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First published online 24 July 2008
doi: 10.1242/dev.020453
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1 Departments of Pathology and Immunology, Washington University School of
Medicine, 660 South Euclid Avenue, St Louis, MO 63110, USA.
2 Department of Developmental Biology, Washington University School of Medicine,
660 South Euclid Avenue, St Louis, MO 63110, USA.
Author for correspondence (e-mail:
stappenb{at}pathology.wustl.edu)
Accepted 25 June 2008
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Fibroblast growth factor 9 (Fgf9), Gut development, Tgfβ signaling, Epithelial-mesenchymal crosstalk, Mesenchymal stem cells, Follistatin, Mouse
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). From E15.5 to
E18.5, a proximal-to-distal wave of cytodifferentiation converts the
epithelium to a simple columnar histology, organized into nascent villi with
intervillus regions (Schmidt et al.,
1988; Calvert and Pothier,
1990). A less appreciated developmental change that also occurs
during this time period is a remarkable cephalo-caudal lengthening of the
organ. Early in embryonic development, the lengthening of the small intestine
occurs in proportion to total body length. However, in the latter third of
embryogenesis, the rate of intestinal lengthening far outpaces that of the
body, becoming exponential in both humans and rodents
(Sbarbati, 1979;
Weaver et al., 1991). As
postnatal development commences, the rate of intestinal lengthening slows with
a resumption of a linear relationship between the two.
In humans, the issue of intestinal lengthening becomes clinically important
in individuals with short bowel syndrome
(Ladefoged et al., 1996). This
condition most often occurs when a section of intestine is surgically removed
postnatally for a variety of conditions in which a portion of the small
intestine is no longer viable (Nightingale
and Lennard-Jones, 1993). Following resection, an adaptive
response occurs in which the diameter of the small intestine increases in
response to loss of surface area. In mouse models of this process, crypts and
villi lengthen owing to increased epithelial progenitor proliferation and
decreased epithelial turnover (Erwin et
al., 2006; Dekaney et al.,
2007; Wang et al.,
2007). However, overall small intestinal length is never
regenerated to any appreciable degree. Because the adaptive response is often
inadequate to restore sufficient surface area for absorption, individuals with
short bowel syndrome are often treated with total parenteral nutrition, which
has a 3-year survival rate of 65-80%
(Howard et al., 1995). Thus,
understanding the exponential lengthening of the small intestine in the mouse
during late-stage embryonic development has potential clinical
implications.
However, little is known regarding the developmental pathways that regulate
small intestinal lengthening during this crucial period. Most well-studied
major developmental pathways play important roles in the proper patterning of
the crypt-villus axis or the development of the outer muscle wall during
embryogenesis. The formation of villus structures during the late prenatal
period is dependent on an intact Pdgf pathway
(Karlsson et al., 2000). The
canonical Wnt pathway is essential for the regulation of epithelial stem cell
homeostasis (Korinek et al.,
1998). The Bmp arm of the Tgf-β superfamily regulates crypt
census, and thus indirectly impacts stem cell number in the small intestine
(Haramis et al., 2004). The
ultimate cell fate decisions of committed stem cell daughters are regulated at
least in part by the Notch signaling pathway
(van Es et al., 2005;
Fre et al., 2005;
Vooijs et al., 2007). Finally,
the outer muscular layer of the intestine depends upon hedgehog signals for
its proper formation (Ramalho-Santos et
al., 2000).
More recent studies have begun to elucidate how these disparate
developmental pathways interact during intestinal embryogenesis. For example,
Wnt and Notch perform synergistic roles in the maintenance of proliferation
and homeostasis of the epithelial stem cell compartment
(van Es et al., 2005). Wnt and
Indian hedgehog play antagonistic roles in the determination of epithelial
cell differentiation in the colon (van den
Brink et al., 2004). Finally, the Pten/Akt family interacts with
Wnt/β-catenin signaling to regulate epithelial homeostasis
(He et al., 2004;
He et al., 2007). Thus, the
interaction of multiple developmental pathways plays a crucial role in several
aspects of small intestinal development.
Fibroblast growth factors (Fgfs) comprise a large family of polypeptide
growth factors that are found in organisms ranging from C. elegans to
humans (Ornitz and Itoh,
2008). Specifically, Fgf9 plays a key role in embryonic
development of the lung, heart, cecum and testes
(Colvin et al., 2001a;
Colvin et al., 2001b;
Lavine et al., 2005;
White et al., 2006;
Zhang et al., 2006). However,
Fgf9 expression is not limited to these organs. We and others have previously
shown that this growth factor is also expressed throughout the length of the
intestine during late stage embryogenesis
(Zhang et al., 2006;
Sala et al., 2006). A careful
study of the role of Fgf9 in the development of the intestine outside of the
cecum has not been undertaken.
