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First published online 15 February 2006
doi: 10.1242/dev.02277
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,
1 Samuel Lunenfeld Research Institute, Mount Sinai Hospital, 600 University
Avenue, Toronto, Ontario M5G 1X5, Canada.
2 Department of Medical Genetics and Microbiology, University of Toronto,
Toronto, Ontario M5S 1A8, Canada.
Author for correspondence (e-mail:
janet.rossant{at}sickkids.ca)
Accepted 5 January 2006
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SUMMARY |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Key words: Ets2, Trophoblast, Anteroposterior axis, Embryo patterning
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INTRODUCTION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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; Georgiades et al.,
2002; Rossant and Cross,
2001). At the blastocyst stage, when the trophoblast lineage is
first established, the trophoblast plays a key role in promoting implantation
in the uterus. Soon after implantation, but still well before the
establishment of the definitive placental stage, the PPT can be subdivided
into two main regions: the ectoplacental cone (EPC; the half of PPT that is
largely in contact with maternal tissues) and the extraembryonic ectoderm
(EXE; the half of PPT that contacts the epiblast). However, the role and
development of the PPT from implantation until the establishment of the
definitive placenta is largely unexplored.
During this period, the embryonic region undergoes the major events of
gastrulation and anteroposterior (AP) axis formation. The first morphological
sign of mouse AP patterning within the epiblast is the appearance of
ingressing mesoderm at its posterior end of the primitive streak (PS) at E6.5.
This signifies the onset of gastrulation, which, in turn, sets in motion many
of the subsequent embryonic patterning events. However, for the
postimplantation conceptus as a whole, the earliest known morphological and
gene expression AP asymmetries are evident within the visceral endoderm (VE),
which envelops the epiblast prior to and during epiblast AP patterning. At
E5.5, the distal VE (DVE) constitutes a localized VE thickening and displays a
unique expression profile (Rivera-Perez et
al., 2003). By E5.75, the DVE shifts anteriorly to reach the
anterior-most epiblast and becomes known as anterior VE (AVE)
(Thomas and Beddington, 1996).
It is well established that interactions between the epiblast and the VE are
necessary for AP axis formation. For example, Nodal expression within
the epiblast is required for mesoderm initiation, DVE formation and anterior
DVE shift, and Cripto is necessary in the epiblast for the anterior
shift of the DVE (Ding et al.,
1998; Lu and Robertson,
2004; Yamamoto et al.,
2004). Signaling from the DVE/AVE has been proposed to induce
anterior and suppress posterior identity in the epiblast, based on gene
knockout, tissue ablation and transplantation experiments
(Dufort et al., 1998;
Hallonet et al., 2002;
Kimura et al., 2000;
Perea-Gomez et al., 1999;
Thomas and Beddington, 1996).
Moreover, the anterior AVE shift per se is necessary for epiblast AP axis
formation, as its failure results in posterior epiblast characters being
localized proximally and anterior ones distally
(Ding et al., 1998;
Perea-Gomez et al., 2001).
The involvement of the PPT as a possible source of signals for these events
was suggested by several recent findings
(Beck et al., 2002;
Donnison et al., 2005;
Fujiwara et al., 2002;
Rodriguez et al., 2005). For
example, the Spc1 and Spc4 genes encoding secreted proteases
of the subtilisin-like proprotein convertase (SPC) family, were shown to be
required in EXE for the elongation of a pre-existing, mesoderm-producing PS
(Beck et al., 2002), although
it was unclear from this study whether EXE signaling is also required for
mesoderm and PS initiation (Beck et al.,
2002; Guzman-Ayala et al.,
2004). Another study reported that mutation of the
trophoblast-specific Ets-related factor Elf5 resulted in a loss of
EXE in all cases, and in varying defects in mesoderm formation and AP
patterning (Donnison et al.,
2005). Rodriguez and colleagues demonstrated that removal of the
EXE at pre-gastrulation stages resulted in failure to express markers of the
primitive streak and also in expansion of the AVE
(Rodriguez et al., 2005).
