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First published online January 26, 2007
doi: 10.1242/10.1242/dev.02778
1 Natural Medicines Research Center, Korea Research Institute of Bioscience and
Biotechnology (KRIBB), Daejeon 305-333, Korea.
2 Department of Biological Sciences, Korea Advanced Institute of Science and
Technology, Daejeon 305-701, Korea.
3 Division of Molecular and Life Sciences, Pohang University of Science and
Technology, Pohang, Kyungbuk, Korea.
4 Laboratory of Cell Regulation and Carcinogenesis, National Cancer Institute,
Bethesda, MD 20892-5055, USA.
Authors for correspondence (e-mail:
hjlee{at}kribb.re.kr;
jkh{at}postech.ac.kr)
Accepted 6 December 2006
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: SRF, Germ-layer formation, Activin and Nodal signaling, Xenopus
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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;
Heasman, 2006). Several
members of the TGF-ß growth factor superfamily, including Activin, Vg1
and Nodal-related proteins are responsible for the induction of mesoderm and
endoderm germ layers as well as for the subsequent patterning of the embryo
(Schier, 2003). These ligands
bind to a serine-threonine kinase receptor complex leading, intracellularly,
to the phosphorylation and activation of the receptor-Smad family of signal
transducers (R-Smads) that includes Smad2 and Smad3. Upon activation, these
R-Smads translocate into the nucleus where they control gene expression in
association with Smad4 and certain transcription regulators
(Schier, 2003;
Shi and Massague, 2003;
Whitman, 2001). However, a
detailed understanding of how the cellular responses to TGF-ß ligands are
modified by the differential combination of distinct Smad partners remains
elusive.
Recent studies in the mouse and frog suggest that the intra- or
extracellular inhibition of mesoderm-inducing signals is crucial for
appropriate germ-layer specification. Inactivation of mouse Lefty2,
an extracellular feedback inhibitor of Nodal signaling, results in expansion
of the primitive streak and mesoderm migration defects
(Meno et al., 1999). Knockdown
of Xenopus Lefty also causes the fate domains of the organizer and
dorsal mesoderm to expand, leading to exo-gastrulation
(Branford and Yost, 2002;
Cha et al., 2006). In the frog,
the loss of function of intracellular factors such as Ectodermin and
Xema expands mesoderm at the expense of ectoderm specification
(Dupont et al., 2005;
Suri et al., 2005). The
phenotype of mice mutant for DRAP1, a transcriptional corepressor, resembles
that of Lefty2 mutants (Iratni et
al., 2002). These proteins have been shown to limit the spatial or
temporal extent of the response to Activin/Nodal signaling in vertebrate
embryos.
Serum response factor (SRF) is a MADS box-containing transcription factor
that binds to a serum response element (SRE) found in the promoters of a
variety of genes, including immediate early genes, neuronal genes and muscle
genes (Shore and Sharrocks,
1995). SRF contains a highly conserved N-terminal DNA-binding and
dimerization domain termed the MADS box - owing to its homology among yeast
(MCM1, Agamous), plant (Deficiens) and vertebrate (SRF) proteins - and a
C-terminal transactivation domain
(Johansen and Prywes, 1993;
Norman et al., 1988;
Shore and Sharrocks, 1995).
SRF controls cell growth and differentiation, neuronal transmission and muscle
development, and functions by regulating the expression of its target genes
(Carson et al., 2000;
Castillo et al., 1997;
Treisman, 1986). SRF-deficient
mouse embryos display early embryonic lethality owing to the absence of
mesodermal cells, and this has led to the proposal that SRF is required for
mesoderm formation during mouse gastrulation
(Arsenian et al., 1998).
Interestingly, experiments using SRF-/- embryonic stem cells
suggest that the phenotype of SRF mutants may be due to a non-cell-autonomous
defect in differentiation toward mesoderm, rather than any impairment in the
cellautonomous induction of the mesoderm program
(Weinhold et al., 2000).