Here, by using mice that lack mesenchymal Fgf9 signaling (either through loss of the Fgf9 ligand or its mesenchymal receptors), we show that Fgf9 signals are required for longitudinal growth of the small intestine during late-stage embryogenesis. Fgf9 signaling to the intestinal mesenchyme regulates proliferation of fibroblasts, which in turn appears to drive gut lengthening. Additionally, we found that Fgf9 inhibits the fibroblast to myofibroblast transition in the small intestinal mesenchyme during embryogenesis. Interestingly, this process is regulated by an indirect cellular mechanism that controls the Tgfβ signaling pathway in fibroblasts.
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MATERIALS AND METHODS |
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SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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;
Trokovic et al., 2003;
White et al., 2006;
Yu et al., 2003) and all are
on a C57Bl/6 background.
Histochemistry and BrdU labeling
For embryo collections, time-mated females were sacrificed at E14.5-E18.5,
1 hour after an intraperitoneal injection of bromodeoxyuridine (BrdU, 120
µg/gm, Sigma). The GI tract was isolated from embryos, pinned on black wax,
fixed in 2% paraformaldehyde for 4 hours and processed as previously described
(Stappenbeck and Gordon,
2000). Tissue samples were embedded in paraffin and 5 µm serial
sections were prepared. Sections were then stained with Hematoxylin and Eosin.
Adjacent, unstained sections were used for immunohistochemical studies (see
below).
Paraffin sections were processed for immunohistochemistry using an antibody to BrdU (BD Biosciences). Sections were counterstained with Hematoxylin. Proliferation was measured in three cellular compartments in the proximal and distal small intestines as illustrated in Fig. 2 by scoring the number of BrdU-labeled nuclei to the number of total nuclei in each compartment.
Immunohistochemistry
Slide preparations for immunohistochemistry varied depending on primary
antibody used. For antibodies directed against 
-SMA (clone 1a4;
Sigma-Aldrich; 1:2000 dilution), prolyl 4-hydroxylase (clone ER-TR7; BMA
Biomeidcal; 1:100), desmin (clone ZC18; Zymed; 1:100), I-Fabp, Fgfr1 (Santa
Cruz,; 1:50), Fgfr2 (Santa Cruz; 1:100), Fst (Santa Cruz; 1:10), Fstl1 (Santa
Cruz; 1:10), Pecam1 (rat anti-mouse CD31; clone MEC13.3, BD Pharmingen;
1:100), p-Smad1/5/8 (Cell Signaling; 1:100) and p-Smad2/3 (Cell Signaling;
1:100), embryonic small intestines were embedded in paraffin and sectioned at
5 µm prior to staining. Sections were then deparaffinized in three washes
of xylene, and rehydrated in isopropanol. Antigen retrieval was performed by
varying methods depending on primary antibody used. For I-Fabp, Fst, Fstl1,
p-Smad2/3 and p-Smad1/5/8, slides were heated at 95°C for 20 minutes in 10
mM sodium citrate buffer (pH=6.0). Slides were then washed in double distilled
H2O for 5 minutes prior to washing in blocking buffer (composed of
1% BSA, 0.1% Triton in PBS) for 20 minutes. For primary antibody to Pecam1,
antigen retrieval was carried out by a 10-minute incubation with a 1%
chymotrypsin solution, followed by a 10 minute wash in double distilled
H2O prior to application of blocking buffer. Antibodies to
-SMA, prolyl 4-hydroxylase, Fgfr2 and desmin required no antigen
retrieval.
For antibodies to p-Erk1/2 (Cell Signaling; 1:100), CD29 (BD Biosciences; 1:100), CD44 (BD; 1:100), CD45 (BD; 1:100), CD54 (BD; 1:100), CD105 (BD; 1:100) and CD106 (BD; 1:100), embryonic small intestines were embedded in OCT compound (Sakura) and immediately frozen in liquified Cytocool (Richard-Allan Scientific). Frozen sections of 5 µm were then fixed for 5 minutes in 4% paraformaldehyde, rinsed in PBS and then immersed in blocking buffer for 20 minutes prior to incubation with primary antibody. Primary antibodies were incubated at 4°C overnight, followed by secondary antibody incubation for 1 hour at 24°C using a 1:500 dilution. Slides were then counterstained with bis-benzimide and coversliped with a 1:1 glycerol/PBS solution. Sections were viewed with a Zeiss Axiovert 200 with Axiocam MRM camera and Apotome optical sectioning slider.