To elucidate further the role of trophoblast signaling in embryonic
patterning and to better understand early PPT development, we examined the
development of mouse conceptuses homozygous for the previously reported,
targeted mutation of Ets2
(Yamamoto et al., 1998).
Ets2, which encodes a member of the Ets family of nuclear
transcription factors (Sharrocks,
2001), was shown to be expressed exclusively in PPT during the
period of embryonic patterning, and Ets2 mutant embryos were reported
to show reduced growth of the EPC and to lack the chorion, the derivative of
the EXE (Yamamoto et al.,
1998). In view of these PPT defects, the possibility that epiblast
and VE patterning processes could be affected by the loss of Ets2 function in
the trophoblast remained an unexplored possibility.
We report here that loss of Ets2 in the trophoblast can lead to the loss of EXE identity and a failure to produce the appropriate patterning signals from the EXE to the epiblast, thereby resulting in a failure to form a PS or to undergo normal AVE spatial orientation. Our results provide new genetic insights into the pathways of EXE maintenance and function.
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MATERIALS AND METHODS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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) were maintained by crossing to outbred ICR strain mice.
Homozygous Ets2db1/Ets2db1 conceptuses
were collected from intercrosses of
Ets2db1/Ets2+ heterozygotes.
Genotyping was performed on individually isolated embryos, on Richert's
membrane directly, or on isolated embryos following in situ hybridization.
This involved two separate PCR reactions with a primer set that produced
either the wild-type (220 bp) or mutant (200 bp) sequences, which were
resolved on a 1.5% agarose gel as previously described
(Yamamoto et al., 1998). The
conditions for both PCR reactions were: 94°C for 3 minutes; 35 cycles of
94°C for 30 seconds, 64°C for 30 seconds and 74°C for 4 seconds;
then 72°C for 5 minutes. For the detection of the wild-type DNA the
primers used were: 5'-CGTCCCTACTGGATGACAGCGG-3' and
5'-TGCTTTGGTCAAATAGGAGCCACTG-3'. For the detection of the mutant
DNA the primers used were: 5'-CGTCCCTACTGGATGACAGCGG-3' and
5'-AATGACAAGACGCTGGGCGG-3'.
Whole-mount in situ hybridization and histology
Single-color whole-mount in situ hybridization, with either one probe or
two probes simultaneously, was carried out essentially as previously described
(Lickert et al., 2002), with
the exception that the proteinase digestion step was replaced by a 15-minute
incubation at room temperature with RIPA reagent (150 mM NaCl, 1% Nonidet
P-40, 0.5% Na deoxycolate, 0.1% SDS, 1 mM EDTA, 50 mM Tris pH 8). Double-color
whole-mount in situ hybridization was carried out as above with an additional
color development stage, as described previously
(Yamamoto et al., 2004). Color
development time varied from 4 to 48 hours depending on the probe. The probes
used for the detection of Ets2, T, Nodal, Hex, cerberus
(Cer1), Otx2, Bmp4, Oct4, Mash2, Fgf4, Foxh1, Spc4, Cdx2, Cripto,
Wnt3, Sox1, Sox2, Hoxb1 and Errß (Esrrb - Mouse
Genome Informatics) expression have been described before
(Beck et al., 2002;
Ding et al., 1998;
Donnison et al., 2005;
Guzman-Ayala et al., 2004;
Liu et al., 1999;
Luo et al., 1997;
Yamamoto et al., 1998;
Yamamoto et al., 2001;
Yamamoto et al., 2004). For
histology after in situ hybridization, embryos were post-fixed in 4%
paraformaldehyde, dehydrated through an ethanol series and embedded in wax
before sectioning. For semithin (1 µm) sections, embryos were fixed in 4%
paraformaldehyde, processed and stained according to standard methodology
(Georgiades et al., 2001).
Generation of chimeric conceptuses
Wild-type 129S6/B6-F1 hybrid ES cells ubiquitously expressing GFP
(Vintersten et al., 2004) were
aggregated with tetraploid embryos (one tetraploid embryo per ES cell clump),
and were subsequently transferred to recipient ICR females and the resulting
chimeras collected at E7.75. The tetraploid embryos were produced from
two-cell stage embryos derived from intercrosses of
Ets2db1/Ets2+ heterozygotes, followed
by electrofusion, as previously described
(Nagy, 2002). Upon embryo
isolation, the Richert's membrane, which is exclusively derived from the
tetraploid embryo, was genotyped as described above.
