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First published online 16 May 2007
doi: 10.1242/dev.002642
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1 Institut de Recherche, Signalisation, Biologie du Développement et
Cancer, CNRS UMR 6543, Centre de Biochimie, Faculté des Sciences,
Université Nice Sophia-Antipolis, Nice, France.
2 Wolfson Institute for Biomedical Research and Department of Biology,
University College London, Gower Street, London WC1E 6BT, UK.
3 Laboratoire d'Embryologie Cellulaire et Moléculaire, CNRS UMR 7128,
Nogent sur Marne, France.
* Author for correspondence (e-mail: billon{at}unice.fr)
Accepted 3 April 2007
 |
SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Adipocyte, Differentiation, Development, Origin, Neural crest, Mouse, Quail
|
INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). 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 MacDougald,
2006).
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 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, 1975).
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.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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)
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 103 cells per 20 µl of medium for 3 days. Cell
aggregates were then pooled and cultured in suspension with RA
(10-7 M) 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 µM), a treatment referred to herein as DIF1.
Neural differentiation protocol on genetically engineered ES cells
Genetically engineered, selectable sox2-ßgeo/oct4-tk
ES cells (OSG) have been described elsewhere
(Billon et al., 2002).
Basically, these cells have a ßgeo gene inserted into the
Sox2 locus 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 (10-6 M) 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 µg/ml) and Ganciclovir (2.5 µM) 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 µg/ml)- and laminin (10 µg/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 µg/ml transferrin. Adipocyte differentiation was induced using DIF1 (see
above) or DIF2 treatments. In DIF2 treatment, cells were first treated with
dexamethasone (1 µM), 1-methyl-3-isobutylmethyl-xanthine (IBMX, 0.5 mM),
insulin (170 nM), T3 (2 nM) and roziglitazone (0.5 µM) 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
5x103 cells/well (20 µl) 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.
Animals
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-Cre mice 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 CO2 and 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 µm thickness were cut on a
cryostat and mounted on SuperFrost® Plus glass slides (VWR
International).
RT-PCR analysis
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%.
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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-ßIII-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).
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). 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, Sox10 and FoxD3 mRNAs 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 progenitors
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-ßgeo/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 and Sox2 mRNAs 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, Sox10 and
FoxD3 mRNAs (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 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).
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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 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.
|
, 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 mouse development
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 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-YFP reporter 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).
|
). 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 ß-galactosidase activity in
18-month-old Sox10-Cre/R26-lacZ males revealed
ß-galactosidase-positive adipocytes around the salivary gland, but not in
truncal anatomic regions including subcutaneous, perirenal, periepididymal and
interscapular adipose depots (data not shown). |
DISCUSSION |
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|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
).
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
lineage.
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. 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).
|
). 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-Cre fate 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.
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-YFP mice. 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 potentials.
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.
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
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
|
REFERENCES |
|---|
|
TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Abderrahim-Ferkoune, A., Bezy, O., Chiellini, C., Maffei, M.,
Grimaldi, P., Bonino, F., Moustaid-Moussa, N., Pasqualini, F., Mantovani, A.,
Ailhaud, G. et al. (2003). Characterization of the long
pentraxin PTX3 as a TNFalpha-induced secreted protein of adipose cells.
J. Lipid Res. 44,994
-1000.
Abzhanov, A., Tzahor, E., Lassar, A. B. and Tabin, C. J.
(2003). Dissimilar regulation of cell differentiation in
mesencephalic (cranial) and sacral (trunk) neural crest cells in vitro.
Development 130,4567
-4579.
Ailhaud, G. and Hauner, H. (2003). Handbook of Obesity. New York: Marcel Dekker.
Ailhaud, G., Grimaldi, P. and Negrel, R. (1992). Cellular and molecular aspects of adipose tissue development. Annu. Rev. Nutr. 12,207 -233.[CrossRef][Medline]
Bain, G., Kitchens, D., Yao, M., Huettner, J. E. and Gottlieb, D. I. (1995). Embryonic stem cells express neuronal properties in vitro. Dev. Biol. 168,342 -357.[CrossRef][Medline]
Baroffio, A., Dupin, E. and Le Douarin, N. M.
(1988). Clone-forming ability and differentiation potential of
migratory neural crest cells. Proc. Natl. Acad. Sci.
USA 85,5325
-5329.