Isolation of embryonic small intestinal mesenchymal stem cells
A single wild-type E18.5 small intestine was minced with a razor blade
followed by treatment with collagenase (Gibco) in Dulbecco's modified eagle
medium (DMEM) containing 10 mM HEPES buffer for 1 hour at 37°C. After
passage through a 70 µm filter, the cell suspension was centrifuged for 10
minutes at 400 g. The cell pellet was then suspended in DMEM
containing 10% FBS and plated on standard plastic tissue culture plates. After
incubation for 2 hours at 37°C, the growth media containing nonadherent
cells was removed and discarded. Adherent cells were replenished with fresh
media and passaged once weekly.
To differentiate MSCs into adipocytes, isolated cultured embryonic small intestinal MSCs were treated with 10-8 M dexamethasone and 5 µg/ml insulin for 21 days. Cells were then stained with Oil Red O to verify presence of lipid stores. To differentiate MSCs into osteocytes, MSCs were treated with 10-8 M dexamethasone, 5 µg/ml ascorbic acid 2-phosphate, 10 mM β-glycerophosphate for 21 days. Cells were then stained with Alizarin Red to verify presence of Ca2+ deposits in the extracellular matrix.
FACS analysis
Cultured cells were treated for 5 minutes with a trypsin/EDTA solution.
Cells were pelleted and fixed in 2% formaldehyde in PBS for 10 minutes at
37°C. Cells were then chilled at 4°C for 1 minute, and pelleted by
centrifugation at 250 g for 5 minutes. Cells were then
permeabilized by 30 minute incubation in 90% methanol at 4°C. Cells were
then washed twice with PBS, and then re-suspended in 1% BSA in PBS, containing
the primary antibody for 1 hour at 24°C. The cells were then washed twice
in PBS before incubation in fluorescently-conjugated secondary antibody
(diluted 1:500 in 1% BSA in PBS for 1 hour). Cells were then rinsed in PBS,
before finally suspending in 1% BSA in PBS for flow cyotometric analysis. All
analysis was performed on a FACSCalibur flow cytometer and analyzed with
FlowJo software.
Immunoblotting
MSCs were serum starved for 2 days in DMEM containing 0.5% FCS. Following
starvation, 10 ng/ml Fgf9 (Peprotech) was added to the growth media. Following
treatment, cells were lysed in RIPA buffer for 5 minutes on ice and then
centrifuged at 14,000 g for 10 minutes. Protein content of the
supernatant was quantified using a Bio-Rad Dc colorimetric protein
assay (Bio-Rad). Protein (10 µg) was then reduced and loaded into 10%
Bis-Tris gels and electrophoresed. Samples were then transferred onto PVDF
membranes and blocked in a solution containing 4% BSA (gels and membranes from
Invitrogen), 0.1% Tween in TBS for 1 hour at 24°C. Blots were then
incubated in primary antibody overnight, washed in TBST and then probed with
HRP-conjugated secondary antibody (Bio-Rad) for 1 hour at 24°C before
development with the SuperSignal West Pico chemiluminescent kit (Pierce).
qRT-PCR analysis
Cells were serum starved and treated with Fgf9 as above. Total cellular RNA
was collected and purified using a Qiagen RNeasy mini kit. cDNAs were then
synthesized using Superscript III reverse transcriptase (Invitrogen) and
random primers according to the manufacturer's protocol. qRT-PCR that was
performed in triplicate for each sample using SYBR-green master mix
(Invitrogen). The following primers were used: 18S
(5'-CATTCGAACGTCTGCCCTATC and 5'-CCTGTGCCTTCCTTGGA), Fst
(5'-TACTGTGTGACCTGTAATCGG-3' and
5'-TGATACACTTTCCCTCATAGGC-3') and Fstl1
(5'-CACGGCGAGGAGGAACCTA-3' and
5'-TCTTGCCATTACTGCCACACA-3').
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) was
co-cultured with MSCs using a transwell system (Corning). MSCs
(5.0x104) were plated on the upper chamber, which was
separated from the lower chamber by a membrane containing 0.4 µm pores. The
lower chamber was seeded with 1.0x104 myofibroblasts.
Experiments were performed was carried out in DMEM containing 0.5% FCS. When
confluent (
3 days), myofibroblasts were collected for FACS analysis as
described above. Fst was obtained from R&D Systems for treatment of
Mic216.
