© 2000 American Heart Association, Inc.
Vascular Biology |
Fibroblast Growth Factor Plays a Critical Role in SM22
Expression During Xenopus Embryogenesis
From the Department of Cardiovascular Medicine, Graduate School of Medicine (T.O., I.S., K.M., S.K., Y.H., R.N., Y.Y., I.K.), the Laboratory of Molecular Embryology, Department of Biological Sciences, Graduate School of Science (K.S.), and the Department of Life Sciences (Biology), Graduate School of Arts and Sciences (M.A.), University of Tokyo, Tokyo, Japan.
Correspondence to Dr Issei Komuro, Department of Cardiovascular Medicine, University of Tokyo Graduate School of Medicine, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-8655, Japan. E-mail komuro-tky{at}umin.ac.jp
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Abstract |
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Abstract—Although smooth muscle cells (SMCs) are critical components of the circulatory system, the regulatory mechanisms of SMC differentiation remain largely unknown. In the present study, we examined the mechanism of SMC differentiation by using Xenopus laevis SM22
(XSM22) as an
SMC-specific marker. XSM22 cDNA contained a 600-bp open reading
frame, and the predicted amino acid sequences were highly conserved in
evolution. XSM22 transcripts were first detected in heart anlage,
head mesenchyme, and the dorsal side of the lateral plate mesoderm at
the tail-bud stage, possibly representing the precursors of
muscle lineage. At the tadpole stage, XSM22 transcripts were
restricted to the vascular and visceral SMCs. XSM22 was strongly
induced by basic fibroblast growth factor (FGF) in animal caps.
Although expressions of Xenopus cardiac actin were not
affected by the expression of a dominant-negative FGF receptor, its
injection dramatically suppressed the XSM22 expression. These
results suggest that XSM22 is a useful molecular marker for the SMC
lineage in Xenopus and that FGF signaling plays an
important role in the induction of XSM22 and in the differentiation
of SMCs.
Key Words: smooth muscle cells • SM22 • basic fibroblast growth factor • dominant-negative fibroblast growth factor receptor
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Introduction |
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
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Smooth muscle (SM) cells (SMCs) are the critical component of the circulatory, genitourinary, respiratory, and digestive systems. Unlike skeletal or cardiac muscle cells, which stably express cell type–specific genes after terminal differentiation and lose their ability to proliferate, SMCs exhibit remarkable phenotypic plasticity and retain the ability to reenter the cell cycle even after terminal differentiation. This unique property of SMC phenotypic modulation is associated with the change of expression patterns of SMC-specific markers and is implicated in the pathogenesis of a wide variety of cardiovascular diseases, including atherosclerosis, restenosis after angioplasty, and hypertension.1 It is important to identify the genetic pathways that govern SMC differentiation to understand the pathophysiology of cardiovascular diseases.
During embryogenesis, skeletal, cardiac, and SMCs are derived from distinct populations of myogenic precursor cells. Recent advances in developmental biology have revealed a number of regulatory determinants that control the differentiation of skeletal or cardiac muscle cell lineage. The MyoD family of skeletal muscle–specific transcription factors, including MyoD, myf-5, myogenin, and MRF-4, induces the differentiation of skeletal muscle cells in the somites.2 Cardiomyocyte precursor cells arise from anterior lateral mesoderm, and cardiac transcription factors, such as Csx/Nkx2.5, GATA-4, and myocyte-specific enhancer factor (MEF)2C, regulate the process of cardiomyocyte differentiation.3 4 However, the embryonic origins of SMCs are less clear, in part because they arise from different precursor populations in multiple regions of the embryo. Vascular SMCs are thought to originate from 2 origins, the mesenchymal neural crest5 6 and the lateral plate mesoderm,7 8 and visceral SMCs are thought to locally arise in several distinct regions.9
In addition to the lack of our knowledge regarding the ontogeny of the
SMCs, there is only a limited information on the regulatory mechanisms
of SMC-specific gene expression. Cell type–specific transcription
factors have been demonstrated to play critical roles in organ
development2 4 ; however, no SMC-specific transcription
factor has been isolated. The earliest known marker of differentiated
SMCs is SM -actin, which is induced concomitantly with the
recruitment of presumptive SMC precursors into the vessel
wall.1 Other differentiation-specific marker proteins,
such as SM22, calponin, and h-caldesmon, are sequentially induced
during vascular development. SM1, one of the SM myosin heavy chain
isoforms, is a marker of the differentiation/maturation of the SMC, and
another isoform, SM2, which is produced from the same gene as SM1 by
the alternative splicing mechanism, is a marker of maturation after
birth. Analysis of the promoter of SM -actin and SM1/2 genes
has suggested that serum response factor (SRF), which binds to the CArG
box, and the MEF2 family of MADS box transcription factors, which bind
to AT-rich elements, play critical roles in SMC-specific gene
expression.10 11 12 13 14 15 However, because SRF and MEF2 are not
SMC specific, the mechanism of SMC-specific gene regulation remains to
be elucidated.
