10 somites
formed. Explanted neural tubes (including premigratory NC) were cultured
in cloning medium, as described (Trentin et al., 2004). After 48 hours, the
neural tubes were removed, leaving behind the outgrowth of migratory NC
cells, which constitute the primary cultures.
After five days of primary culture, CNCC or TNCC were either left in
cloning medium (control) or switched to various combinations of media
known to be permissive for adipocyte development (Rodriguez et al., 2004;
Student et al., 1980): L1 medium contains DMEM (Invitrogen) supplemented
with 10% FCS (Dutscher). Hmads medium is a 50:50 mixture of DMEM and
Ham’s-F12 (Invitrogen) supplemented with 10 ?g/ml transferrin. Adipocyte
differentiation was induced using DIF1 (see above) or DIF2 treatments. In
DIF2 treatment, cells were first treated with dexamethasone (1 ?M), 1-methyl-
3-isobutylmethyl-xanthine (IBMX, 0.5 mM), insulin (170 nM), T3 (2 nM) and
roziglitazone (0.5 ?M) for 2 days and then switched to DIF1 treatment (i.e.
dexamethasone and IBMX were omitted).
To perform secondary cultures, CNCC or TNCC from 2-day primary
cultures (see above) were harvested by treatment with trypsin-EDTA
solution (Trentin et al., 2004) and seeded in four-well culture plates (Nunc)
at a density of 5?103cells/well (20 ?l) in cloning medium. After 4 more
days, adipocyte differentiation was induced using various combinations of
the media described above.
CNCC and TNCC cultures were recorded as positive for adipocyte
differentiation only when they included at least 10 adipocytes per culture.
Development 134 (12)
All animals were housed in the University College London (UCL) animal
facility. Animal experiments were performed according to the UK Animal
Act 1986 and approved by the UCL Care Committee. Rosa26R-YFP mice
were purchased from the Jackson Laboratory (Bar Harbor, Maine, USA).
These mice were crossed with Sox10-Cremice expressing Cre recombinase
under the control of the Sox10 promoter (Matsuoka et al., 2005). Offspring
with the genotype Sox10-Cre/Rosa26R-YFP were used in this study. Eight-
week-old mice were asphyxiated with CO2and tissues were immediately
removed and fixed in 4% (w/v) paraformaldehyde (PFA) in phosphate-
buffered saline (PBS) at room temperature for 1 hour. All tissues were
cryoprotected overnight in 20% (w/v) sucrose in PBS, embedded in Tissue-
Tek optimum cutting temperature (OCT) compound (R. A. Lamb,
Eastbourne, UK) and frozen on dry ice. Sections of 25 ?m thickness were
cut on a cryostat and mounted on SuperFrost®Plus glass slides (VWR
Total RNA was extracted using the TRI-Reagent kit (Euromedex, Souffel
Weyersheim, France) according to the manufacturer’s instructions. cDNAs
were synthesised using SuperScript Reverse Transcriptase (Invitrogen)
according to the supplier’s instructions and were used as templates for the
polymerase chain reaction (PCR). All primer sequences are detailed in Table
1. For semi-quantitative PCR, parameters were 94°C for 30 seconds for the
denaturing step, 60°C for 30 seconds for the annealing step and 72°C for 1
minute for the elongation step. Real-time PCR assays were run on an ABI
Prism 7000 real-time PCR machine (PerkinElmer Life Sciences). Reactions
were performed according to the manufacturer’s instructions using SYBR
green master mix (Eurogentec, Angers, France). PCR conditions were as
follows: 2 minutes at 50°C, 10 minutes at 95°C, followed by 40 cycles of 15
seconds at 95°C and 1 minute at 60°C. Gene expression was quantified using
the comparative-?Ct method. Data were normalised relative to Gapdh
amplification and the highest expression was defined as 100%.
