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First published online 31 October 2007
doi: 10.1242/dev.003798
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Max-Planck Institute of Immunobiology, Department of Developmental Biology, Freiburg i. Br., Germany.
Author for correspondence (e-mail:
hiiragi{at}mpi-muenster.mpg.de)
Accepted 31 August 2007
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Embryonic patterning, Lineage specification, Mouse pre-implantation embryos
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The mechanisms of morphogenesis and patterning of the blastocyst have been
addressed in many studies during the last decades
(Rossant, 2004
;
Rossant and Tam, 2004), but
conclusive models for these processes are still lacking. Three models
currently exist. The pre-patterning model suggests that blastocyst patterning
and morphogenesis can already be anticipated at the egg stage
(Zernicka-Goetz, 2002;
Gardner and Davies, 2003;
Gardner, 2007). A certain
cleavage pattern may establish differences in the developmental potential of
4-cell stage blastomeres, possibly through an epigenetic mechanism
(Piotrowska-Nitsche et al.,
2005; Piotrowska-Nitsche and
Zernicka-Goetz, 2005;
Torres-Padilla et al., 2007).
However, no reproducible molecular organization of the egg with relevance to
lineage segregation in the blastocyst has been identified thus far. The
regulative model suggests that differences between blastomeres with relevance
for lineage divergence appear after compaction (8-cell), consistent with the
highly regulative nature of the early mouse embryo
(Johnson and McConnell, 2004).
It has been suggested that the TE and ICM are specified on the basis of the
blastomeres' positions [inside-outside hypothesis
(Tarkowski and Wroblewska,
1967)], so that those on the outside will form TE, and those on
the inside ICM. Epithelial maturation may be involved in the specification
process (Johnson and Ziomek,
1981; Plusa et al.,
2005; Vinot et al.,
2005). Epithelialization and/or microtubule reorganization at
compaction results in an apical-basal organization of the cells along the
radial axis, and cleavage perpendicular to this axis may give rise to two
distinct daughter cells: a polar cell on the outside and an apolar cell on the
inside (Johnson and Ziomek,
1981; Houliston et al.,
1989; Yamanaka et al.,
2006). Evidence suggesting that morphogenesis of the blastocyst is
not pre-determined (Alarcon and Marikawa,
2003; Chroscicka et al.,
2004; Louvet-Vallee et al.,
2005; Motosugi et al.,
2005; Kurotaki et al.,
2007) supports the regulative model. Eccentric positioning of the
blastocyst cavity, driven by mechanical constraints during cavity expansion,
specifies its embryonic-abembryonic axis
(Alarcon and Marikawa, 2003;
Motosugi et al., 2005;
Kurotaki et al., 2007). The
third, so-called cryptic pre-formation model suggests that cleavage patterns
influence the future allocation of cells within the blastocyst and thereby
their fate (Graham, 1971).
Although both blastomeres at the 2-cell stage contribute to the inside and
outside population at morula and blastocyst stage, a preference of the
descendants of the earlier dividing cell to locate on the inside has been
observed (Surani and Barton,
1984). This model forms a bridge between the other two models,
since it appreciates the highly regulative nature of the mouse embryo while
also accommodating the influence of the history of a cell on its fate.
The TE versus ICM population has been proposed to be regulated by the
POU-domain transcription factor Oct4 (also known as Pou5f1 - Mouse Genome
Informatics) (Okamoto et al.,
1990; Rosner et al.,
1990; Scholer et al.,
1990) and the caudal-like transcription factor Cdx2
(Beck et al., 1995;
Strumpf et al., 2005), with
slight differences between inside and outside cells in Oct4 and Cdx2 protein
levels that are amplified through reciprocal inhibition to give a mutually
exclusive pattern of Cdx2 on the outside and Oct4 in the inside
(Niwa et al., 2005). Although
Cdx2 was thought to be the key molecule for the TE lineage, this protein
appears to be dispensable for TE specification, since
Cdx2-/- embryos form an expanded blastocyst including the
TE but fail to maintain TE function
(Strumpf et al., 2005;
Tolkunova et al., 2006) (A.
Tomilin, personal communication). Epiblast (EPI) and primitive endoderm (PE)
are marked by the homeodomain protein Nanog
(Chambers et al., 2003;
Mitsui et al., 2003;
Strumpf et al., 2005;
Chazaud et al., 2006) and the
transcription factor Gata6, respectively
(Chazaud et al., 2006).