Embryo culture and microsurgery
E5.5 or E6.0 conceptuses, derived from matings between mice homozygous for
a GFP transgene under the control of Oct4 regulatory
elements (generous gift of A. Nagy, Samuel Lunenfeld Research Institute,
Toronto, Canada), were isolated free from their Richert's membrane and
cultured intact or after trophoblast ablation for either 32-36 (for E5.5) or
24 (for E6.0) hours. E5.5 conceptuses were typically isolated between 11 am
and 1 pm on the fifth day (plug day=day 0) and were morphologically staged so
that only those conceptuses with a strictly distal VE thickening were chosen.
E6.0 conceptuses were isolated around midnight of the fifth day. They were
cultured in serum-free conditions in a pre-equilibrated medium containing 60%
(v/v) serum replacement for embryonic stem cells (GIBCO), and 40% (v/v) of
DMEM, 1 mM Sodium pyruvate, 4 mM l-glutamate and 100 mM
ß-mercaptoethanol. The conditions were 37°C, 5% CO2 in
atmospheric oxygen. For trophoblast ablation, conceptuses were immobilized and
a cut was made along the embryonic-extraembryonic junction using finely pulled
1-mm-thick glass rods (World Precision Instruments). Most trophoblast-ablated
conceptuses healed within 30 minutes and only those that healed were kept for
further study.
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RESULTS |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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Ets2db1/Ets2db1 conceptuses display two main types of phenotypes
We generated and genotyped
Ets2db1/Ets2db1 conceptuses using a
PCR-based approach, as previously described
(Yamamoto et al., 1998). In
accordance with previous observations, heterozygous mutants were
indistinguishable from their wild-type littermates. Gross examination of all
homozygous mutants from E6.75 onwards revealed small size, reduced
accumulation of maternal blood around the implantation site and resorption by
E9.5 (Yamamoto et al., 1998).
Although all mutant embryos were much smaller than wild types from E6.75, two
classes of mutants could be detected. The majority of
Ets2db1/Ets2db1 conceptuses,
henceforth referred to as type I mutants, had a small PPT region relative to
rest of the conceptus, whereas the PPT in type II mutants was in proportion to
the size of the rest of the conceptus (Fig.
1F-H; Table 1).
Based on histology at E6.75, type I, but not type II, mutants showed an
absence of mesoderm and their VE was abnormally thickened in the
anterior-distal tip region (Fig.
1F-H).
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Examination of 84 conceptuses at E5.5 derived from Ets2db1/Ets2+ intercrosses revealed that 12 (14.3%) had a small PPT. Genotyping confirmed that these E5.5 conceptuses were Ets2db1/Ets2dbl (Fig. 2K and data not shown). Double in situ hybridization with either Hex or Cerberus, and either Cdx2 or Bmp4 (n=7), revealed an absence of EXE markers prior to the DVE-to-AVE transition (Fig. 2L,M).
Expression of Nodal and Fgf4 in type-I Ets2 mutants
Our findings demonstrate that, in type I Ets2 mutant embryos, the
distal PPT fails to display any EXE identity, but does not adopt an EPC
phenotype. Recent experiments suggested that secreted signaling molecules from
the epiblast, Nodal and Fgf4, are involved in EXE maintenance, with Fgf4
inhibiting EPC identity and Nodal plus Fgf4 promoting EXE fate
(Guzman-Ayala et al., 2004).
Examination of Fgf4 expression at E6.75 revealed that in contrast to
the wild-type situation, type I mutants show no expression of this gene
(n=3; Fig. 2N),
perhaps due to the fact that these mutants lack a PS, as Fgf4 at this
stage normally marks the distal PS rather than the epiblast. However,
Fgf4 expression was detected in the epiblast of mutants
(n=3) at E5.5 (Fig.