Immunocytochemistry and stainings
mES cells were cultured on PDL-coated glass coverslips and fixed in 4% PFA
in PBS for 5 minutes at room temperature. After washing with PBS, cells were
incubated for 30 minutes in 10% normal goat serum/0.1% Triton X-100 to
block non-specific staining. They were then incubated for 1 hour in the first
antibody, washed in PBS, and incubated for 1 hour in FITC-coupled goat anti-
mouse immunoglobulin (Ig) or goat anti-rabbit Ig antibodies (diluted 1:100;
Jackson ImmunoResearch Laboratories) and bisbenzamide (5 ng/ml; Hoechst
No. 33342; Sigma). Coverslips were mounted in Citifluor mounting medium
(CitiFluor, London, UK) and examined with a Zeiss fluorescence microscope.
The following antibodies were used: monoclonal anti-rat nestin antibody
(diluted 1:100, Pharmingen), polyclonal anti-Sox9, anti-Sox10 and anti-
FoxD3 antibodies (diluted 1:100, Chemicon).
Quail NC cell cultures were fixed at day 20 in 4% PFA in PBS for 30
minutes and cell phenotypes were analysed by immunocytochemistry as
described previously (Trentin et al., 2004). Briefly, glial cells and
myofibroblasts were identified using anti-Schwann cell myelin protein
(SMP) (Dulac et al., 1988) and an antibody to ?-smooth muscle actin
(Sigma), respectively; neurons and adrenergic cells were stained with anti-
?III-tubulin (Chemicon) and anti-quail tyrosine hydroxylase (Fauquet and
Ziller, 1989), respectively; melanocytes were recognised by the presence of
pigment granules whereas unpigmented melanoblasts were labelled with
anti-melanocyte/melanoblast early marker (Nataf et al., 1993). Secondary
FITC- and Texas Red-conjugated antibodies were purchased from Southern
Biotech (Birmingham, AL, USA). Fluorescence was observed under an X70
Olympus inverted microscope.
For immunofluorescence staining of yellow fluorescent protein (YFP)
and perilipin, cryostat sections of transgenic mice were air-dried and
immersed in PBS, pH 7.4, containing 0.1% Triton X-100 for 5 minutes to
remove excess OCT compound. Non-specific binding sites were blocked by
incubating slides with 10% heat-inactivated sheep serum/0.5% Triton X-
100/PBS for at least 1 hour at room temperature. After blocking, sections
were incubated overnight at 4°C with rabbit anti-GFP polyclonal antibody
(ab290, Abcam; diluted 1:8000) and guinea pig anti-perilipin polyclonal
antibody (PROGP20, RDI, Concord, MA, USA; diluted 1:2000). After
washing the sections several times in PBS containing 0.1% Triton X-100,
they were incubated with a secondary antibody mixture of fluorescein-
conjugated anti-rabbit IgG and rhodamine-conjugated anti-guinea pig IgG
(both at 1:200; Pierce Biotechnology) for 1 hour at room temperature. After
several washes in PBS containing 0.1% Triton X-100, cell nuclei were
counterstained with Hoechst (0.01 mg/ml; Sigma). Sections were
coverslipped with an antifade mounting medium (Dako Cytomation
Fluorescent) and observed under a confocal microscope (Leica TCS SP).
Neutral lipid accumulation was assessed in mES cells and NC cell
cultures by Oil Red O staining as previously described (Abderrahim-
Ferkoune et al., 2003).
Early steps of adipogenesis in mES cells are
correlated with downregulation of mesoderm
markers and upregulation of neural/NC markers
To explore the origin of adipocytes derived from mES cells, we first
induced adipogenesis using a standard protocol involving treatment
of early differentiating EBs with RA (Dani et al., 1997). We treated
EBs with RA between day 3 and day 6 of differentiation, a period
known to be permissive for adipocyte development, and we examined
expression of mesodermal (Brachyury), mesenchymal (Sox9),
neuroepithelial (Sox1) and NC (FoxD3, Sox9 and Sox10) marker
genes. Consistent with previous reports, we observed that, whereas
treatment of nascent EBs with RA resulted in marked reduction of
Brachyury expression (Fehling et al., 2003; Kawaguchi et al., 2005),
Sox1, Sox9, Sox10and FoxD3mRNAs were all detected in RA-treated
EBs (data not shown) (Kawaguchi et al., 2005). Together, these data
support the idea that RA treatment, largely used to induce adipocyte
development in mES cells, reduces mesoderm formation and favours
neuroepithelial/NC development in EBs.