Baroffio, A., Dupin, E. and Le Douarin, N. M. (1991). Common precursors for neural and mesectodermal derivatives in the cephalic neural crest. Development 112,301 -305.[Abstract]
Billon, N., Jolicoeur, C., Tokumoto, Y., Vennstrom, B. and Raff, M. (2002). Normal timing of oligodendrocyte development depends on thyroid hormone receptor alpha 1 (TRalpha1). EMBO J. 21,6452 -6460.[CrossRef][Medline]
Bradley, A., Evans, M., Kaufman, M. H. and Robertson, E. (1984). Formation of germ-line chimaeras from embryo-derived teratocarcinoma cell lines. Nature 309,255 -256.[CrossRef][Medline]
Brook, F. A. and Gardner, R. L. (1997). The
origin and efficient derivation of embryonic stem cells in the mouse.
Proc. Natl. Acad. Sci. USA
94,5709
-5712.
Caplan, A. I. (1991). Mesenchymal stem cells. J. Orthop. Res. 9,641 -650.[CrossRef][Medline]
Chai, Y., Jiang, X., Ito, Y., Bringas, P., Jr, Han, J., Rowitch, D. H., Soriano, P., McMahon, A. P. and Sucov, H. M. (2000). Fate of the mammalian cranial neural crest during tooth and mandibular morphogenesis. Development 127,1671 -1679.[Abstract]
Dani, C., Smith, A. G., Dessolin, S., Leroy, P., Staccini, L., Villageois, P., Darimont, C. and Ailhaud, G. (1997). Differentiation of embryonic stem cells into adipocytes in vitro. J. Cell Sci. 110,1279 -1285.[Abstract]
Doetschman, T. C., Eistetter, H., Katz, M., Schmidt, W. and Kemler, R. (1985). The in vitro development of blastocyst-derived embryonic stem cell lines: formation of visceral yolk sac, blood islands and myocardium. J. Embryol. Exp. Morphol. 87,27 -45.[Medline]
Dulac, C., Cameron-Curry, P., Ziller, C. and Le Douarin, N. M. (1988). A surface protein expressed by avian myelinating and nonmyelinating Schwann cells but not by satellite or enteric glial cells. Neuron 1,211 -220.[CrossRef][Medline]
Dupin, E., Baroffio, A., Dulac, C., Cameron-Curry, P. and Le
Douarin, N. M. (1990). Schwann-cell differentiation in clonal
cultures of the neural crest, as evidenced by the anti-Schwann cell myelin
protein monoclonal antibody. Proc. Natl. Acad. Sci.
USA 87,1119
-1123.
Dupin, E., Creuzet, S. and Le Douarin, N. M. (2006). The contribution of the neural crest to the vertebrate body. Adv. Exp. Med. Biol. 589,96 -119.[Medline]
Evans, M. J. and Kaufman, M. H. (1981). Establishment in culture of pluripotential cells from mouse embryos. Nature 292,154 -156.[CrossRef][Medline]
Fauquet, M. and Ziller, C. (1989). A monoclonal
antibody directed against quail tyrosine hydroxylase: description and use in
immunocytochemical studies on differentiating neural crest cells.
J. Histochem. Cytochem.
37,1197
-1205.
Fehling, H. J., Lacaud, G., Kubo, A., Kennedy, M., Robertson,
S., Keller, G. and Kouskoff, V. (2003). Tracking mesoderm
induction and its specification to the hemangioblast during embryonic stem
cell differentiation. Development
130,4217
-4227.
Ferguson, C. A. and Graham, A. (2004). Redefining the head-trunk interface for the neural crest. Dev. Biol. 269,70 -80.[CrossRef][Medline]
Green, H. and Meuth, M. (1974). An established pre-adipose cell line and its differentiation in culture. Cell 3,127 -133.[CrossRef][Medline]
Green, H. and Kehinde, O. (1975). An established preadipose cell line and its differentiation in culture. II. Factors affecting the adipose conversion. Cell 5, 19-27.[Medline]
Green, H. and Kehinde, O. (1976). Spontaneous heritable changes leading to increased adipose conversion in 3T3 cells. Cell 7,105 -113.[CrossRef][Medline]
Greenberg, A. S., Egan, J. J., Wek, S. A., Garty, N. B.,
Blanchette-Mackie, E. J. and Londos, C. (1991). Perilipin, a
major hormonally regulated adipocyte-specific phosphoprotein associated with
the periphery of lipid storage droplets. J. Biol.
Chem. 266,11341
-11346.
Hamburger, V. and Hamilton, H. L. (1951). A series of normal stages in the development of the chick embryo. J. Morphol. 88,49 -92.[CrossRef]
Hausman, D. B., DiGirolamo, M., Bartness, T. J., Hausman, G. J. and Martin, R. J. (2001). The biology of white adipocyte proliferation. Obes. Rev. 2, 239-254.[CrossRef][Medline]
Hausman, G. J. and Kauffman, R. G. (1986). The
histology of developing porcine adipose tissue. J. Anim.