Establishment of the reciprocal pattern within the ICM is mediated through the
Grb2-Mapk signaling pathway (Chazaud et
al., 2006). How initial differences among cells of the ICM are
established is not known.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Control embryos developed in vivo were isolated at 70 (n=9 for Cdx2 and Nanog, n=5 for Oct4; about 10 hp-c), 90 (n=4 for Cdx2 and Nanog, n=3 for Oct4; about 30 hp-c) and 115 (n=6 for Cdx2 and Nanog; about 55 hp-c) hp-hCG.
Immunohistochemistry
Embryos were washed in PBS and fixed in 4% paraformaldehyde (PFA; Sigma,
P6148) in PBS for 10-15 minutes at room temperature. For single blastomere and
denuded embryo stainings 1-3% bovine serum albumin (BSA; Sigma, A9647) was
added to all solutions except PFA. After washing with 0.1% Tween 20 (Sigma,
P7949) in PBS (PBS-T), embryos were permeabilized in 0.25% Triton X-100
(Sigma, T8787) in PBS for 30 minutes at room temperature, washed in PBS-T,
blocked with 3% BSA in PBS-T (blocking solution), and incubated with the
following primary antibodies for 1 hour at room temperature in blocking
solution: mouse monoclonal anti-Oct4 (Santa Cruz Biotechnology, sc-5279, 1:100
dilution); rabbit polyclonal anti-Cdx2
(Beck et al., 1995) (kindly
provided by A. Tomilin, Max-Planck Institute for Immunobiology, Freiburg,
Germany; 1:100 dilution) or mouse monoclonal anti-Cdx2 (BioGenex, MU392-UC,
1:100-200 dilution); rabbit polyclonal anti-Nanog antibody (kindly provided by
S. Yamanaka, Institute for Frontier Medical Sciences, Kyoto University, Japan;
1:400 dilution). Nanog expression pattern was confirmed using goat anti-mouse
Nanog (R&D Systems, AF2729; 0.4 µg/ml) on embryos fixed at 80 hp-hCG
(data not shown). After washing in blocking solution, embryos were incubated
with the following secondary antibodies for 1 hour at room temperature: Alexa
Fluor 488 goat anti-rabbit (Molecular Probes, A11034, 1:100 dilution), Alexa
Fluor 546 goat anti-mouse (Molecular Probes, A11030, 1:100 dilution) or
Alexa-Fluor 488 donkey anti-goat (Molecular Probes, A11055, 1:100 dilution) in
blocking solution. In addition, 0.9 U/100 µl Alexa Fluor 633 phalloidin
(Molecular Probes, A22284) was added to the secondary antibody solution to
visualize cell membranes and 10 µM 4,6-diamidino-2-phenylindole dilactate
(DAPI; Molecular Probes, D3571) to stain DNA.
Microscopy and image analysis
Embryos were washed in blocking solution followed by PBS and placed in a
drop of PBS on a glass-bottom dish (WillCo Wells, GWSt-5040) covered with
mineral oil. Laser scanning microscopy was performed using a Zeiss LSM 510,
with optical sections obtained every 1 µm. Auto z-brightness correction was
used to adjust fluorescence along the z-axis. Unmodified images were
analyzed using IMARIS imaging software version 4.2-5.5 (Bitplane AG). Nuclei
were segmented in 3D reconstructions based on DAPI staining with the
isosurface or spot tool (5-10 µm, depending on nucleus size). In the case
of isolated blastomeres, nuclei were segmented using the spot tool and region
growing function. The number of cells staining positive for Cdx2, Nanog or
Oct4 was evaluated visually (0 and 5 hp-c) or by segmentation (IMARIS
software) based on the respective staining (10-55 hp-c). Only blastomeres with
more intense nuclear than cytoplasmic staining were considered positive.
Protein levels were analyzed from unmodified data sets as mean fluorescence
intensities within segments. DAPI ratios were used to minimize error caused by
staining and LSM. Data were normalized with respect to background levels,
defined as the average of the mean fluorescence intensities of 3-5 randomly
chosen cytoplasmic spots divided by the average of all mean DAPI fluorescence
intensities within defined segments. To minimize variation based on tissue
depths, only nuclei within the central 2/4 along the z-axis were
included in the fluorescence intensity analysis until 45 hp-c; at 55 hp-c, all
cells were included in the analysis to allow inclusion of the whole ICM. The
images shown in some of the figures were modified using a low-pass filter
(Kernel size 3x3x3; Zeiss LSM Image Examiner 4.0.0.91) and
contrast enhancement (Adobe Photoshop CS or CS2) or gauss filter (Kernel size
3; Axiovision Ver. 4.5) and brightness (Adobe Photoshop CS or CS2). QuickTime
movies were made using ImageReady (Adobe, San Jose, CA).