Recently, SM22, a 22-kDa protein originally identified in the chick
gizzard, has been characterized as an SMC-specific molecular marker in
birds and mammals.7 16 17 18 19 20 21 22 SM22 is structurally related
to the actin- and tropomyosin-binding protein calponin23
and Drosophila mp20, which is expressed specifically in
synchronous oscillatory flight muscles.24 Because
SM22 is abundantly and predominantly expressed in vascular and
visceral SMCs,25 26 27 transcriptional regulation of SM22
has also been studied. Transgenic analysis of the SM22
promoter revealed that CArG boxes and SRF may play critical roles in
the transcriptional regulation of SM22 in arterial
SMCs.28 29 These results indicate that the SM22 gene
can be a useful molecular marker for the differentiation of SMCs,
although the precise function of its protein is unknown at
present.
In addition to tissue-specific transcription factors, secreted molecules are important as determinants of embryonic development, because the inductive signals between different germ layers or different tissues have been implicated in the differentiation of specific cell types. In fact, skeletal muscle differentiation is controlled by the mutually antagonistic molecules bone morphogenetic protein (BMP) and Noggin, which are secreted from the ectoderm and neural tube,30 and cardiomyocyte precursor cells are induced in the anterior lateral mesoderm by BMPs secreted from the endoderm.31 These results suggest that the differentiation of SMCs may also be regulated by secreted molecules. Although it has been reported that transforming growth factor-ß and platelet-derived growth factor play a critical role in the differentiation of neural crest cells into SMCs and in the maturation of mesenchymal cells into vascular SMCs, respectively,32 33 it is unclear whether these growth factors are also involved in the determination of SMC lineage during early embryonic development.
Xenopus laevis is an excellent experimental model system to
investigate the early embryonic development. One can easily examine the
roles of a molecule during embryogenesis by enhancing or inhibiting the
functions of its protein with the injection of wild-type or mutated
mRNA, and the effects of peptide growth factors on tissue
differentiation can also be easily examined by animal cap assays. In
the present study, to elucidate the regulatory mechanisms of SMC
differentiation, we used X laevis as an experimental system
and selected SM22 as a molecular marker of differentiated SMCs.
X laevis SM22 (XSM22) transcripts were detected in
presumptive SMC precursor cells within the lateral plate mesoderm at
the tail-bud stage. And at the almost same stage, the presumptive
skeletal and cardiac muscle precursor cells within the head mesenchyme
and the heart anlage expressed XSM22. Thereafter, expression of
XSM22 was restricted to SMCs at the tadpole stage. In animal cap
assays, XSM22 was strongly induced by basic fibroblast growth factor
(FGF), and the overexpression of a dominant-negative form of FGF
receptor resulted in the dramatic reduction of the XSM22 expression
not only in the abnormal trunk but also in the normal head. The
present study indicates that SM22 is a useful molecular marker
for the SMC lineage and suggests that FGF signaling may play an
important role in the induction of XSM22 and in the differentiation
of SMCs in Xenopus.