Sci. 63,642
-658.
Hausman, G. J., Campion, D. R. and Martin, R. J. (1980). Search for the adipocyte precursor cell and factors that promote its differentiation. J. Lipid Res. 21,657 -670.[Medline]
Hausman, G. J., Wright, J. T., Jewell, D. E. and Ramsay, T. G. (1990). Fetal adipose tissue development. Int. J. Obes. 14 Suppl. 3, 177-185.[Medline]
Hole, N. (1999). Embryonic stem cell-derived haematopoiesis. Cells Tissues Organs 165,181 -189.[CrossRef][Medline]
Jiang, X., Rowitch, D. H., Soriano, P., McMahon, A. P. and Sucov, H. M. (2000). Fate of the mammalian cardiac neural crest. Development 127,1607 -1616.[Abstract]
Joseph, N. M., Mukouyama, Y. S., Mosher, J. T., Jaegle, M.,
Crone, S. A., Dormand, E. L., Lee, K. F., Meijer, D., Anderson, D. J. and
Morrison, S. J. (2004). Neural crest stem cells undergo
multilineage differentiation in developing peripheral nerves to generate
endoneurial fibroblasts in addition to Schwann cells.
Development 131,5599
-5612.
Kawaguchi, J., Mee, P. J. and Smith, A. G. (2005). Osteogenic and chondrogenic differentiation of embryonic stem cells in response to specific growth factors. Bone 36,758 -769.[Medline]
Keller, G. (2005). Embryonic stem cell
differentiation: emergence of a new era in biology and medicine.
Genes Dev. 19,1129
-1155.
Lafontan, M. and Berlan, M. (2003). Do regional differences in adipocyte biology provide new pathophysiological insights? Trends Pharmacol. Sci. 24,276 -283.[CrossRef][Medline]
Lahav, R., Dupin, E., Lecoin, L., Glavieux, C., Champeval, D.,
Ziller, C. and Le Douarin, N. M. (1998). Endothelin 3
selectively promotes survival and proliferation of neural crest-derived glial
and melanocytic precursors in vitro. Proc. Natl. Acad. Sci.
USA 95,14214
-14219.
Keller, G. M. (1995). In vitro differentiation of embryonic stem cells. Curr. Opin. Cell Biol. 7, 862-869.[CrossRef][Medline]
Kuhlbrodt, K., Herbarth, B., Sock, E., Hermans-Borgmeyer, I. and
Wegner, M. (1998). Sox10, a novel transcriptional modulator
in glial cells. J. Neurosci.
18,237
-250.
Le Douarin, N. M. and Teillet, M.-A. (1974). Experimental analysis of the migration and differentiation of neuroblasts of the autonomic nervous system and of neurectodermal mesenchymal derivatives, using a biological cell marking technique. Dev. Biol. 41,163 -184.
Le Douarin, N. M. and Kalcheim, C. (1999). The Neural Crest. Cambridge: Cambridge University Press.
Le Douarin, N. M., Creuzet, S., Couly, G. and Dupin, E.
(2004). Neural crest cell plasticity and its limits.
Development 131,4637
-4650.
Le Lievre, C. S. and Le Douarin, N. M. (1975). Mesenchymal derivatives of the neural crest: analysis of chimaeric quail and chick embryos. J. Embryol. Exp. Morphol. 34,125 -154.[Medline]
Li, M., Pevny, L., Lovell-Badge, R. and Smith, A. (1998). Generation of purified neural precursors from embryonic stem cells by lineage selection. Curr. Biol. 8, 971-974.[CrossRef][Medline]
Lumsden, A. G. (1988). Spatial organization of the epithelium and the role of neural crest cells in the initiation of the mammalian tooth germ. Dev. Suppl. 103,155 -169.
MacDougald, O. A. and Lane, M. D. (1995). Adipocyte differentiation. When precursors are also regulators. Curr. Biol. 5,618 -621.[CrossRef][Medline]
Mandrup, S. and Lane, M. D. (1997). Regulating
adipogenesis. J. Biol. Chem.
272,5367
-5370.
Martin, G. R. (1981). Isolation of a
pluripotent cell line from early mouse embryos cultured in medium conditioned
by teratocarcinoma stem cells. Proc. Natl. Acad. Sci.
USA 78,7634
-7638.