Blastomere dissociation
Blastomere dissociation was performed at the 8-cell stage before compaction
at 60-65 hp-hCG directly after oviduct flush. To separate blastomeres, the
zona pellucida was removed by incubating embryos for 2.5 minutes in pronase at
37°C, followed by washing in H-KSOM. Embryos were then transferred to
calcium- and magnesium-free (-CM) KSOM medium for about 10 minutes. Pipetting
embryos up and down within the -CM KSOM with a narrow glass pipette
dissociated blastomeres. After dissociation, blastomeres were kept in KSOM
until fixation at 70 (n=5), 78 (n=11) or 90 (n=5)
hp-hCG.
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Cell-cell adhesion was also inhibited by antibodies against E-cadherin. Embryos were flushed from oviducts at E1.5 (before 8-cell stage), washed in H-KSOM and then immediately cultured in KSOM with anti-E-cadherin antibody or without antibody for a control. Compaction was reversibly inhibited using mouse monoclonal antibodies ECCD-1 and DECMA-1 [kindly provided by M. Takeichi (Riken, Center for Developmental Biology, Kobe, Japan) and R. Kemler (Max-Planck Institute for Immunobiology, Freiburg, Germany), respectively], both directed against the extracellular portion of E-cadherin. Both antibodies inhibited compaction to a comparable degree (DECMA-1 at a concentration of 1-2 µg/ml; ECCD-1 at a concentration of 3 mg/ml), and DECMA-1 was used in all subsequent experiments. Time-lapse recordings showed that after experimental inhibition of compaction for about 10 hours (until 75 hp-hCG) with 1 µg/ml DECMA-1 the temporal pattern of development after washing in H-KSOM was comparable to that of untreated embryos after compaction (see Fig. S2A and Movie 3 in the supplementary material; n=39/40, control n=16/16). Full-term development was confirmed by transfer of embryos to oviducts after washing in H-KSOM at about 90 hp-hCG as described below. Patterning of embryos treated with 1.5 µg/ml DECMA-1 was assessed by antibody staining of embryos fixed at 80 hp-hCG (see Fig. S2B in the supplementary material; n=3, control n=2). Patterning of developmentally delayed blastocysts was examined after treatment with 1 and 1.5 µg/ml DECMA-1, subsequent washing in KSOM at 78 hp-hCG and fixation at 96 hp-hCG (n=7 embryos). The Nanog and Cdx2 patterns of these embryos were the same as in untreated embryos (data not shown).
Time-lapse recording of the embryos
Time-lapse recordings were started at 75 hp-hCG. Temperature was maintained
by a Tempcontrol 37-2 digital (Carl Zeiss, Oberkochen, Germany) at 37.5°C
in a plastic chamber incubator XL (Zeiss) and a heatable mounting frame M-H
(Zeiss), attached to a Zeiss Axiovert 200M with Narishige manipulators. Zeiss
AxioVision Ver. 4.5 software was used for the acquisition of the time-lapse
images. The halogen lamp was set below 2.4 V to minimize the embryos' exposure
to light. Embryos were recorded every 30 minutes for up to 72 hours. The
time-lapse movies were converted to QuickTime movies using ImageReady
(Adobe).
Embryo transfer
Embryos were transferred into the oviduct of E0.5 pseudopregnant NMRI
females mated to vasectomized NMRI males as described
(Nagy et al., 2003). Live born
mice were obtained from oviduct transfers of untreated embryos
(n=4/16 transferred embryos) and embryos treated with DECMA-1 at a
concentration of 1-1.5 µg/ml (n=6/49 transferred embryos).
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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), we determined the position of blastomeres within the
conceptus in consecutive optical sections using actin staining with phalloidin
to label membranes. `Inside' cells are defined here as those lacking any
contact with the outer surface of the embryo, whereas `outside' cells have
such contact (white arrowheads in Figs
3,
4 and
5 point to examples of inside
blastomeres; green arrowheads mark outside blastomeres of special interest;
see also Movies 1 and 2 in the supplementary material, including consecutive
optical sections of embryos). The first inside cells appeared at 10 hp-c
(15±2 total cells, n=18,
Fig. 2)
(Barlow et al., 1972). In 13 of
18 embryos at this stage, all blastomeres still had contact to the outside,
although some with only a small portion of their surface
(Fig. 3,
Fig. 5A, asterisks). The number
of inside cells increased continuously, and at 25 hp-c, nine cells on average
were inside (9±2 inside cells, 34±4 total cells, n=17;
Fig. 2).