Laser capture microdissection
Ten thousand cells were separately procured from the mesenchyme and
epithelium of control and DCR1R2 E18.5 embryonic small intestines as
previously described (Pull et al.,
2005). DNA was isolated by incubating the captured cells with
proteinase K in TE buffer at 42°C for 10 hours. DNA was amplified by 40
cycles of PCR using primers as previously described
(Yu et al., 2003).
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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).
Fgf9 expression persists in a similar distribution during late stage
embryogenesis (see Fig. S1A in the supplementary material).
Interestingly, the defect in small intestinal length in the
Fgf9-/- embryos was not accompanied by an alteration in
its caliber. We measured caliber at the same relative position along the
length of each individual small intestine
(Fig. 1C). Thus, the
Fgf9-/- small intestine was not a miniaturized version of
controls, but instead was growth altered in only one dimension (length). In
addition, more severe developmental defects such as small intestinal atresia,
duplication or malrotation were never observed as have been previously noted
as frequent malformations in other gene knockouts such as
Shh-/- and Ihh-/-
(Ramalho-Santos et al.,
2000).
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Analysis of the intestinal epithelium provides a sensitive readout for
interaction of Fgf9 with several major developmental pathways. Proliferative
epithelial progenitors (as indicated by the presence of BrdU-positive cells)
were present only in the intervillus epithelium of control and
Fgf9-/- mice (Fig.
1E), indicating that the balance of Wnt and Bmp signaling was not
significantly perturbed. The proper allocation of secretory and absorptive
lineages is controlled by Notch signaling
(van Es et al., 2005). The
small intestinal epithelium of Fgf9-/- embryos contained
similar proportions of goblet cells, enteroendocrine cells and enterocytes as
those seen in their control littermates at E18.5
(Fig. 1F), indicating no severe
defects were present in this pathway.
However, we found that Fgf9-/- small intestines showed
a subtle defect in enterocytic differentiation at E16.5 and E18.5. These cells
expressed lower amounts of intestinal fatty acid binding protein (I-fabp)
(Cohn et al., 1992) in the
proximal small intestine, as well as lower levels of ileal lipid-binding
protein (Ilbp) in the distal small intestine
(Fig. 1G; data not shown for
E16.5). However, the enterocytes displayed no obvious alterations in polarity,
apical junctions or size in either location of the small intestine.
Proliferation of small intestinal mesenchymal fibroblasts is driven by Fgf9 during late stage embryogenesis
Cell proliferation in the small intestine was quantified by BrdU labeling.
We analyzed the three longitudinal layers of this organ (the intervillus
epithelium, its underlying mesenchyme, and the muscularis propria) in control
and Fgf9-/- embryos as one or more of these layers could
drive intestinal elongation (Fig.
2A). We determined the proliferative index (PI) for each
compartment as defined by the number of nuclei incorporating BrdU divided by
the total number of nuclei in that compartment. We performed this analysis in
both the proximal and distal small intestine at three developmental time
points (E14.5, E16.5 and E18.5). The average PI for the intervillus epithelium
and for the mucularis propria was not significantly different when comparing
control and Fgf9-/- embryos at any location or timepoint
(Fig. 2B,C shows data for the
proximal small intestine at all timepoints). However, the PI for the
mesenchyme was significantly reduced (P<0.01) in
Fgf9-/- embryos when compared with their littermate
controls in both the proximal and distal small intestine at all time points
between E14.5 and E18.5 (Fig.
2C). To examine the effects of Fgf9 signaling loss on cell death,
TUNEL stained sections of control and Fgf9-/-small
intestines were examined. We observed no significant differences in apoptosis
in any cellular compartment at any time point (E14.5-E18.5) during late-stage
embryogenesis (data not shown).
The embryonic intestinal mesenchyme contains multiple cell types that may
increase proliferation in response to Fgf9. The vast majority of the
mesenchyme in E18.5 control small intestines was composed of prolyl
4-hydroxylase-positive, smooth muscle actin (-SMA)-negative
fibroblasts (Powell et al.,
2005). Two other smaller populations included Pecam1-positive
endothelial cells and CD45-positive, F4/80-positive monocyte-derived cells. As
expected, the monocyte-derived cells did not proliferate to any appreciable
levels in the mesenchyme of these embryos (data not shown). In the absence of
Fgf9, fibroblasts exhibited a significant decrease in proliferation
(P<0.01; Fig. 2D),
whereas endothelial cells maintained a proliferative rate similar to controls.