To better understand the molecular mechanisms by which SRF affects germ-layer specification during vertebrate embryogenesis, we have investigated SRF function in Xenopus early embryos, where mesoderm induction and patterning are much better characterized than, for example, in mice. SRF expression is restricted to the prospective ectoderm in Xenopus early embryos. Ectopic expression of SRF inhibits mesoderm formation. Conversely, loss-of-function of SRF stimulates mesendoderm induction, thereby expanding the expression of mesendodermal genes toward the ectodermal territory. In addition, SRF, a binding partner of Smad2 and FAST-1, impedes their association in Activin/Nodal signaling. These results suggest that SRF may function to restrict inappropriate germ-layer specification throughout the vertebrate embryo.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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),
and developmental stages of the embryos were determined according to Nieuwkoop
and Faber (Nieuwkoop and Faber,
1994). Microinjection was carried out in 0.33xModified
Ringer (MR) containing 4% Ficoll-400 using a Nanoliter Injector (WPI).
Injected embryos were cultured in 0.33xMR until stage 8 and then
transferred to 0.1xMR for later culture to the appropriate stages.
Whole-mount in situ hybridization and RT-PCR
Whole-mount in situ hybridization was performed with digoxigenin
(DIG)-labeled probes as described
(Harland, 1991). For RT-PCR
analysis, total RNA was prepared from embryos or animal cap explants with TRI
reagent (Sigma) and treated with RNase-free DNaseI (Roche) to remove genomic
DNA. RNA was transcribed using M-MLV reverse transcriptase (Promega). PCR
amplification was performed using Taq polymerase (TaKaRa). Primers used for
RT-PCR analysis are described at the homepage of the De Robertis group
http://www.hhmi.ucla.edu/derobertis/index.html.
The number of PCR cycles for each primer pair was determined empirically to
maintain amplification in the linear range.
Plasmids, RNA synthesis, morpholinos and cell lines
The mammalian expression plasmid for Flag-tagged Smad2 was described
previously (Lee et al., 2004).
The pCGN-HA-SRF construct was kindly provided by Dr Jae-Hong Kim (Korea
University, Seoul, Republic of Korea)
(Johansen and Prywes, 1993).
GST-tagged SRF and deletion mutant constructs were obtained by PCR and cloning
into the BamHI and SpeI sites of eukaryotic expression
vector pEBG. Flag-tagged SRF was generated in the pEF-Flag vector. Myc-tagged
FAST-1 deletion mutant constructs were obtained by PCR using the Myc-FAST-1
construct as template and cloning into the EcoRI and XhoI
sites of pCS2-MT.
For expression in Xenopus embryos, XSRF constructs including
pSP64T-wt XSRF and pSP64T-dn XSRF (kind gifts from Dr Harumasa Okamoto,
Neuroscience Research Institute, AIST, Tsukuba, Japan) were linearized with
XbaI and their capped mRNAs were synthesized using the SP6 mMessage
mMachine kit (Ambion). The XSRF-6Myc construct was generated by subcloning the
sequence containing its coding region and 5' untranslated region with
MO-binding sites into the BamHI and ClaI sites of pCS2-MT.
FAST-VP16A and FAST-EnR constructs were described
previously (Watanabe and Whitman,
1999). The morpholino antisense oligonucleotides (GeneTools)
directed against Xenopus SRF were as follows: XSRF MO1,
CTGGTTACTGGGCAGCATCCCTTG; XSRF MO2, AAATTTA CTAATCTGCCCTTCCTTG. The standard
control MO (CO MO) was CCTCTTACCTCAGTTACAATTTATA.
Mv1Lu, HepG2 and HeLa cells were all maintained in Dulbecco's Modified Eagle Medium (high glucose) supplemented with 10% fetal bovine serum and 100 µg/ml gentamicin. Mv1Lu-SRF cells that stably express SRF were generated by transfection with pCGN-HA-SRF expression plasmid. A day after transfection, cells were split and selected for neomycin resistance. Neomycin-resistant colonies were pooled after 2 weeks of selection, expanded and analyzed.
Luciferase reporter assays
HepG2 or Mv1Lu cells were transiently transfected with different
combinations of plasmid DNA in 12-well plates using Lipofectamin and Plus
Reagent (Invitrogen, Rockville, MD) according to the manufacturer's
instructions. The cells were transfected with SRF, reporter plasmid,
ARELuc/FAST-1, constitutively active HA-tagged ALK4*, pCMV-Gal to
normalize transfection efficiency and pcDNA3 to normalize the amount of
transfected DNA. All transfections were normalized to a total of 1 µg of
DNA in each well. Cells were harvested 36 hours after transfection and the
luciferase activity was measured using Enhanced Luciferase Assay Kit (BD
Biosciences). Values were normalized to the ß-galactosidase activity and
represent the mean of three independent transfections with error bars
indicating the standard deviation. Similar results were obtained in three
separate experiments.