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Methods |
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Isolation of XSM22
cDNAPartial cDNA of XSM22
was isolated by reverse transcriptase
(RT)–polymerase chain reaction (PCR) with Xenopus stomach
RNA used as a template. A pair of degenerative primers (primer 1,
5'-CAG TCC AAG AT(C/T) GAG AAG AAG TA(C/T)-3'; primer 2, 5'-GAG GTC AAC
(A/G)GT CTG GAA CAT GTC-3') were synthesized on the basis of the
previously published amino acid sequences of mouse, rat, chick, and
human SM22.20 By use of the resultant 309-bp PCR
product as a probe, the Xenopus stage-49 whole-embryo
cDNA library was screened with standard protocols. Hybridization was
performed at 42°C for 16 hours in a solution containing 50%
formamide, 2x SSC, 10% dextran sulfate, 1% SDS, and 2.5x
Denhardt’s solution. Filters were washed to a stringency of 0.1x SSC
and 0.1% SDS at room temperature and exposed to x-ray film for 24
hours at -80°C. Positive recombinant phages were purified by
sequential screening at low plaque density. Phage DNA of isolated
positive clones was prepared by use of a Lambda Mini Kit (Qiagen), and
EcoRI-excised cDNA inserts were subcloned into pBluescript
II SK(-) plasmid vector (Stratagene). Sequencing of the isolated cDNA
was performed by the dideoxy chain-termination method with use of the
BigDye Terminator Cycle Sequencing Kit and ABI PRISM 310 Genetic
Analyzer (Applied Biosystems).
Northern Blot Analysis
RNA was extracted from adult Xenopus tissues by the
lithium/urea method.34 Total RNA (20 µg) was
size-fractionated on 1% agarose/formaldehyde gel and transferred to
Hybond N nylon membranes (Amersham). Blots were hybridized with an
-32P–labeled 309-bp fragment of XSM22 at
42°C for 16 hours in 50% formamide, 5x SSC, 1% SDS, and 5x
Denhardt’s solution, washed to a stringency of 0.1x SSC and 0.1% SDS
at 42°C, and exposed to x-ray film for 12 hours at -80°C.
RT-PCR
For RT-PCR analysis, RNA was extracted from
Xenopus embryos or animal caps with the use of RNAzol B
(Tel-Test Inc), and cDNA was synthesized by Superscript II RNaseH(-)
RT (GIBCO-BRL). PCR was performed by using the synthesized cDNA as a
template and a pair of primers (primer 1, 5'-TGA CGA GGA ACT AGA GCA
ACG-3'; primer 2, 5'-AAT CTT CAG CTG CCT TCA GG-3') that were expected
to amplify the 245-bp fragment. RT-PCR of Xenopus elongation
factor 1 (EF-1) was also performed as an internal control by
using a pair of primers (primer 3, 5'-CAG ATT GGT GCT GGA TAT GC-3';
primer 4, 5'-ACT GCC TTG ATG ACT CCT AG-3') that were expected to
amplify the 268-bp fragment. The cycling conditions were 3 minutes at
94°C for the initial denaturation step, followed by 35 cycles of 1
minute at 94°C (denaturation), 45 seconds at 55°C (annealing), and
30 seconds at 72°C (elongation); a DNA Thermal Cycler (Perkin-Elmer)
was used. The PCR products were electrophoresed on 2.0% agarose
gel, stained by ethidium bromide, and photographed on Fuji instant film
FP-3000B (Fujifilm).
In Situ Hybridization
Whole-mount in situ hybridization of Xenopus embryos
was performed essentially as described.35 Briefly,
600 IU of human chorionic gonadotropin (Denka) was injected into
pigmented Xenopus females to induce ovulation. Eggs were
stripped from the ovulating females and fertilized as previously
described,36 and the embryos were fixed at proper
stages in MEMFA buffer (0.1 mol/L MOPS, pH 7.4, 2 mmol/L EGTA,
1 mmol/L MgSO4, and 3.7% formaldehyde). To synthesize the
antisense and sense probes, the 621-bp cDNA fragment including the
XSM22 coding region was isolated and subcloned into the
NotI-SalI site of pBluescript II SK(-). This
construct was linearized by NotI (for the antisense probe)
or SalI (for the sense probe) and subjected to in vitro RNA
transcription by using T7 or T3 RNA polymerase, respectively, in the
presence of digoxigenin-modified UTP of the digoxigenin RNA labeling
kit (Boehringer-Mannheim). Hybridization was performed at
60°C overnight. After they were washed, the embryos were incubated
with anti-digoxigenin antibody (Boehringer-Mannheim) coupled to
alkaline phosphatase at 4°C overnight. The chromogenic
reaction was carried out with BM purple (Boehringer-Mannheim).