These findings suggested that mesenchymal fibroblast proliferation contributed
to small intestinal elongation during embryogenesis.
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; White et al.,
2006; Yu and Ornitz,
2008). To examine non-epithelial targets for Fgf9 and potential
redundancy among Fgfrs, we conditionally inactivated floxed alleles of both
Fgfr1 and Fgfr2 (Pirvola
et al., 2002; Herbert et al.,
2003; Trokovic et al.,
2003; White et al.,
2006) using the Dermo1-Cre allele
(Yu et al., 2003). To define
the targeted lineages in the small intestine, Dermo1-Cre
mice were mated with ROSA26 reporter mice
(Soriano, 1999). At E14.5, the
mesenchyme and muscularis propria expressed Dermo1-Cre, as indicated
by positive X-Gal staining in these regions
(Fig. 3A). All mesenchymal
lineages, including fibroblasts, endothelial cells and macrophages contained
detectable β-galactosidase whereas all epithelial cells were negative
(Fig. 3A,B). Dermo1-Cre; Fgfr1+/-: Fgfr2+/- mice were mated with mice containing two floxed alleles of both Fgfr1 and Fgfr2 to generate mice with the genotype Dermo1-Cre; Fgfr1loxP/-;Fgfr2loxP/- (DCR1R2). This breeding scheme also produced Dermo1-Cre Fgfr1loxP/- Fgfr2LoxP/+ (DCR1), Dermo1-Cre; Fgfr1loxP/+; Fgfr2loxP/- (DCR2) and control mice that lack Dermo1-Cre. We confirmed by immunofluorescence using Fgfr1 and Fgfr2 antisera that DCR1R2 mice contained neither Fgfr1 nor Fgfr2 expression in mesenchymal cells of the small intestine whereas the levels of expression in the epithelium were not perceptibly changed (data not shown). In addition, we performed PCR of laser capture microdissected epithelium and mesenchyme from control and DCR1R2 mice. We could only detect PCR products that indicted the presence of a recombined allele for Fgfr1 and Fgfr2 in the mesenchyme of DCR1R2 mice (Fig. 3C).
Late stage DCR1R2 embryos contained shortened small intestines when compared with littermate controls (P<0.01; Fig. 3D), whereas the crown-rump length was not significantly altered (data not shown). The gross morphology, histological analysis and epithelial differentiation showed defects similar to those described in the Fgf9-/- mouse (data not shown). DCR1 and DCR2 embryos both showed milder but significant reductions in gut length (P<0.01; Fig. 3D). The effects of loss of Fgfr1 and Fgfr2 were additive as the small intestinal lengths of DCR1R2 mice were significantly different than both DCR1 and DCR2 (P<0.01; Fig. 3D). Proliferation data for the DCR1R2 mice showed that these embryos displayed a similar phenotype to Fgf9-/- embryos. Only the mesenchyme contained a significant (P<0.01) decrease in proliferation (Fig. 3E). These data suggest that Fgf9 acts primarily through both Fgfr1 and Fgfr2 in the mesenchyme of the developing small intestine to drive mesenchymal proliferation and small intestinal lengthening.
Fgf9 regulates Tgfβ signaling to control mesenchymal differentiation
As the small intestine transitions to the postnatal period, the majority of
the mesenchymal fibroblasts are replaced by myofibroblasts, which are
characterized by elevated expression of -SMA and their lack of
detectable desmin expression (Powell et
al., 2005). To examine mesenchymal differentiation, sections of
the small intestine of control and Fgf9-/- embryos were
stained for -SMA and desmin (Fig.
4A-F; see Fig. S1B,C in the supplementary material).
Interestingly, at E14.5, E16.5 and E18.5, Fgf9-/- small
intestines displayed a profound increase in -SMA expression in the
mesenchyme when compared with the same region in their littermate controls.
These -SMA-positive cells did not express detectable levels of desmin,
consistent with myofibroblast differentiation (see Fig. S1B,C in the
supplementary material).