Adipocytes can be obtained from a purified
population of mES cell-derived neuroepithelial
We then tested the hypothesis that neuroectoderm, rather than
mesoderm, could be a source of adipocytes in mES cell-derived
cultures. We used genetically engineered, selectable Sox2-?geo/oct4-
tk mES cells that allow enrichment for neuroepithelial progenitors
(Billon et al., 2002; Li et al., 1998). We first used a standard, RA
treatment protocol to promote neural development of these selectable
mES cells (Bain et al., 1995; Billon et al., 2002; Li et al., 1998). After
6 days, we selected EBs with G418 and Ganciclovir to enrich for
neuroepithelial cells and eliminate residual undifferentiated mES cells,
respectively. After 4 days of selection (Fig. 1A, Day 10+selection),
Oct4 mRNA could barely be detected, suggesting that no residual
undifferentiated ES cells were present in the culture. By contrast, Sox1
andSox2mRNAs were readily detected (Fig. 1A), and more than 85%
of the cells expressed the neuroepithelial marker nestin (Fig. 1B),
indicating that a combined negative and positive selection strategy was
extremely efficient to generate highly enriched populations of
neuroepithelial cells (Billon et al., 2002). This treatment also resulted
in a significant increase in Sox9, Sox10and FoxD3mRNAs (Fig. 1A),
suggesting that NC-like cells might develop within the selected
population of neuroepithelial cells. These data were further supported
by the finding that a high percentage of the cells also expressed
FoxD3, Sox9 and Sox10 proteins (Fig. 1B).
To investigate whether selected, mES cell-derived neuroepithelial
cells could develop towards adipogenesis, we plated these cells and
cultured them in the presence of factors known to promote adipocyte
differentiation in ES cell-derived cultures (DIF1) (Dani et al., 1997).
We used accumulation of lipid droplets within the cells, as well as
Developmental origin of adipocytes
fat cell-specific expression of FABP4 mRNA, to monitor adipocyte
differentiation. As shown in Fig. 1C, mature adipocytes containing
lipid droplets could easily be observed after 14 days of treatment
with adipogenic factors. Furthermore, FABP4 mRNA was readily
detected in cells treated with adipogenic factors (Fig. 1D). By
contrast, in the absence of adipogenic factors, mES cell-derived
neuroepithelial cell cultures showed neither lipid accumulation nor
FABP4 expression (data not shown).
Together, these data suggest that, in vitro, adipocytes can develop
from a highly enriched population of mES cell-derived
neuroepithelial cells, possibly through a NC pathway.
Adipocytes can differentiate from quail cephalic
and truncal NC cells in vitro
We then checked whether adipocytes could develop from primary
NC cells isolated from a normal developing embryo. We used in
vitro cultures of quail NC cells because they have been instrumental
in establishing the developmental potentialities of cephalic and trunk
NC cells (Baroffio et al., 1988; Baroffio et al., 1991; Dupin et al.,
1990; Lahav et al., 1998; Trentin et al., 2004).
We first isolated neural tubes (including the premigratory NC)
from the cephalic region of quail embryos at HH stage 9(Hamburger
and Hamilton, 1951), and allowed cephalic NC cells (CNCC) to
migrate away from the neural tubes for 48 hours in explant cultures.
We then removed neural tubes and grew migrating CNCC in culture
medium permissive for differentiation along the main NC derivatives
(i.e. cloning medium) (Trentin et al., 2004). Four days later (at day
6 of culture), we either maintained CNCC under these conditions or
switched them to culture media known to be permissive for
adipocyte differentiation of either mouse preadipocyte cell lines (i.e.
L1 medium) (Student et al., 1980) or human adipose tissue-derived
stem cells (i.e. serum-free hmads medium) (Rodriguez et al., 2004).