Immunoblotting and immunoprecipitation
293T, Mv1Lu and HeLa cells were used for the detection of protein-protein
interaction in vivo. For the treatment with Activin A, HeLa or Mv1Lu cells
were starved overnight in 0.2% serum-containing medium 24 hours after
transfection, and treated with 25 ng/ml Activin A for 2 hours. Cells were
lysed in ice-cold RIPA buffer containing 50 mM Tris-HCl pH 7.4, 150 mM NaCl,
1% NP-40, 1 mM EDTA, 5 mM Na3VO4, 50 mM NaF and Protease
Inhibitor Cocktail (Complete, Roche). Cell lysates were separated by SDSPAGE
and transferred to nitrocellulose membranes. The membranes were immunoblotted
with various antibodies, followed by incubation with HRP-conjugated antibodies
to rabbit or mouse IgG and detected by chemiluminescence according to the
manufacturer's instructions (Pierce).
For immunoprecipitation, cell lysates were incubated with the appropriate antibody for 2 hours, followed by incubation with Protein G Plus-agarose beads (Santa Cruz Biotechnology) for 1 hour at 4°C. The beads were washed four times with RIPA buffer and then boiled for 5 minutes in 2xSDS sample buffer. The eluted immunoprecipitates were analyzed by immunoblotting as described above. For GST pull-down assay, cell lysates were incubated with glutathione-agarose beads (Santa Cruz Biotechnology) for 2 hours. After washing the beads four times with RIPA buffer, immunoblotting was performed.
Antibodies used for immunoprecipitation: anti-SRF rabbit polyclonal antibody (G-20, Santa Cruz Biotechnology); anti-Flag (M2, Sigma); anti-Smad2 (Zymed). Antibodies used for immunoblotting: anti-Smad2 mouse monoclonal antibody (BD Transduction Laboratories); anti-HA (Y-11, Santa Cruz Biotechnology); anti-Smad4 (Santa Cruz Biotechnology).
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To examine the effects of ectopic expression of XSRF on axis formation, we
injected XSRF RNA into the marginal zone of two dorsal or ventral blastomeres
of four-cell stage embryos. Compared with the phenotype of uninjected control
embryos, dorsally XSRF-injected embryos exhibited a severely shortened body
axis and the truncation of anterior structures (72%, n=123)
(Fig. 1E,F). These phenotypes
are similar to those caused by experimental conditions in which Activin or
Nodal signaling is inhibited (Chang et al.,
1997; Osada and Wright,
1999; Piepenburg et al.,
2004). Ventral injection of XSRF, however, resulted in more modest
defects in trunk and tail development (Fig.
1G). To characterize at the molecular level the events leading to
these disrupted phenotypes, we next examined the expression of mesodermal
markers in XSRF-injected embryos. Ventral or dorsal injection of XSRF
suppressed the expression of ventral mesodermal marker Wnt8, pan-mesodermal
marker Xbra, and dorsal mesodermal marker Chordin, in the early gastrula
embryos (Fig. 1H-M). Moreover,
XSRF-injected embryos showed dramatically inhibited formation of the somites,
a paraxial mesoderm derivative, which was evident by the absence of MyoD
expression at the tadpole stages (Fig.
1N,O). Consistently, RT-PCR analysis showed that ectopic XSRF
could reduce the expression of mesodermal markers, such as Wnt8, Mix2 and
Xbra, in the ventral region (Fig.
1P). Taken together, these results indicate that ectopically
injected XSRF could interfere with mesoderm formation, thereby leading to the
defects in axis specification.