Pigmented embryos were fixed again in Bouin’s solution (1% picric
acid, 9.25% formaldehyde, and 5% glacial acetic acid) for 2 hours and
were bleached in 10%
H2O2 with 5% formamide.
For sectioning, these embryos were embedded in O.C.T. compound
(Tissue-Tek) and frozen. Sections (20-µm) were prepared by using
Cryostat Microtome (Leica) and photographed.
Animal Cap Assay
For animal cap assays, the vitelline membrane of
mid–blastula-stage embryos was manually removed with sharp forceps,
and one third of the top portion of the animal hemisphere was isolated
with a tungsten needle. Isolated animal caps were cultured in 1x
Steinberg’s solution (60 mmol/L NaCl, 0.67 mmol/L KCl, 0.34
mmol/L Ca(NO3)2, 0.83 mmol/L MgSO4,
10 mmol/L HEPES, pH 7.4) with 0.1% BSA containing growth factors for 2
hours. The following concentrations of growth factors were used: 5, 50,
and 200 ng/mL basic FGF (human recombinant, Becton Dickinson Labware);
1, 10, and 100 ng/mL activin (generous gift of Dr Yuzuru Eto, Ajinomoto
Co, Inc, Kawasaki, Kanagawa, Japan); and 5, 50, and 200 ng/mL
human recombinant BMPs (BMP cocktail, Sangi Co). After growth factor
treatment, caps were cultured in 0.1x Steinberg’s solution with 0.1%
BSA without growth factors at 23°C until control embryos reached
stage 35, when RNA was extracted from caps by use of RNAzol B.
Injection of Dominant-Negative Mutant of FGF Receptor
XFD/Xss and d50/Xss (generous gift of Dr Enrique Amaya)
containing a dominant-negative mutant of the Xenopus FGF
receptor (XFD) and a negative control of XFD,37
respectively, were linearized by EcoRI. cRNA was transcribed
by using SP6 RNA polymerase and capped with the use of an mCAP RNA
Capping Kit (Stratagene). Fertilized embryos were dejellied in 2.0%
L-cysteine (pH 8.0), transferred to 1x
Steinberg’s solution, and maintained at 14°C. At the 1-cell stage,
the embryos were transferred into 1x Steinberg’s solution containing
3% Ficoll (Nacalai Tesque, Inc) and penicillin-streptomycin
(GIBCO-BRL). Ten nanoliters of 0.1 ng/nL XFD cRNA, 0.1 ng/nL d50 cRNA,
or water was injected into animal hemispheres of the 1-cell embryos as
described previously.38 Injected and uninjected
embryos were collected at stage 39 individually, and RNA was extracted
by using RNAzol B. Furthermore, XFD-injected embryos were divided into
the head-side half, containing the head and heart, and the tail-side
half, containing the gut and the truncated tail, and RNA was extracted
by using RNAzol B. Then RT-PCR was performed by using a pair of primers
for XSM22 and for Xenopus cardiac actin (primer 5, 5'-TCC
CTG TAC GCT TCT GGT CGT A-3'; primer 6, 5'-TCT CAA AGT CCA AAG CCA CAT
A-3') that were expected to amplify the 250-bp fragment as a marker for
the skeletal and cardiac muscle lineages.39 40 These
PCR products were subjected to electrophoresis and quantified by
NIH image.
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Results |
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cDNA Cloning of XSM22
We isolated full-length XSM22
cDNA from the Xenopus
stage-49 whole-embryo cDNA library with a partial cDNA fragment
obtained by PCR as a probe. The nucleotide and amino acid
sequences of XSM22 are shown in Figure 1A
. The initiation methionine codon (ATG)
is in a favorable context for the Kozak41 consensus
sequence, and there is an in-frame stop codon (TGA) 15 bp upstream from
the translational initiation site. The XSM22 cDNA contained a 600-bp
open reading frame that encodes a 200–amino acid polypeptide. The stop
codon (TAA) was followed by a 3'-untranslated region that contained a
consensus polyadenylation signal (AATAAA) at 21 bp upstream from the 3'
terminal poly(A)+ tract. The amino acid sequences
of XSM22 were compared with those of mouse, rat, human, and chick
SM22 (Figure 1B). The XSM22 amino acid sequences were
80% identical to those of the mouse, rat, human, and chick SM22,
indicating that SM22 is highly conserved in evolution. XSM22 was
also 37% identical to Drosophila mp20.24
The mp20 protein has 2 EF hand motifs that are thought to be the
potential calcium-binding sites (Figure 1B, underlined), and 1
of them (site II) is conserved in XSM22 (Figure 1C).