We induced adipocyte differentiation in each of these three media
using two well-described adipogenic protocols (i.e. DIF1 and DIF2,
see Materials and methods section). After 15 more days, we assessed
for the presence of adipocytes using their characteristic
morphological feature (lipid droplet-filled cytoplasm) and by
staining the cultures with Oil Red O to reveal neutral lipid droplets.
As shown in Fig. 2A, typical mature adipocytes could readily be
detected in CNCC cultures stimulated to differentiate in cloning
Development 134 (12)
Fig. 1. Development of adipocytes from ES cell-derived neuroepithelial precursors. Sox2-?geo/oct4-tk genetically engineered ES cells
were treated with RA and then selected between day 6 (D6) and day 10 (D10) to enrich for neuroepithelial cells and to eliminate residual
undifferentiated ES cells. They were then induced to differentiate towards the adipocyte lineage. (A) At various times, the cells were processed for
quantitative PCR analysis using Oct4, Sox1, Sox2, Sox9, Sox10, FoxD3 or GAPDH probes. Data were normalised relative to GAPDH amplification
and the highest expression was defined as 100%. Similar results were obtained in two independent experiments. +sel, with selection; –sel, no
selection. (B) After selection (D10), neuroepithelial precursors were stained with anti-FoxD3, anti-Sox9 or anti-Sox10 antibody (red) to identify
NC-like cells, and with bisbenzimide to identify cell nuclei (blue). 14 days after induction of adipocyte differentiation (+), adipocytes were
identified either using their characteristic morphology (C) or by RT-PCR to detect FABP4 mRNA (D). The results in D are shown for two
independent experiments. Scale bar: 100 ?m.
medium, as well as in L1 medium (data not shown). By contrast,
hmads medium could not sustain CNCC survival, resulting in the
death of most of the cultures. Quantification of the number of CNCC
primary cultures containing adipocytes in each medium condition
revealed that up to 40% of adipocyte-containing cultures could be
obtained in cloning medium, and 37% in L1 medium (Fig. 2B). No
adipocyte developed in the presence of hmads medium. The addition
of 1% serum to this medium (hmads+S medium), however,
enhanced CNCC survival and formation of up to 50% of adipocyte-
containing cultures in differentiating conditions (Fig. 2B). Together,
these data suggest that adipocytes can readily develop from primary
cultures of CNCC under various culture conditions.
To reduce the variability of cell differentiation between the
different primary CNCC cultures, we next investigated the
adipogenic potentialities of CNCC in secondary cultures, prepared
by replating primary CNCC at the initial density of 2.5?105cells
per ml. As shown in Fig. 2C, 89-100% of day 20 secondary cultures
contained adipocytes when induced to differentiate in either cloning
or L1 medium. Furthermore, several markers of adipocyte
differentiation, including CEBP?, PPAR? and FABP4 mRNAs,
could readily be detected in these conditions after 9 days of culture,
when the first differentiating adipocytes were detected under
microscopy (Fig. 2D). Therefore, our results clearly indicate that, at
least in vitro, CNCC can differentiate into adipocytes with a high
efficiency. Analysis of CNCC cultures at day 20 with lineage-
specific markers indicated that other CNCC derivatives, such as glial
cells, neurons, melanocytes and myofibroblasts, also differentiated
in adipocyte-containing cultures (data not shown).
To investigate whether the trunk NC cells (TNCC) can
differentiate into adipocytes in vitro, we isolated quail NC cells from
the thoracic level, replated them into secondary cultures and treated
them as above. As shown in Fig. 3A, adipocytes could readily be
observed after 20 days in adipogenic conditions. These cultures also
comprised SMP-positive Schwann cells (Fig. 3A), as well as other
cell types known to arise from TNCC (data not shown).
Quantification of TNCC cultures containing adipocytes revealed that
more than 40% of TNCC cultures contained adipocytes when
submitted to DIF2 treatment in cloning or L1 media (Fig. 3B).
Therefore, trunk NC cells in culture exhibit adipogenic
developmental potential, although with a lower frequency than
cephalic NC cells in similar conditions.