2O). We next investigated Nodal expression and showed
that in type I (n=3), but not type II (n=2) mutants,
Nodal expression is reduced and fails to become posteriorized
(Fig. 2P). A similar
abnormality in the locality of the Nodal expression pattern was seen
in a subset of mutants lacking one of the Nodal enhancers that
contains binding sites for the transcription factor Foxh1
(Norris et al., 2002), or in a
subset of homozygous Foxh1 mutants
(Yamamoto et al., 2001). We
therefore examined the expression of the Foxh1 transcription factor,
which is known to be involved in Nodal signaling
(Norris et al., 2002;
Yamamoto et al., 2001), and
show that it is also not expressed in type I mutants (n=3;
Fig. 2Q). These results
indicate that, in the epiblast of type I mutants, the upregulation and
posteriorization of Nodal, but not the epiblast expression of
Fgf4, are dependent on Ets2.
Lack of PS and mesoderm initiation in type I, but not type II, Ets2 mutants
We examined the expression of Oct4 (n=3; an
undifferentiated epiblast marker), T (n=4), Hoxb1
(n=3) and Cripto (n=3) (a PS and mesoderm markers)
from E6.75 to E7.75. The type I epiblast remained undifferentiated and failed
to express any mesoderm markers (Fig.
3A-F). Interestingly, Wnt3, a gene previously shown to be
necessary in vivo for both PS and mesoderm initiation
(Liu et al., 1999), is not
expressed in type I mutants (n=3)
(Fig. 3G). Histologically, type
II mutants showed signs of PS and mesoderm formation
(Fig. 1H), and this was
confirmed molecularly by the expression of T (n=3) at E6.75
in these mutants (Fig. 3H), and
by double in situ hybridization using T- and Cdx2-specific
probes (n=3; Fig. 3I).
In addition, the type I epiblast remains undifferentiated, as judged by the
expression of Oct4 at E7.75 (Fig.
3B), and fails to express any anterior neural markers such as
Sox1 (n=3; Fig.
3J), Sox2 (n=3;
Fig. 2G) and Gbx2,
either at E7.75 (n=3) or E8.5 (n=3;
Fig. 3K,L).
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Having already shown that the DVE in E5.5 mutants forms normally in the absence of EXE (Fig. 2L,M), we used Hex and cerberus expression to investigate the subsequent anterior DVE shift in Ets2 homozygous mutants. We confirm by molecular markers (n=5 for Hex; n=3 for cerberus), that the AVE shift is incomplete (Fig. 4G,H). Furthermore, the expression of AVE markers is not well confined to the midline but shows a broad expansion of expression over the surface of the anterior epiblast (Fig. 4G,H). Using either Hex-Cdx2 (n=2) or Hex-T (n=2) double in situ hybridization, we demonstrate that this VE abnormality is present only in type I mutants (Fig. 4J,K). By E7.75, the majority of type I mutants examined (4/6) show a strictly distal Hex expression domain (Fig. 4I), whereas a minority (2/6) display the same abnormal localization of the AVE as is seen at E6.75 (data not shown). The type I AVE defect therefore does not reflect a developmental delay in AVE formation, and, by E7.75, the initially incomplete anterior displacement of AVE becomes distally localized in the majority of cases.
The effect of the abnormal AVE localization in type I mutants on AP patterning in the epiblast was studied by examining the localization of Otx2 expression. As expected from the abnormal localization of the AVE, Otx2 expression in E7.75 type I (n=3) mutants is confined to the distal epiblast; this is in contrast to their wild-type counterparts, where it is expressed anteriorly (Fig. 4M). The positioning of posterior epiblast markers cannot be assessed because the absence of PS, mesoderm and related posterior epiblast genes from type I mutants means that there are no posterior epiblast characteristics to be abnormally localized.
Ets2 function is solely required in trophoblast for embryo AP patterning
The type I patterning defects, the early lethality of both types of
Ets2 mutant, and the trophoblast-specific expression of Ets2
prior to and during patterning, in conjunction with the previously reported
rescue of this lethality in chimeric mice with wild-type extraembryonic
tissues and Ets2db1/Ets2db1 embryo
proper (Yamamoto et al.,
1998), suggest that Ets2 function in PPT is required for
all the patterning defects reported here. However, the possibility that the
type I phenotype could require defective Ets2 function in both the
epiblast and PPT cannot be excluded.