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Tissue Distribution of XSM22 in the Adult
To examine the tissue distribution of the XSM22 gene
expression, Northern blot analysis was performed with 20 µg
of total RNA prepared from various adult tissues (Figure 2A). Among 8 adult Xenopus
tissues examined, XSM22 transcripts were detected at high levels in
aorta, intestine, kidney, and stomach and at lower levels in lung and
skeletal muscle. Other transcripts with larger sizes (3.0 kb) were
also detected in aorta, intestine, kidney, and stomach, which may
represent an alternatively spliced variant or unspliced mRNA.
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Temporal Expression Pattern of XSM22 During
Xenopus Embryogenesis
To examine the temporal expression pattern of XSM22 during
embryogenesis, RT-PCR was performed with use of the total RNA prepared
from whole embryos of each developmental stage (Figure 2B
). The
staging of embryos was determined according to Nieuwkoop and
Faber.42 RT-PCR analysis revealed that XSM22
transcripts were first detected at the tail-bud stage (stage 30), and
they continued to be expressed thereafter.
Spatial Expression Pattern of XSM22 During
Xenopus Embryogenesis
To examine the spatial expression pattern of XSM22 during
embryogenesis, whole-mount in situ hybridization analysis was
performed. By in situ hybridization analysis, XSM22
transcripts were first detected at stage 28 in the premyocardial tissue
of the heart anlage and in the dorsal side of the lateral plate
mesoderm lying at the ventral margin of the myotome (Figure 3A). At stage 38, in addition to the
expression in the heart and in the lateral plate mesoderm, XSM22 was
also detected in the head region (Figure 3B), and XSM22 was
predominantly expressed at the branches of the arterial
trunk at stage 49 (Figure 3C). In the sections of the head
region in the stage-39 embryo, XSM22 was detected symmetrically in
the head mesenchyme between the prosencephalon and the notochord,
possibly representing somitic precursors that differentiate
into head muscle later in development (Figure 4A). In the sections of body region at
stage 39, XSM22 was detected in the dorsal part of the lateral plate
mesoderm (Figure 4B and 4C). At the tadpole stage (stage 45),
XSM22 was detected in the thin layer surrounding the epithelium of
the gut (Figure 4D), representing the visceral SMCs
of the digestive system.
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Basic FGF–Induced XSM22 in the Animal Caps
To determine whether the growth factors known to have
mesoderm-inducing activities may also induce the expression of
XSM22, we performed animal cap assays. Although XSM22 was not
detected in untreated animal caps, it was induced in the caps treated
with basic FGF at concentrations of 50 and 200 ng/mL. XSM22 was also
induced in caps treated with activin (1, 10, and 100 ng/mL) or BMPs (5,
50, and 200 ng/mL) but at much lower levels than XSM22 induced by
bFGF (Figure 5), suggesting that FGF is
implicated in the induction of XSM22 gene.
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XFD Suppressed XSM22 Expression
To elucidate the role of FGF in the induction of the XSM22 gene
in vivo, we injected a cRNA of the dominant-negative FGF receptor,
XFD,37 into 1-cell embryos and examined the expression of
XSM22 by RT-PCR. The injection of XFD strongly decreased the
expression of XSM22 in whole embryos (n=5) compared with embryos
injected with d50 (n=3), a negative control of XFD, or water (n=3) and
compared with uninjected embryos (n=3, Figure 6A). The XFD-injected embryo shows major
deficiencies in trunk and posterior development but a normal head,
representing the importance of FGF signaling in the
formation of the posterolateral mesoderm.37 So we divided
these injected embryos into 2 parts, normal head (n=3) and abnormal
trunk (n=3), and performed RT-PCR. The expression of XSM22 in the
heads and trunks of XFD-injected embryos was significantly suppressed
compared with XSM22 expression in water-injected embryos. In
contrast, the expression of Xenopus cardiac actin in heads
(n=3) and trunks (n=3) was not affected by XFD injection
(P<0.05 and P<0.01, Figure 6B and 6C).