Some adipocytes are derived from the NC during
To investigate whether subsets of adipocytes originate from the NC
during normal development, we adopted a recombinase-mediated
lineage labelling strategy in transgenic mice. We used Sox10-Cre
transgenic mice to map NC derivatives because to date, Sox10 is
Developmental origin of adipocytes
Fig. 2. Development of adipocytes in primary cultures of quail cephalic NC cells (CNCC). (A,B) Primary cultures of CNCC were obtained
from mes-rhombencephalon of HH stage 9 quail embryo in explant culture and expanded for 5 days in cloning medium. Adipocyte differentiation
was then induced using different media (cloning, L1 and hmads) and adipogenic treatments (DIF1 or DIF2). Adipocytes were identified after
15 days. (A) Typical adipocytes show lipid droplet-filled cytoplasm (left) and are stained with Oil Red O, which reveals neutral lipids (right).
Scale bar: 100 ?m. (B) Quantification of CNCC primary cultures containing adipocytes after treatment with the mentioned media and adipogenic
treatments. A total of 100 cultures were analysed. (C,D) Secondary cultures of quail CNCC were isolated from 48-hour primary cultures. Adipocyte
differentiation was then induced at day 6 using the mentioned media and adipogenic treatments. (C) Quantification of secondary CNCC cultures
containing adipocytes after 15 days of treatment. A total of 84 cultures were analysed. (D) Expression of CEBP?, PPAR?, FABP4 and 18s RNAs after
3 days in cloning medium+DIF1. The results are shown for two independent CNCC cultures out of 10 distinct experiments.
considered as the best bona fide NC marker (Matsuoka et al., 2005).
Indeed, Sox10 is expressed strongly in premigratory and migratory
NC cells at all rostro-caudal levels of the neural axis during early
mouse embryonic development (Kuhlbrodt et al., 1998). Most
importantly, it is not expressed in cephalic and somitic mesoderm
(Ferguson and Graham, 2004). We crossed Sox10-Cre transgenic
founders to a Cre-conditional R26-YFPreporter line (Srinivas et al.,
2001) to identify the YFP+ NC-derived population at a single-cell
resolution. Extensive analysis of Cre and Sox10 expression on the
double transgenic offspring has been previously conducted by
Matsuoka et al. to ascertain that Sox10 activation of the Cre
transgene accurately reflects endogenous gene expression
(Matsuoka et al., 2005).
We examined Sox10-Cre/R26-YFP offspring for the presence of
YFP+ adipocytes at postnatal day 28 (P28), because at this stage the
adipose tissue can be detected at both cephalic (jaws) and trunk
(subcutaneous and perigonadal depots) levels. We confirmed
adipocyte identity by immunolabelling sections with perilipin, an
adipocyte marker that is specifically expressed at the periphery of
lipid droplets (Greenberg et al., 1991). As shown in Fig. 4A-D,
YFP+, perilipin+ adipocytes could be detected in the area around the
salivary gland and ears in Sox10-Cre/R26-YFP mice. Perilipin
showed complete coexpression with YFP, suggesting that the
majority of adipocytes in this site are derived from NC, and not from
mesoderm. By contrast, YFP expression could not be detected in
either peri-ovarial (Fig. 4E-H) or subcutaneous (Fig. 4I-L)
adipocytes, suggesting that these two truncal fat depots do not arise
from NC. Consistent with these findings, staining for ?-
galactosidase activity in 18-month-old Sox10-Cre/R26-lacZ males
revealed ?-galactosidase-positive adipocytes around the salivary
gland, but not in truncal anatomic regions including subcutaneous,
perirenal, periepididymal and interscapular adipose depots (data not
Throughout the past century, observations about the development of
adipose tissue have been recorded in a variety of species. Early on,
it was noticed that fat depots develop in many regions during
mammalian postnatal life, generally in areas composed of loose
connective tissues such as the subcutaneous layers between muscles
and dermis, but also around internal organs (Rosen et al., 2000).