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4N
Ets2db1/Ets2db1). In this type of
chimera, the epiblast and all its derivatives are ES-cell derived and GFP
positive, whereas the VE and PPT are tetraploid-embryo derived and GFP
negative. Chimeras were PCR genotyped using the exclusively extraembryonic
parietal endoderm on Reichert's membrane. 2N ES+/+4N
Ets2db1/Ets2db1 conceptuses at E7.75
display all the hallmarks of natural mated-derived E7.75 type I mutants. They
do not express the PS and mesoderm marker T (n=2), their
Hex-positive AVE is either distal (2/3) or anterior-distal,
resembling that in E6.75 type I mutants (1/3; data not shown), and their
epiblast still expresses Oct4 (n=2;
Fig. 5A-D). Chimeras with a
type II phenotype were also detected (P.G., unpublished). These findings indicate directly that the lack of PS and mesoderm, and the abnormal localization of AVE in type I Ets2db1/Ets2db1 conceptuses, are solely due to the absence of a functional Ets2 in the PPT.
Cultured wild-type conceptuses lacking EXE from E5.5 phenocopy the type-I embryonic defects
To determine whether the loss of epiblast patterning in type I
Ets2db1/Ets2db1 mutants was due to the
absence of EXE character, we investigated whether wild-type conceptuses that
develop in the absence of EXE, phenocopy the type I patterning defects.
We developed a serum-free embryo culture system to culture E5.5 intact or
trophoblast-ablated conceptuses for 30-34 hours, by which time intact
conceptuses reached the early-to-mid PS stage. Only those E5.5 conceptuses
with a strictly distal VE thickening were chosen to ensure that the experiment
started prior to the beginning of the anterior shift of DVE
(Fig. 6A). This thickening was
shown previously to express Hex, a gene that marks the DVE/AVE prior
to and during its anterior shift
(Rivera-Perez et al., 2003).
Conceptuses transgenic for GFP under the control of the epiblast-specific gene
Oct4 (Fig. 6A) were
dissected into embryonic (Epi+VE, Fig.
6B) and abembryonic portions (PPT,
Fig. 6B) using glass needles.
To assess the accuracy of trophoblast-ablation, some conceptuses were examined
immediately after surgery for the expression of the EXE marker Cdx2.
Ninety-three percent (26/28) of trophoblast-ablated embryos showed no
Cdx2 expression (Fig.
6C). Conversely, very few dissected conceptuses showed carry-over
of Oct4-GFP expression in the extraembryonic fragments
(Fig. 6A). This demonstrates
the accuracy of the approach and ensures that any effects of the separation of
embryonic and abembryonic regions are not likely to be due to either EXE
remnants still attached to the epiblast or the removal of the proximal
epiblast during dissection.
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DISCUSSION |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES |
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), where the entire post-implantation trophoblast region
adopted an EPC fate when Nodal signaling from the epiblast was lost. The epiblast patterning defects seen in type I Ets2 mutants can be solely attributed to the defects in the development of the EXE, thus strongly supporting a direct role for the EXE in signaling to the epiblast for PS and mesoderm initiation. First, Ets2 expression is detectable specifically in the trophoblast before and during the early streak stages of development, when the mutant phenotype starts to become apparent in the epiblast. Second, our tetraploid-ES chimera work shows directly that absence of Ets2 in the trophoblast can lead to epiblast patterning defects. Third, early patterning defects only occur in type I, but not type II mutants, correlating directly with the absence or presence of EXE markers. Fourth, absence of EXE marker gene expression is evident from E5.5 in the prospective type I mutants, well before any of the AP patterning processes affected in type I mutants normally begin, excluding the possibility that the absence of EXE markers is a consequence rather than a cause of type I embryo patterning defects. Fifth, E5.5, but not E6.0, wild-type trophoblast-ablated conceptuses lacking EXE phenocopy the type I epiblast defects.