Tgfβ signals are well known to drive fibroblast to myofibroblast
transitions in other model systems, such as wound repair in skin
(Powell et al., 2005). Smad2
and Smad3 serve as intermediate molecules in the Tgfβ signaling cascade
that are phosphorylated in response to active binding of Tgfβ and activin
ligands to their receptors, while Bmp signaling phosphorylates and activates
Smad1, Smad5 and Smad8 (Waite et al., 2003). To assess the level of Tgfβ
signaling in intestinal mesenchyme of control and Fgf9-/-
mice, we examined phospho (p)-Smad2/3 expression using antibodies specific for
the active phosphorylated forms. At E14.5, E16.5 and E18.5, both
Fgf9-/- and control embryos showed high level expression
of p-Smad2/3 in the nuclei of villus epithelial cells
(Fig. 4G-L). At all three of
these time points, Fgf9-/- small intestines showed robust
p-Smad2/3 staining in nearly all mesenchymal cells, whereas control mice
showed only scattered p-Smad2/3-positive cells. By comparison, absence of Fgf9
did not similarly alter the levels of p-Smad1/5/8 in the mesenchyme (see Fig.
S1D,E in the supplementary material). The staining pattern of p-Smad1/5/8 in
control embryos corresponded to previously shown patterns in adult small
intestines (Haramis et al.,
2004).
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; Que et al.,
2007). If the mechanism of Fgf9 action was exclusively directed
towards fibroblasts, then we would predict that p-Erk1/2 stained sections
should label these cells. However, stained sections of E18.5 small intestines
from control mice showed that only a small subset of mesenchymal cells showed
detectable p-Erk1/2 staining (Fig.
5A, denoted as high p-Erk1/2). We found that these high p-Erk1/2
cells were located closer to the epithelium than the muscle wall and were
typically clustered at the base of villi. Significantly fewer of these high
p-Erk1/2 mesenchymal cells were detected in Fgf9-/- small
intestines (Fig. 5B,C), as well
as in DCR1R2 mice (see Fig. 2A in the supplementary material). In
control small intestines, the high p-Erk1/2 mesenchymal cells did not express
well-characterized markers for either of the less abundant mesenchymal
lineages [CD45-positive macrophages and Pecam1-positive endothelial cells
(Fig. 5D,E)]. In addition, the
p-Erk1/2 high cells were neither BrdU positive
(Fig. 5F) nor prolyl
4-hydroxylase positive (data not shown) in control small intestines,
suggesting these cells were distinct from the proliferative fibroblasts in the
mesenchyme.
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). We screened for
mRNA expression of these candidates in the whole small intestines of wild-type
mice by RT-PCR. We detected expression of Fst and Fstl (data not shown). Both
of these genes were previously identified using expression profiles as
preferentially expressed in the mesenchyme (versus the epithelium) of E15.5
small intestines (Li et al.,
2007). We then evaluated expression of Fst and Fstl1 in control
embryonic small intestines by immunohistochemistry. At E14.5 we found
scattered mesenchymal Fst- and Fstl1-positive cells (see, for example, Fig.
S2B in the supplementary material). At later times in embryonic development,
we observed clusters of both Fst-and Fstl 1-positive mesenchymal cells near
the base of villi of control (Fig.
5G,H) but not Fgf9-/- mice. Double-label
immunofluorescence for p-Erk1/2 with either Fst or Fstl1 showed that the
subset of stromal cells that expressed high levels of pErk1/2 showed
colocalization with both of these proteins
(Fig. 5G,H). In addition, Fgfr2
colocalized with both p-Erk and Fst, indicating that epithelial Fgf9 signals
through mesenchymal receptors in cells that contain high levels of Fst and
Fstl1 (see Fig. S2C,D in the supplementary material).
The p-Erk/Fst/Fstl1-expressing cells near the villus base are mesenchymal stem cells
One mesenchymal cell type that cannot, as yet, be identified on tissue
sections with a single marker is the mesenchymal stem cell (MSC). Though these
cells were originally considered to function as the source of many different
types of mesenchymal lineages, recent work has suggested that they may play a
role in signaling, particularly in injury repair
(Gnecchi et al., 2006). Most
mouse tissues from either adults (da Silva
Meirelles et al., 2006; Wang
et al., 2006a) or embryos (Lee
and Tarantal, 2006) contain MSCs that can be readily isolated by
well-established protocols (da Silva
Meirelles et al., 2006). Therefore, we isolated MSCs from late
stage wild-type embryonic small intestines using these same protocols.
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; Wang et al.,
2006a). Last, the iMSCs differentiated into either adipocytes or
osteocytes when cultured under inductive conditions (see Fig. S3A,B in the
supplementary material) (da Silva Meirelles
et al., 2006). The iMSCs also displayed similar characteristics as the high p-Erk/Fst/Fstl1 small intestinal mesenchymal cells. When cultured alone in 10% serum, the iMSCs expressed much higher relative amounts of mRNA encoding Fst (ranging from 10-128 fold higher than mRNAs from epithelial, macrophage and myofibroblast cell lines) and Fstl1 (ranging from 47-3500 fold higher than the same set of cell lines; Fig. 6B). Importantly, the iMSCs responded in vitro to Fgf9 stimulation in a time- and dose-dependent manner by increasing p-Erk1/2 expression (see Fig. S3C in the supplementary material; data not shown). We found that Fst was co-expressed with p-Erk1/2 in the iMSCs in vitro (see Fig. S3D in the supplementary material). Furthermore, exogenous Fgf9 increased expression of both Fst and Fstl1 in these iMSCs (Fig. 6C).