Extensive histological and histochemical studies in pig foetuses have
revealed that immature fat cells, containing small lipid droplets,
appear in subcutaneous areas from the last third of the gestation
period, always in close association with blood vessels (Hausman and
Kauffman, 1986; Hausman et al., 1990). Several mesodermal cell
types within the foetus have been proposed to be adipocyte
precursors, including fibroblasts and endothelial cells (reviewed by
Hausman et al., 1980). However, the diffuse nature of adipose
tissues, as well as the lack of molecular markers to prospectively
identify adipocyte precursors, have left the origin of the adipocyte
lineage uncertain. In the present study, we have used several
approaches to address this issue in mice and avians and we provide
direct evidence for the contribution of the NC to the adipocyte
To gain insight into the ontogeny of fat cells, we first used
mouse ES cells, because they provide a powerful system to model
the earliest stages of mammalian development (Keller, 2005).
Early and transient treatment with RA turned out to be crucial for
quantitative induction of adipocytes and other mesenchymal cell
types from ES cells (Dani et al., 1997; Kawaguchi et al., 2005).
However, the basis for this effect of RA is uncertain. It has been
proposed that RA might posteriorise nascent mesoderm in
developing EBs, without promoting somitogenesis nor sclerotome
development (Kawaguchi et al., 2005). RA treatment also favours
the formation of neuroectoderm derivatives within EBs, including
NC-like cells expressing Sox9, Sox10 and FoxD3 (this study) (see
also Kawaguchi et al., 2005). Because some mesenchymal cells
are known to develop from the NC during normal development
(reviewed by Dupin et al., 2006; Le Douarin et al., 2004; Le
Douarin and Kalcheim, 1999), it has been proposed that either a
mesodermal subset or NC, or both, may be the source(s) of
mesengenesis in ES cells (Kawaguchi et al., 2005). In support of
the second hypothesis, we show here, using genetically
engineered ES cells, that it is possible to derive adipocytes from
a highly enriched population of ES cell-derived neuroepithelial
precursors. These precursors express NC markers, suggesting that
adipocytes developing from RA-treated ES cells in vitro might
follow a NC, rather than a mesodermal, differentiation pathway.
However, the elucidation of the molecular events involved in RA-
induced adipogenesis in mES cells is unclear at present and
requires further investigation. Interestingly, RA-independent
formation of major neuroectodermal derivatives, including NC
cells, has been reported from mouse and primate ES cells in vitro
(Mizuseki et al., 2003). NC induction involved exposure to
stromal cell-derived inducing activity (SDIA) and BMP4, and
allowed the generation of PNS neurons and smooth muscle cells.
Development 134 (12)
Fig. 3. Development of adipocytes in secondary cultures of quail
trunk NCC (TNCC). TNCC that had migrated from cultured thoracic
neural tubes were replated into secondary cultures and treated as in
Fig. 2. (A) Typical adipocytes showing Oil Red O-stained lipid droplets
(left) and SMP-positive glial cells (right) were identified after 20 days in
adipogenic conditions. Scale bar: 100 ?m. (B) Quantification of TNCC
cultures containing adipocytes after treatment with the mentioned
media and adipogenic treatments. A total of 42 cultures were analysed.
The formation of other mesenchymal derivatives, however, was
not assessed in this study. Furthermore, in this system too,
the molecular pathways underlying NCC induction and
differentiation remain to be clarified (Mizuseki et al., 2003).
Regardless of the uncertainty about NCC and adipocyte formation
in mES cell cultures, independent observations indicate that some
adipocytes might normally develop through a NC pathway in vivo.