While this work was in progress, two recent papers have also provided
evidence that trophoblast signaling is required for inducing posterior
epiblast identity. All conceptuses with a targeted deletion of another
trophoblast-specific Ets-related gene, Elf5
(Donnison et al., 2005), were
reported to lack EXE from E5.5, although only about half of them were shown to
fail to form mesoderm as judged by T and Cripto expression.
This work therefore left open the question of whether EXE signaling is
required for mesoderm formation, because those Elf5 mutants with
mesoderm, were reported to also lack EXE. Our work, however, using double in
situ hybridization with mesoderm and EXE markers on both types of
Ets2 homozygous mutants, plus our culture of wild-type
trophoblast-ablated conceptuses lacking EXE, provides strong evidence that EXE
signaling is indeed required for mesoderm formation. Consistent with our
results, Rodriguez and colleagues
(Rodriguez et al., 2005) have
also demonstrated that removal of the EXE at pre-gastrulation stages followed
by culture in the presence of serum results in failure to express markers of
the PS, and that the ectopic transplantation of EXE cells could induce ectopic
T expression.
Rodriguez et al. also reported an expansion of the AVE in
trophoblast-ablated conceptuses (Rodriguez
et al., 2005). Our results with such cultured EXE-deleted embryos
do not show such a dramatic expansion of DVE/AVE markers. This difference may
be a result of the serum-free conditions we used, which may not promote full
expansion of the DVE/AVE. However, we do see defects in the localization of
the AVE in type I mutant Ets2 embryos in vivo, supporting for the
first time an in vivo role for the EXE in the proper localization of the
AVE.
All of these results strongly suggest that the EXE is a key source of signals to pattern the early embryo. Our work, taken together with the findings of others, provides evidence for a bidirectional signaling model in which Fgf4 and Nodal from the epiblast signal to maintain the EXE, which in turn produces direct signals to the epiblast, such as Bmp4, and indirect signals, such as the Spc proteases, which act on the epiblast to promote mesoderm development and posterior epiblast identity, probably largely by modulating Nodal activity.
Fgf4 is known to be required for maintenance of the trophoblast stem cells
of the EXE both in vivo (Goldin and
Papaioannou, 2003) and in vitro
(Tanaka et al., 1998). Recent
evidence from the Constam Laboratory suggests that the default state of
pre-gastrulation PPT is EPC, and that the EXE is maintained by Fgf4-mediated
inhibition of EPC formation in cooperation with active Nodal signaling
(Spc-mediated processed Nodal)
(Guzman-Ayala et al., 2004).
Our type I PPT phenotype is consistent with this model because Fgf4 expression
is still present in the pre-gastrulation epiblast of type I Ets2
mutants, suggesting that correct Fgf4 signaling from the epiblast can still
inhibit the distal PPT from adopting an EPC fate. The reduced Nodal expression
in type I mutants provides an explanation for the apparent absence of EXE
identity in the distal PPT of type I mutants: Fgf4 alone would be sufficient
to block EPC markers but not sufficient to induce EXE markers if Nodal
signaling is defective (Guzman-Ayala et
al., 2004). The barely detectable Nodal expression in
Ets2 mutants suggests defective Nodal signaling from the epiblast to
the trophoblast, as well as within the epiblast, in view of the known
autoinductive function of Nodal for amplifying its own expression
(Beck et al., 2002;
Guzman-Ayala et al., 2004;
Norris et al., 2002). The
reduced Nodal expression in type I mutants can be attributed to an
intrinsic requirement for Ets2 in PPT for the induction of the EXE signals
responsible for elevating Nodal expression in the epiblast. These
signals would include the secreted proprotein convertases Spc1/Spc4. Spc1/Spc4
in the EXE are required for the post-translational activation of the secreted
form of Nodal, which is necessary for elevating functional Nodal levels in the
epiblast and Bmp4 levels in the EXE (Beck
et al., 2002; Guzman-Ayala et
al., 2004), and their expression is lost in Ets2 mutants.