Matsuoka, T., Ahlberg, P. E., Kessaris, N., Iannarelli, P., Dennehy, U., Richardson, W. D., McMahon, A. P. and Koentges, G. (2005). Neural crest origins of the neck and shoulder. Nature 436,347 -355.[CrossRef][Medline]
McGonnell, I. M. and Graham, A. (2002). Trunk neural crest has skeletogenic potential. Curr. Biol. 12,767 -771.[CrossRef][Medline]
Mizuseki, K., Sakamoto, T., Watanabe, K., Muguruma, K., Ikeya,
M., Nishiyama, A., Arakawa, A., Suemori, H., Nakatsuji, N., Kawasaki, H. et
al. (2003). Generation of neural crest-derived peripheral
neurons and floor plate cells from mouse and primate embryonic stem cells.
Proc. Natl. Acad. Sci. USA
100,5828
-5833.
Montague, C. T. and O'Rahilly, S. (2000). The perils of portliness: causes and consequences of visceral adiposity. Diabetes 49,883 -888.[Abstract]
Mountford, P., Zevnik, B., Duwel, A., Nichols, J., Li, M., Dani,
C., Robertson, M., Chambers, I. and Smith, A. (1994).
Dicistronic targeting constructs: reporters and modifiers of mammalian gene
expression. Proc. Natl. Acad. Sci. USA
91,4303
-4307.
Nakamura, H. and Ayer-le Lievre, C. S. (1982). Mesectodermal capabilities of the trunk neural crest of birds. J. Embryol. Exp. Morphol. 70,1 -18.[Medline]
Nataf, V., Mercier, P., Ziller, C. and Le Douarin, N. M. (1993). Novel markers of melanocyte differentiation in the avian embryo. Exp. Cell Res. 207,171 -182.[CrossRef][Medline]
Negrel, R., Grimaldi, P. and Ailhaud, G.
(1978). Establishment of preadipocyte clonal line from epididymal
fat pad of ob/ob mouse that responds to insulin and to lipolytic hormones.
Proc. Natl. Acad. Sci. USA
75,6054
-6058.
Okabe, S., Forsberg-Nilsson, K., Spiro, A. C., Segal, M. and McKay, R. D. (1996). Development of neuronal precursor cells and functional postmitotic neurons from embryonic stem cells in vitro. Mech. Dev. 59,89 -102.[CrossRef][Medline]
Otto, T. C. and Lane, M. D. (2005). Adipose development: from stem cell to adipocyte. Crit. Rev. Biochem. Mol. Biol. 40,229 -242.[CrossRef][Medline]
Owen, M. (1988). Marrow stromal stem cells. J. Cell Sci. Suppl. 10,63 -76.[Medline]
Phillips, B. W., Vernochet, C. and Dani, C. (2003). Differentiation of embryonic stem cells for pharmacological studies on adipose cells. Pharmacol. Res. 47,263 -268.[CrossRef][Medline]
Pittenger, M. F., Mackay, A. M., Beck, S. C., Jaiswal, R. K.,
Douglas, R., Mosca, J. D., Moorman, M. A., Simonetti, D. W., Craig, S. and
Marshak, D. R. (1999). Multilineage potential of adult human
mesenchymal stem cells. Science
284,143
-147.
Rodriguez, A. M., Elabd, C., Delteil, F., Astier, J., Vernochet, C., Saint-Marc, P., Guesnet, J., Guezennec, A., Amri, E. Z., Dani, C. et al. (2004). Adipocyte differentiation of multipotent cells established from human adipose tissue. Biochem. Biophys. Res. Commun. 315,255 -263.[CrossRef][Medline]
Rohwedel, J., Guan, K. and Wobus, A. M. (1999). Induction of cellular differentiation by retinoic acid in vitro. Cells Tissues Organs 165,190 -202.[CrossRef][Medline]
Rosen, E. D. and MacDougald, A. (2006). Adipocyte differentiation from the inside out. Nat. Rev. 7,885 -896.[CrossRef]
Rosen, E. D., Walkey, C. J., Puigserver, P. and Spiegelman, B.
M. (2000). Transcriptional regulation of adipogenesis.
Genes Dev. 14,1293
-1307.
Santagati, F. and Rijli, F. M. (2003). Cranial neural crest and the building of the vertebrate head. Nat. Rev. Neurosci. 4,806 -818.[Medline]
Shah, N. M., Groves, A. K. and Anderson, D. J. (1996). Alternative neural crest cell fates are instructively promoted by TGFbeta superfamily members. Cell 85,331 -343.[CrossRef][Medline]
Smith, A. G., Heath, J. K., Donaldson, D. D., Wong, G. G., Moreau, J., Stahl, M. and Rogers, D. (1988). Inhibition of pluripotential embryonic stem cell differentiation by purified polypeptides. Nature 336,688 -690.[CrossRef][Medline]
Spiegelman, B. M. and Farme