Indeed, quail-chick grafting experiments performed by Le Lievre
and Le Douarin indicated that the cephalic NC generates adipose
cells in the skin over the calvarium and in the ventral part of the neck
(Le Lievre and Le Douarin, 1975). In the present report, we revisited
these observations through modern cell fate mapping experiments
in the mouse. Using Sox10-Cre/R26 transgenic mice (Matsuoka et
al., 2005), we have demonstrated that a subset of adipocytes in the
face (salivary gland and ear) indeed originate from NC, and not from
mesoderm, during normal development. The contribution of NC to
mesenchymal cell types during development is not restricted to the
adipocyte lineage and has been established in various classes of
vertebrates. The replacement of the cephalic NC by its quail
counterpart in chick embryos showed that the facial and visceral
skeleton, including the hyoid cartilages, as well as the frontal,
parietal and squamosal bones, are NC derived; only the occipital and
otic (partly) regions of the skull are of mesodermal origin. Moreover,
much of the dermis, all of the connective components of facial
musculature and the wall (except endothelium) of the blood vessels
that irrigate the face and forebrain have a NC origin (reviewed in Le
Douarin et al., 2004). The contribution of cephalic NC cells to
building the head skeleton and the cardiac outflow tract has been
confirmed in the mouse, using genetic Wnt1-Crefate mapping (Chai
et al., 2000; Jiang et al., 2000; Santagati and Rijli, 2003). Our
findings that mature adipocytes in the ear region arise from the NC
during mouse development provide further evidence of the crucial
role of the cephalic NC in generating the various mesenchymal
derivatives forming head structures and tissues, and identify the
adipose tissue as being one of them.
Developmental origin of adipocytes
Fig. 4. Development of adipocytes from
the NC during mouse development.
Permanent genetic lineage labelling of pre-
and post-migratory NC was achieved by
crossing transgenic mice carrying a Sox10-Cre
construct into a R26-YFP reporter. Double
immunolabelling of P28 Sox10-Cre/R26-YFP
offspring with anti-GFP (green) and anti-
perilipin (red) antibodies was used to identify
NC derivatives and adipocytes, respectively.
Bisbenzimide was used to identify cell nuclei
(blue). Sections show salivary gland and ear
(A-D), peri-ovarial (E-H) and trunk
subcutaneous (I-L) regions. There is almost
complete colocalisation of YFP and perilipin in
the salivary gland area, whereas no
overlapping can be detected in the ovary and
trunk subcutaneous adipose depots.
Scale bar: 50 ?m.
Table 1. PCR primer sequences
GeneSpecificity Forward primer (5?-3?) Reverse primer (5?-3?)Size (bp)Accession number
(B) Quantitative RT-PCR
NCC fate is not the same along the neural axis, as established by
mapping the anteroposterior origin of NC derivatives in quail-chick
chimera (Le Douarin et al., 2004). The generation of mesenchymal
derivatives was found to be restricted to the cephalic NC region
(from mid-diencephalon to somite 4 inclusive) in higher
vertebrates. Indeed, when the quail trunk NC is orthotopically
implanted in the chick, no quail mesenchymal cells are ever present
in the host (Le Douarin and Teillet, 1974; Nakamura and Ayer-le
Lievre, 1982). In order to confirm these data, Matsuoka et al.
analysed 23 truncal skeletal structures known to derive entirely
from mesoderm (including the pelvic bone, ilium, humerus, ulna,
tibia, fibula, digits and sacral vertebrae) in sox10-Cre/R26-GFP
mice (Matsuoka et al., 2005). They found that none of these
elements were GFP positive, whereas dorsal root ganglia and
Schwann cells surrounding motor axons, both of which have a trunk
NC origin, were GFP labelled. In accordance with these
observations, we report here that NC-derived adipocytes could be
found in the cephalic region, but not in truncal, subcutaneous and
perigonadal fat depots of Sox10-Cre/R26-YFPmice. These findings
suggest that truncal adipocytes do not arise from NC. They further
indicate that, similarly to other mesenchymal cells such as
chondrocytes and osteocytes, adipocytes have a different origin
along the anteroposterior axis: in the trunk they derive from the
mesoderm, whereas in the cephalic region adipocytes exhibit an
alternative origin in the NC. Whether these different origins reflect
functional differences is unknown at present. Of note,
morphological and functional differences have been reported for
different fat depots in rodents and humans: visceral adipose tissue
(VAT) and subcutaneous adipose tissue (SAT), for instance, differ
in various biochemical properties, such as insulin and adrenergic
response (Lafontan and Berlan, 2003; Montague and O’Rahilly,
2000). Whether cephalic versus truncal adipose depots also present
site-specific regulation remains to be assessed. It also remains to be
ascertained whether, in addition to the NC, the cephalic mesoderm
and/or the anteriormost somites also contribute to the formation of
some adipose tissues in the head and neck.