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Inhibition of XSRF function expands mesendoderm
To examine the effects of loss of XSRF function on germ-layer formation, we
first employed a dominant-negative (DN) XSRF construct that mainly comprises
the DNA-binding domain, lacking the C-terminal half of its wild-type form
(Belaguli et al., 1997;
Watanabe et al., 2005). Dorsal
injection of DN XSRF at the four-cell stage resulted in embryos with the
stout, shortened body axis and microcephaly
(Fig. 3A,B), which is similar
to the phenotypes caused by the increase in Activin/Nodal signaling. By
contrast, ventrally DN XSRF-injected embryos showed no dramatic changes in
phenotype (data not shown). Interestingly, overexpression of DN XSRF in the
animal region of four-cell stage embryos ectopically induced mesodermal
(Chordin and VegT), endodermal (Sox17ß) and neural (Zic3) markers in
ectoderm, with the reverse effect on the expression of epidermal keratin, an
epidermal marker (Fig. 3C). In
addition, DN XSRF enhanced the inducing activity of Activin protein in animal
cap cells, increasing the expression of two dorsal markers, Goosecoid and
Chordin, and decreasing that of Xbra which is dependent upon the relatively
low levels of Activin signals (Fig.
3D). Together, these data suggest that inhibition of XSRF function
may augment the mesendoderm-inducing signals in early embryos.
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SRF associates with Smad2 and FAST-1
To elucidate the molecular mechanism by which SRF suppresses Activin/Nodal
signaling, we first tested whether SRF could interact with Smad2, a crucial
mediator of these signaling pathways (Shi
and Massague, 2003). Indeed, coimmunoprecipitation experiments
using epitope-tagged proteins from 293T cells showed that SRF binds Smad2
(Fig. 5A). We also found that
endogenous Smad2 coimmunoprecipitates with endogenous SRF in Mv1Lu and HeLa
cells (Fig. 5B,C), showing that
their interaction occurs at physiological protein levels. The interaction
between Smad2 and SRF was enhanced by constitutively active Activin type-I
receptor (ALK4*) or Activin A. To locate the domain within Smad2
responsible for interaction with SRF, we evaluated the abilities of a series
of Smad2 deletion mutants to bind SRF using GST pull-down assay. These
indicated that it is the MH2 domain of Smad2 that retains the ability to bind
SRF (Fig. 5D,E).
Furthermore, we investigated whether SRF could also bind FAST-1, a
DNA-binding partner of Smad2 in Activin/Nodal signaling
(Whitman, 2001). GST pull-down
assays demonstrated that SRF interacts with FAST-1
(Fig. 6A,B). This association
was not affected by constitutively active Activin type-I receptor (data not
shown). To identify the region within FAST-1 required for this interaction, we
examined whether SRF could coimmunoprecipitate with a series of deletion
fragments of FAST-1 in GST pull-down experiments.
Fig. 6B shows that the Forkhead
DNA-binding domain is required for FAST-1 to associate with SRF.
|
;
Shi and Massague, 2003). Thus,
we checked whether SRF could suppress the phosphorylation of Smad2 by Activin.
Smad2, however, was phosphorylated by Activin in stable Mv1Lu cells
overexpressing SRF as strongly as in control Mv1Lu cells
(Fig. 7A), indicating that SRF
does not affect Activin-dependent phosphorylation of Smad2. Since SRF associates with the MH2 domain of Smad2 that mediates its interaction with Smad4, we also examined the possible inhibitory effects of SRF on the ligand-induced formation of the Smad2-Smad4 complex that is essential for TGF-ß or Activin signaling. However, formation of the Smad2-Smad4 complex induced by Activin was not reduced in Mv1Lu cells stably overexpressing SRF as compared with that in control Mv1Lu cells (Fig. 7B).
We also tested whether SRF could interfere with the ability of FAST-1 to bind Smad2, as both of them associate with SRF. As shown in Fig. 7C, Myc-tagged FAST-1 coimmunoprecipitated with Flag-tagged Smad2 and this interaction was enhanced by constitutively active Activin type-I receptor (ALK4*). However, the level of Myc-tagged FAST-1 that coimmunoprecipitated with Flag-tagged Smad2 was markedly reduced by the co-transfection of SRF (Fig. 7C), suggesting a negative role of SRF in the association of FAST-1 and Smad2. Since this indicates the possible inhibitory effects of SRF on FAST-mediated transcription, we speculated that an activated form of FAST-1 (FAST-VP16A) would recover the defective phenotypes caused by overexpression of XSRF in Xenopus early embryos. Accordingly, we found that disruption of axial structure by XSRF could be rescued by coexpression of FAST-VP16A (29% defective, n=52) (Fig. 7D,E). Conversely, co-injection of an inhibitory form of FAST-1 (FAST-EnR) could rescue the gastrulation-defective phenotypes caused by XSRF MO (45% defective, n=38) (Fig. 7F,G). Overall, these results suggest that SRF may negatively regulate Activin/Nodal signaling by inhibiting the formation of a functional complex between FAST-1 and Smad2.