These results strongly suggest that FGF plays an important role in the
induction of the XSM22 gene and that the regulatory mechanisms of
XSM22 gene expression are possibly different from those of the
cardiac actin gene.
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Discussion |
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In the present study, we isolated XSM22
cDNA and
investigated the regulatory mechanisms of XSM22 expression. XSM22
was detected at the tail-bud stage in the head mesenchyme and
splanchnic layer of the lateral plate mesoderm,
consistent with the previous notion that precursors of
SMCs originate from these tissues. The expression of XSM22 became
thereafter restricted to the area where SMCs differentiate and
continued to be expressed in the adult tissues containing SMCs,
indicating that XSM22 can be a useful molecular marker of SMC
precursors and differentiated SMCs. We also found that FGF could induce
XSM22 gene expression in animal caps and that the FGF signaling is
necessary for XSM22 gene expression in the Xenopus
embryo.
SM22 was originally identified as a calponin-related 22-kDa protein
abundantly expressed in chick gizzard.16 43 44 The amino
acid sequences of the isolated Xenopus SM22 were >80%
identical to the amino acid sequences of SM22 from other species,
indicating that SM22 is highly conserved in evolution. In addition
to the observation that SM22 is related to calponin, part of the
SM22 amino acid sequences also show similarity to those of
Caenorhabditis elegans unc-87, a thin-filament–associated
protein, mutations of which affect the function of body wall
muscle.45 However, because SM22 is easily
extracted from the contractile apparatus, it does not
appear to be a structural component tightly bound to the cytoskeleton.
In addition, SM22 also shows relatively high homology with
Drosophila mp20, which contains 2 potential calcium-binding
sites,24 and 1 of these motifs is conserved in
SM22 of vertebrates. Therefore, although the precise
physiological functions of SM22 are unclear at
present, SM22 may be implicated in the control of SMC function
via the regulation of intracellular calcium handling.
Northern blot analysis revealed that XSM22 was abundantly
expressed in aorta, intestine, kidney, and stomach, which contain a
large amount of SMCs. Slight expression of XSM22 was also detected
in lung and skeletal muscle, which may reflect the XSM22 expression
in the vessels of these organs. These expression patterns of XSM22
in adult tissues were in general agreement with those of SM22 in
other species. Previous studies in rat indicated that SM22 was
uniformly expressed in all tissues that contain SMCs, whereas
expression of SM -actin was not detected in some tissues, such as
bladder, ovary, and lung.46 These observations suggest
that SM22 can be a useful molecular marker of differentiated SMCs
and may have several advantages compared with SM -actin as an
SMC-specific marker.
Whole-mount in situ hybridization analysis revealed that
XSM22 transcripts were first detected in the heart anlage and in the
lateral plate mesoderm at the mid tail-bud stage (stage 28) and were
also observed in the head region at the tadpole stage (stage 38). In
the sections of the stage-39 embryo, XSM22 transcripts were detected
symmetrically in the lateral plate mesoderm and in the head mesenchyme
around the notochord. Although the precise origin or the timing of
differentiation of SMCs is unclear at present, previous studies
using interspecies grafting experiments suggest that vascular SMCs are
derived from the splanchnic layer of the ventrolateral plate mesoderm
and from neural crest mesoectodermal cells.5 Therefore, it
is possible that the XSM22-positive cells observed in the lateral
plate mesoderm and in the head mesenchyme represent the
precursor cells of vascular SMCs. In situ hybridization
analyses in mice, however, have indicated that SM22 is not
detected in the region thought to be the origin of vascular SMCs.
Murine SM22 was detected in the primitive heart tube at embryonic
day 8.0, in the myotomal component of the somites at embryonic day 9.5,
and in the SMCs of developing dorsal aorta at embryonic day
9.5.20 Therefore, there may be species difference in the
expression patterns of the SM22 gene among vertebrates.