We report here that quail NCC, upon stimulation with defined
adipogenic factors, can efficiently differentiate into adipocytes in
vitro. It is worth noting that the ability to generate adipocytes in vitro
was not restricted to CNCC, but was also shared by TNCC, although
adipogenic potential was higher in CNCC. In vivo, trunk NC cells
give rise to melanocytes, PNS neurons and glial cells, and to adrenal
medullary cells. By contrast, the trunk NC does not form an axial
and appendicular trunk skeleton nor associated connective tissues
(Le Douarin et al., 2004). A recent study, however, suggested that
trunk NC cells provide a small contribution to mesenchymal tissues
by generating a subset of fibroblasts in the mouse sciatic nerve
(Joseph et al., 2004). Therefore, although during normal
development the property of NC to form mesenchyme is almost
restricted to the cephalic part of the neural axis, our finding that
TNCC in culture can generate adipocytes suggests that a hidden
capacity of trunk NC to yield mesenchymal cells can be revealed by
appropriate environmental cues. In support of this hypothesis, it has
been reported that small subsets of mesenchymal cells can be
derived from TNCC after unilateral heterotopic grafting, provided
that these cells develop in close relationship with host cephalic
migratory NC cells (Nakamura and Ayer-le Lievre, 1982). In
addition, long-term in vitro culture of avian TNCC can trigger their
differentiation into chondrocytes (Abzhanov et al., 2003; McGonnell
and Graham, 2002). Moreover, mouse TNC explants yield dentine
and bone when recombined with branchial arch 1 epithelium in
intraocular grafts (Lumsden, 1988). Finally, clonogenic cells from
avian and mammalian TNC generate myofibroblasts in addition to
neural and melanocytic cell types in vitro (Shah et al., 1996; Trentin
et al., 2004). Together, these data support the idea that the trunk NC
of higher vertebrates has cryptic mesenchymal differentiation
Our findings that quail NCC can generate adipocytes in vitro opens
exciting new opportunities to study the events regulating the earliest
stages of adipocyte lineage induction and specification from the NC.
Adipocytes produced from CNCC accumulate intracellular lipids as
multiple droplets. Furthermore, they express key adipocyte
differentiation regulators such as CEBP? and PPAR?, as well as the
specific adipocyte molecular marker FABP4. This pattern of gene
expression is entirely consistent with that observed upon
adipogenesis of murine clonal preadipocyte cell lines (Student et al.,
1980) and human adipose tissue-derived stem cells (Rodriguez et al.,
2004), suggesting that the regulatory pathways involved in adipocyte
terminal differentiation are conserved between these three species.
As a conclusion, we present here several lines of evidence
showing that adipocytes differentiate from the NC in vivo and in
vitro, thus providing important insight into the developmental origin
of the adipocyte lineage. First, the derivation of adipocytes from
mES cells was found to involve a neuroepithelial/NC-like, rather
than mesodermal, differentiation pathway. Second, we have shown
that quail cephalic and trunk NC cells can generate adipocytes in
vitro. Finally, we have determined that mature adipocytes in the ear
region arise from the NC during mouse development. Given that
clonogenic NC cells include multipotent progenitors yielding
chondrocytes, myofibroblasts and neural/melanocytic cells (Baroffio
et al., 1991; Trentin et al., 2004), it is tempting to speculate that the
NC might be a source of mesenchymal stem cells giving rise to
adipocytes, chondrocytes and possibly other mesenchymal cells, as
described in adult bone marrow (Caplan, 1991; Owen, 1988;
Pittenger et al., 1999). A major challenge now is to discover how and
when the adipocytic lineage segregates in NC-derived cells.
Analysis of the progeny of single NC cells should allow us to
determine whether adipocytes arise from multipotent progenitors or
from early committed cells.
We thank members of the Dani and Ailhaud laboratories for critical advice and
comments on the manuscript. This research was supported by CNRS and by
funding under the Sixth Research Framework Programme of the European
Union, Project FunGenES (LSHG-CT-2003-503494). Work at UCL was
supported by the UK Medical Research Council and the Wellcome Trust.
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