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Onuma et al.,
2002; Osada and Wright,
1999). Consistently, these gain-of-function phenotypes of XSRF can
be rescued by coexpression of an activated mutant of the FAST-1 co-factor
(Fig. 7). Furthermore, SRF
inhibits Activin/Nodal signal-dependent transcription as analyzed by our
reporter assays and RT-PCR (Fig.
2). These demonstrate that SRF antagonizes Activin/Nodal signaling
upstream of, or in parallel to, the FAST-1 factor.
Conversely, XSRF loss-of-function results in a shift in the cellular fates
along the animal-vegetal axis, causing the mesoderm that normally exists at
the equator of the embryo to spread toward the animal pole at the expense of
ectoderm (Figs 3, 4). This
seems to be responsible for the defective embryos that show microcephaly and
alterations in gastrulation movement. Recent evidence has shown that loss of
inhibitors of mesoderm-inducing signals can lead to inappropriate germ-layer
development. Depletion of Xenopus Lefty, an extracellular inhibitor
of Nodal signaling, expands the organizer and mesendodermal tissues, with
consequent exo-gastrulation (Branford and
Yost, 2002; Cha et al.,
2006). The maternal protein Ectodermin acts as a ubiquitin ligase
for Smad4 to antagonize, intracellularly, TGF-ß signaling for ectoderm
specification (Dupont et al.,
2005). In addition, knockdown of maternal Zic2 transcription
factor causes the same phenotypes as excess Nodal signaling, as this protein
functions to negatively regulate Nodal-related gene expression during
anteroposterior patterning (Houston and
Wylie, 2005). This antagonism of TGF-ß/Nodal signaling for
proper germlayer formation is conserved in mice, and mutation of the
transcriptional co-repressor DRAP1 or mouse Lefty2 leads to expansion of the
primitive streak and severe gastrulation defects
(Iratni et al., 2002;
Meno et al., 1999). Given that
ectopic XSRF precludes mesoderm formation, both in vivo and caused by
Activin/Nodal signaling in animal caps, and that its knockdown leads to
expansion of mesoderm into the prospective ectoderm, it is reasonable to
suggest that XSRF has the same function of limiting the response to the
mesoderm-inducing signals during germ-layer specification.
|
).
However, it could be challenged by the fact that ectopic overexpression of
XSRF has an inhibitory effect on mesoderm formation as shown in our results. A
likely explanation for this paradox is that XSRF might function to establish
the vegetal-to-animal gradient of the mesoderm- and endoderm-inducing
signaling. In this regard, note that Activin and Nodal ligands induce target
genes in a concentration-dependent manner, with dorsal mesoderm and endoderm
at high concentrations and ventral mesoderm at low concentrations
(Agius et al., 2000;
Gurdon and Bourillot, 2001).
Although these mesoderm- and endoderm-inducing molecules are induced by VegT,
which is a transcription factor inherited by all vegetal cells and uniformly
distributed in the vegetal region, the cell fates, mesodermal or endodermal,
might be determined by either distinct transcription regulators or a gradient
of inhibitors of these signals along the animal-vegetal axis
(Heasman, 2006;
Zhang et al., 1998). The
existence of these inhibitors that counteract the activity of mesoderm- and
endoderm-inducers is suggested by the phenotype of VegT-depleted embryos, in
which the marginal zone cells differentiate as ectoderm at the expense of
mesoderm and the vegetal cells differentiate as mesoderm and ectoderm instead
of endoderm (Zhang et al.,
1998). Thus, it is possible that XSRF might function to set the
activity threshold of mesoderm-inducing signaling through the antagonizing
mechanism to enable the marginal zone to adopt a mesodermal cell fate but not
an endodermal one. Therefore, high expression of XSRF could suppress
mesodermal cell fates in the marginal zone instead, whereas its knockdown
could stimulate them in the animal pole region as demonstrated in our results.