To test the hypothesis that the iMSCs could affect myofibroblast
differentiation, we co-cultured them with a line of well characterized
intestinal myofibroblasts (Mic216)
(Plateroti et al., 1998) using
a transwell system. We found that the presence of the iMCSs resulted in a
de-differentiation of myofibroblasts towards fibroblasts, based on decreased
expression of -SMA in the Mic216 cells
(Fig. 6D,E). This finding was
partially recapitulated by the incubation of Mic216 cells with Fst
(Fig. 6E). Mic216 cells did not
significantly alter their SMA expression in response to Fgf9
(Fig. 6E). Last, we performed
double-label immunofluorescence on sections of wild-type small intestines with
p-Erk1/2 and markers of iMSCs. We found that iMSC markers colocalized with the
high p-Erk1/2 mesenchymal cells (Fig,
6F,G; data not shown). iMSCs are also targeted by Dermo1-Cre, as
shown by the colocalization of β-galactosidase with markers of this cell
type (e.g. Fig. 6H). Taken
together, these data support the hypothesis that the
p-Erk/Fst/Fstl1-expressing small intestinal stromal cells are iMSCs.
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DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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). Surprisingly, the response to Fgf9 signaling in the small
intestinal mesenchyme did not occur uniformly. Instead the response to Fgf9
signaling occurred robustly in a select subset of mesenchymal cells that also
expressed high levels of Tgfβ inhibitors, Fst and Fstl1. We found that
these cells most closely resembled iMSCs and that these cells appeared to act
as a cellular intermediary in the signaling of Fgf9 to intestinal mesenchymal
fibroblasts (Fig. 7).
An important feature of Fgf9 signaling in the small intestine is that it
occurs across an epithelial-mesenchymal boundary to regulate elongation of
this organ during late-stage embryogenesis. This type of crosstalk is a common
mechanism of action for fibroblast growth factors. For example, epithelial to
mesenchymal Fgf9 signaling, with reciprocal mesenchymal to epithelial Fgf10
signals, is known to be essential for formation of a cecum
(Burns et al., 2004;
Zhang et al., 2006). In the
lung, epithelial-to-mesenchymal and mesothelial-to-mesenchymal Fgf9 and
reciprocal Fgf10 signaling is required for airway branching
(Bellusci et al., 1997;
Colvin et al., 2001a;
White et al., 2006). Finally,
it has recently been shown that epithelial-to-mesenchymal Fgf9 is necessary
for myocardial growth and proper coronary vascular development
(Lavine et al., 2005). The
function of Fgf9 in the small intestine is somewhat distinct from these other
organs, as no branching of a targeted structure (epithelium or blood vessel)
occurs. Thus far, we have not uncovered a functional reciprocal Fgf partner
that is produced in the intestinal mesenchyme that signals back to the
epithelium. Further studies will be ongoing to determine if this occurs.
Our report suggests that a single molecule (Fgf9) targets the mesenchyme to
affect the elongation of the entire organ during development. Importantly,
communication between the three functional layers of the intestine is
absolutely required, as proper functioning of the organ as a whole requires
each layer to be properly oriented to each other. Thus, extensive
communication across epithelial-mesenchymal boundaries is necessary for proper
intestinal development and adult homeostasis. An unanticipated interaction of
Fgf9 was with Tgfβ but not the Bmp signaling pathways. Previous studies
have demonstrated that development of the mouse small intestine uses
mesenchymal-to-epithelial signaling to control epithelial proliferation and
differentiation during development. Knockout mice for the transcription
factors Nkx2.3 (Pabst et al.,
1999) and Foxl1 (Kaestner et
al., 1997) each show diminished expression of Bmps in the
intestinal mesenchyme. In turn, loss of Bmp signaling has several effects,
including increased epithelial proliferation and dysmorphic villi. In
addition, Wang et al. (Wang et al.,
2006b) have shown that expression of epimorphin in mesenchymal
myofibroblasts is required for proper epithelial maintenance in adult animals,
at least in part by mesenchymal modulation of the Bmp pathway. The
relationship between Fgf9 signaling and Nkx2.3, Foxl1 and epimorphin is
currently unknown but they appear to be distinct pathways as Fgf9 does not
appear to affect Bmps based on the pattern of expression of p-Smad1/5/8 and on
the absence of epithelial/villus defects in Fgf9-/- mice.