Recently, Xenopus eHAND (XeHAND), a basic helix-loop-helix
transcription factor implicated in the formation of the left ventricle
in mice,47 48 has been characterized as an early
marker for the vascular SMCs.49 XeHAND expression precedes
that of SM22, being first detected at stage 24 in the broad domain
of lateral mesoderm, and is condensed to the presumptive posterior
cardinal veins at stage 33/34. Interestingly, XeHAND is not expressed
in the head region and in other neural crest–derived tissues,
suggesting that XeHAND may regulate expression of the XSM22 gene in
lateral mesoderm–derived SMCs.
It also should be noted that the transient expression of SM22 was
detected in the developing heart but not in the adult heart.
Essentially the same expression pattern in the heart was observed with
SM22 in the mouse and with SM -actin in chick, mouse, and
Xenopus.7 Because HAND genes are expressed
in the heart at the early embryonic stage, it is possible that HAND
induces the expression of SM22 and SM -actin in the developing
heart. Further investigation is required to determine what
transcription factors regulate expression of these SM-specific
genes.
In our animal cap assays, XSM22 expression was strongly induced by
basic FGF. Previous studies showed that SM -actin was also induced
in animal caps by basic FGF treatment.7 In addition, the
expression of XSM22 was dramatically reduced in the XFD-injected
embryos, further supporting the notion that the signals provoked by FGF
are implicated in the process of SMC differentiation. Because we did
not examine the expression of XSM22 by whole-mount in situ
hybridization after XFD injection, it is not clear where the expression
of XSM22 is reduced. However, RT-PCR analysis revealed that
XFD injection strongly and specifically suppressed XSM22 expression
not only in the abnormal trunk but also in the normal head. In
addition, the expression of cardiac actin was not affected by
overexpression of XFD. These results suggest that the effects of XFD on
SM22 expression may not be due to the defects of mesoderm induction
in the posterolateral portion of the embryo38 and that FGF
is critically involved not only in posterolateral mesoderm induction
but also in the differentiation of SMCs, irrespective of cell origins.
On the other hand, the expression of SM -actin is induced by basic
FGF7 but is not suppressed by the XFD
injection.12 Furthermore, the expression of
Xenopus cardiac actin was not affected by XFD injection
(Figure 6B and 6C). These results may indicate that the
expression of SM22 was differently and specifically governed by the
FGF signaling pathway compared with the contractile proteins, such as
SM -actin and cardiac actin.
Several molecules, including the Src-family of tyrosine kinases
and the components of the mitogen-activated protein kinase
pathways, have been reported to mediate the mesoderm induction by
FGF.50 51 Several transcription factors, including SRF and
MEF2 family members, have been identified to mediate the SMC-specific
gene expression in vivo and in vitro.15 19 27 28 29 The
roles of these intracellular signaling molecules in the FGF-induced
differentiation of SMCs and the link between FGF signaling and these
transcription factors remain unclear. Recently, it has been reported
that the transcription factor MEF2C is required for the differentiation
of SMCs.52 The expression of SM22
promoter–driven LacZ is diminished in the arterial SMCs
but is maintained in the heart of MEF2C mutant mice. However, it
remains unclear whether MEF2C directly regulates SMC-specific genes,
because the vascular phenotype of the MEF2C mutant is similar
to that of vascular endothelial growth factor or flt-1
mutant mice.53 54 55 In vascular endothelial
growth factor or flt-1 mutant mice, endothelial cells
can differentiate, but the later aspects of vasculogenesis are
disrupted, suggesting the possibility that the defects in the SMC
differentiation in MEF2C mutant mice may be the secondary effects of
disorganized vasculogenesis.
Our present study is the initial step forward to elucidate the
regulatory mechanisms of SMC differentiation. Further detailed
analyses on the expression of XSM22 will pave the way to the
identification of the molecular mechanisms that control the process of
SMC differentiation.
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Acknowledgments |
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We thank Dr Hiroki Hikasa, University of Tokyo, for helpful comments on the manuscript and Dr Takashi Ariizumi, University of Tokyo, for technical advice on the manipulation of the embryo. We are grateful to Chika Masuo for technical assistance.
Received August 19, 1999; accepted November 2, 1999.
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