Given this activity of XSRF in germ-layer specification, the absence of
mesoderm in the homozygous SRF-/- mice
(Arsenian et al., 1998), which
is in contrast to the expansion of mesoderm in SRF-depleted Xenopus
embryos, may be in part due to the over-induction of endoderm at the expense
of mesoderm. In support of this, strong staining for the ectodermal marker
Oct-6 and for the endodermal marker HNF-3ß was observed in SRF-mutant
mice (Arsenian et al., 1998).
Furthermore, the phenotypic difference between frog and mice mutants might
stem from the variable completeness of depletion in knockout and MO injection
methods. In addition, it has been shown that loss of the ability of
SRF-/- embryonic stem (ES) cells to differentiate into mesodermal
cell fates can be recovered by treatment with external factors or their
subcutaneous injection into nude mice
(Weinhold et al., 2000). This
suggests non-cell-autonomous impairment of SRF-/- embryonic cells
in mesoderm formation, the nature of which might be partly understood through
our proposal that SRF might function as a modulator of the levels of input
signals to determine cell fate.
|
|
).
Moreover, Ski could also repress Smads directly by recruiting the
transcriptional repressor N-CoR as well as the histone deacetylase complex
(HDAC) (Liu et al., 2001).
DRAP1 interacts with FAST-1, thereby preventing FAST-Smad2-Smad4 complex from
binding to its cognate DNA targets (Iratni
et al., 2002). In addition, inhibitory Smads (Smad6 and Smad7)
compete with R-Smads for binding to activated type-I receptors and thus
inhibit the phosphorylation of R-Smads
(Shi and Massague, 2003;
ten Dijke and Hill, 2004). Our
data show that SRF precludes the association of Smad2 and FAST-1 induced by
Activin signal (Fig. 7). This
suggests that SRF could function to impede Smad2-FAST complex-mediated
transcription in Activin/Nodal signaling. Supporting this, gain-of-function
phenotypes of SRF are similar to those of maternal FAST-depleted embryos
(Kofron et al., 2004) and can
be rescued by coexpression of an activated mutant of FAST-1
(FAST-VP16A) (Fig.
7). We cannot, however, exclude the possibility that SRF could
also affect FAST-independent transcription as FoxH1 depletion in animal caps
has no effect on the induction of Nodal target genes in response to Xnr1 or
Activin ligands (Kofron et al.,
2004), whereas overexpression of XSRF significantly inhibits the
same response (Fig. 2). Given
that Smad2 binds to SRF via its MH2 domain, which associates with the general
transcriptional co-activators p300 and CAATT-binding factor (CBF)
(Pouponnot et al., 1998;
Shi and Massague, 2003), SRF
may repress the Smad2-mediated transcription of various genes in a cell
context-dependent manner by preventing the interaction of the MH2 domain and
these co-activators. By contrast, a recent study shows that SRF associates
with Smad3 and activates TGF-ß1-dependent transcription during
myofibroblast differentiation (Qiu et al.,
2003). On the other hand, the general mechanism by which SRF
regulates gene transcription is known to involve cooperation with the ternary
complex factors (TCF), which are phosphorylated and activated by MAP kinase
cascades (Chai and Tarnawski,
2002). In Xenopus, SRF was shown, together with the
TCF-type Ets protein Elk-1, to regulate the transcription of Xegr-1,
an organizer-specific gene, downstream of the FGF-initiated MAP kinase pathway
(Panitz et al., 1998).
Interestingly, TGF-ß receptors can activate MAP kinase signaling pathways
(Derynck and Zhang, 2003).
These activated MAP kinase cascades inhibit or enhance Smad activity by
phosphorylating it, depending on the cell signaling context; but, in some
cases, they regulate Smad-independent transcription. It will be interesting to
examine whether the TCF- and the Smad-dependent SRF regulation of gene
transcription involve distinct signaling cascades or whether both of them
could be controlled via MAPK pathways by TGF-ß signaling. In addition, it
remains to be investigated in more detail how SRF could regulate gene
expression in a positive or negative fashion depending on its
transcription-factor binding-partners.
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|
ACKNOWLEDGMENTS |
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|
Footnotes |
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Present address: Brookdale Department of Molecular, Cell and Developmental
Biology, Mount Sinai School of Medicine, One Gustave L. Levy Place, New York,
NY 10029, USA
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