Because it is known that Nodal plays a key role in establishing AP patterning
in the embryo (Brennan et al.,
2001; Liu et al.,
1999; Lu and Robertson,
2004), most of the effects on epiblast patterning in Ets2
mutants can be explained by the reduced signaling of activated Nodal in the
epiblast. Given that Bmp4 is also known to induce certain posterior mesoderm
cell types, including extraembryonic mesoderm and germ cells
(Fujiwara et al., 2001;
Lawson et al., 1999), the
major defects in posterior development seen in Ets2 mutants are
probably caused by a combination of the loss of both Bmp4 and Nodal signaling.
A reduction in active Nodal in the epiblast would also lead to a loss of
signaling back to the trophoblast for the induction and/or maintenance of EXE
identity. Thus the initial cell intrinsic loss of EXE markers in the
trophoblast of Ets2 mutants would be compounded by a subsequent loss of the
Nodal signaling in the epiblast that would normally act to help maintain the
EXE region.
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;
Perea-Gomez et al., 2001)
(Fig. 4M).
One scenario to explain the DVE/AVE defect of type I mutants (localized
beneath the distal tip epiblast and the distal half of the anterior epiblast,
as opposed to beneath the proximal half of the anterior epiblast) could be
that the reduced levels of Nodal observed in these mutants as a
result of the lack of EXE [caused by the apparent absence of EXE-derived
Spc1/Spc4, previously shown to be required for the elevation, but not for the
initial induction of Nodal levels in the epiblast
(Beck et al., 2002;
Guzman-Ayala et al., 2004)]
could be sufficient for DVE formation and for some limited anterior DVE shift,
and that EXE-dependent Cripto expression could be responsible for the
completion of this DVE shift so that it becomes correctly localized. This
scenario is consistent with the following findings. (1) Although a complete
lack of Nodal signaling results in an absence of the DVE
(Brennan et al., 2001), reduced
Nodal signaling [as is the case for the majority of double mutants lacking
both functional alleles of Foxh1 and one allele of Nodal
(Yamamoto et al., 2001;
Yamamoto et al., 2004)] is
sufficient for DVE formation, albeit abnormally localized. (2) Low Nodal
levels could be sufficient for an initial anterior DVE shift, because during
normal development, between E5.5 and E5.7, when the DVE just begins to shift
anteriorly, Nodal levels are low
(Norris and Robertson, 1999;
Varlet et al., 1997;
Yamamoto et al., 2004). (3) An
absence of Cripto could be responsible, in a genetic
background-dependent fashion, for either no effect on AVE formation, a
complete failure of AVE formation, or a partial failure of AVE formation, as
in one genetic background Cripto nulls displayed normal AVE formation
(Xu et al., 1999), whereas in
a different, but mixed, genetic background the Cripto mutation
resulted in either a distally localized AVE, or a `slightly askew' or `broad
and asymmetric' DVE/AVE localization (Ding
et al., 1998). The proximal, as opposed to posteriorized,
expression pattern of Nodal in type I mutants is consistent with the
proposed role of Nodal in correctly localizing the DVE/AVE by having a
proliferative/anti-migratory effect on the VE cells close enough to
`experience' the Nodal signal (Yamamoto et
al., 2004) because it provides an explanation (not mutually
exclusive with the absence of Cripto) as to why the incomplete shift
of type-I DVE/AVE does not progress to reach the anterior
embryonic-extraembryonic junction.
The growth retardation observed in both types of Ets2 mutant
embryos could be attributed to the previously shown abnormal interactions
between trophoblast and maternal cells, as judged by the defective anatomy and
gene expression at the implantation site within both the EPC and the maternal
decidua. The apoptosis detected within the Ets2 mutants was also
suggested to contribute to this (Yamamoto
et al., 1998).
In conclusion, type I Ets2 mutant mice provide new insights about early trophoblast development and how the trophoblast influences fundamental embryonic AP patterning events.
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ACKNOWLEDGMENTS |
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Footnotes |
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Present address: Program in Developmental Biology, Hospital for Sick
Children, 555 University Avenue, Toronto, Ontario M5G 1X8, Canada
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REFERENCES |
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TOP
SUMMARY
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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