The antagonistic relationship of Fgf9 with Tgfβ that we observed in the
developing mouse small intestine is the opposite of the synergistic
interaction that has been extensively documented for Fgfs with Tgfβ in
other systems, most notably the early development of Xenopus and
zebrafish during gastrulation (e.g. Cornell
and Kimelman, 1994; Mathieu et
al., 2004). Antagonistic interactions of Fgf and Tgfβ
signaling have been observed much later in development such as during
melanocyte formation and maturation
(Stocker et al., 1991).
Interestingly, we show that an important downstream function of Fgf9 is the
inhibition of Tgfβ signaling that we hypothesize occurs through an
indirect mechanism using iMSCs as cellular intermediaries. It is known that
Erk, a downstream target of the Fgf receptor, and other MAP kinases, have the
ability to directly modulate the activity of the Tgfβ superfamily through
the targeting of Smad1 molecules for degradation by phosphorylation and later
ubiquination (Kretzschmar et al.,
1997). Interestingly, this ability of Erk to target Smad for
degradation has been shown to be a property of Smads associated with the Bmp
signaling pathway (Smad1, Smad5 and Smad8), and not the Tgfβ and activin
specific Smad2 and Smad3. However, this is still a possible mechanism as both
Smad2 and Smad3 have the ability to be ubiquitinated and degraded following
prolonged activity (Lo and Massagué, 1999). Here, we propose an
indirect mechanism that involves Fgf9 regulation of secreted inhibitors of
Tgfβ. The overall effect of Fgf9 on the small intestinal mesenchyme may
involve other mechanisms whereby Fgf9 targets all fibroblasts and additionally
inhibits Tgfβ signaling in a cell autonomous manner. Such mechanisms need
not be mutually exclusive. Indeed, considering the importance of Tgfβ
signaling in GI tract development and the multiple levels through which
Tgfβ is known to be regulated, such multifaceted control is very
likely.
MSCs are most probably present in all adult and fetal organs and share the
ability to differentiate into several mesenchymal cell types ex vivo
(McTaggert and Atkinson,
2007). However, their exact role in development is still unclear.
Fgfs are known to modulate the behavior of MSCs, serving to increase
proliferation and suppress differentiation of bone marrow-derived MSCs in
vitro (Tsutsumi et al., 2001;
Farré et al., 2007). We
propose that iMSCs play a role in development by regulating the
differentiation of adjacent mesenchymal cells (fibroblasts) through expression
of secreted Tgfβ inhibitors. The location of the iMSCs near the base of
villi may be important for this function as the range of signaling for
Tgfβ is very limited (Reilly and
Melton, 1996).
This potential function of MSCs is in stark contrast to the traditional
role of stem cells, which is to divide only occasionally in order to produce
daughter cells that, through repeated divisions via transit amplification,
produce committed lineages of mature differentiated cells. However, growing
evidence suggests that MSCs have the ability to function outside of this
paradigm. For example, it has been recently shown that MSCs have the potential
to play a role in disease by functioning as a nidus of paracrine signaling
(Gnecchi et al., 2006;
Karnoub et al., 2007). Here,
we show that this idea of MSCs as paracrine signaling centers can be extended
to developmental biology, and that these cells play an important role in gut
development by functioning as cellular transceivers. In this role, iMSCs
preferentially receive an active Fgf9 signal, and in turn communicate to the
surrounding mesenchyme by secreting Tgfβ superfamily inhibitors, thereby
preventing premature mesenchymal differentiation. In this manner, they create
a permissive environment through which mesenchymal expansion occurs, which we
propose contributes to the lengthening of the intestine.
It has been reported that the mesenchymal response to Fgf in other organ
systems, particularly the trachea, occurs in a scattered population similar to
what we observe in this study (Que et al.,
2007). Based on our observations in the intestine, we would
speculate that these scattered mesenchymal cells represent mesenchymal stem
cells residing in these organs. Thus, the concept of Fgf preferentially
signaling to MSC populations might extend to other organ systems.
Understanding the regulation of the signaling function of MSCs in response to
Fgfs may be informative in our understanding of their normal physiological
roles in development and disease.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/135/17/2959/DC1
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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