Establishment of the blastocyst is characterized by formation and
positioning of a cavity. The first small cavities were detectable
stereomicroscopically at 15 hp-c (20±5 cells, n=13),
increasing in size continuously thereafter and reaching blastocyst stage III
by 30 hp-c [59±3 cells, n=11; for classification see Motosugi
et al. (Motosugi et al.,
2005)]. The late blastocyst is organized to accommodate the first
lineages, marked by the restricted expression of transcription factors such as
Oct4 (EPI and PE), Nanog (EPI) and Cdx2 (TE). Oct4 was present in all cells
until 35 hp-c (58±6 cells, n=12; n=6 for Oct4),
declining gradually in the TE, so that by 55 hp-c only the ICM showed intense
Oct4 staining (n=4; Fig.
2, Fig. 3A). After
compaction, embryos exhibit intense cytoplasmic Cdx2 staining, occasionally
making it difficult to distinguish between cytoplasmic and nuclear staining.
It is unclear whether the cytoplasmic staining represents specific Cdx2 signal
or background (Strumpf et al.,
2005). Only the cases in which nuclear staining was clearly
distinguishable from cytoplasmic staining were considered positive in this
study. Cdx2 was absent (n=9/13) or detectable only at low levels at 0
hp-c (n=4/13, Fig.
3B). Nuclear staining increased during the following hours, and,
by 10 hp-c, all cells of some embryos (n=7/19) were Cdx2 positive
(Cdx2+) and on average 12 of 16 cells showed clear nuclear Cdx2 staining
(n=19 embryos, Fig.
3B). By 25 hp-c only cells located on the outside were strongly
Cdx2+ (n=13 embryos, Fig.
3B). Nanog was present in all blastomeres until 35 hp-c
(n=3/6 embryos, and on average 98.5±2% of all non-mitotic
cells were Nanog+; Fig. 2,
Fig. 3C), whereas by 55 hp-c,
high levels of Nanog were restricted to a cluster of cells embedded in the
ICM, presumably the EPI (n=5 embryos;
Fig. 3C).
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).
Another study, however, reported localized expression of Cdx2 in the cytoplasm
of oocytes and zygotes, and asymmetric segregation of Cdx2 in the nucleus of
only half of the blastomeres generated during the subsequent cell divisions
(Deb et al., 2006). We cannot
reproduce these observations with two Cdx2 antibodies, one of which has been
used in all three groups (Strumpf et al.,
2005; Deb et al.,
2006), but instead find upregulation of Cdx2 in all 8-cell
blastomeres (see also isolated 1/8 blastomere study below). Indeed, the report
of Deb et al. has been recently retracted
(Kennedy, 2006;
Roberts et al., 2007). With
the appearance of inside cells, there is a preference for strongly Cdx2+ cells
to be located on the outside (Fig.
3B, Fig. 4A,
Fig. 5A)
(Niwa et al., 2005;
Strumpf et al., 2005).
Nevertheless, single blastomeres with high Cdx2 levels were found in the
inside until 20 hp-c (Fig. 3B
10 and 20 hp-c, Fig. 4A 15
hp-c, Fig. 5A 15 and 20 hp-c;
and see Fig. S1B in the supplementary material, 70hp-hCG, white arrowheads),
whereas some cells on the outside showed low-level Cdx2 staining
(Fig. 3B 10-20 hp-c,
Fig. 4A 15 and 20 hp-c,
Fig. 5A 15 and 20 hp-c and see
Fig. S1B in the supplementary material, 70hp-hCG, green arrowheads). This
`position-independent' variability was restricted to a period from 10 to 20
hp-c. By 25 hp-c, no inside cells with high Cdx2 were found (n=13
embryos), but single cells on the outside occasionally had low levels of Cdx2
(in n=6/13 embryos, Fig.
3B 25 hp-c). By 35 hp-c, all cells located on the outside were
strongly Cdx2+ and those on the inside were Cdx2-negative (Cdx2-; n=6
embryos, Fig. 3B,
Fig. 4A,
Fig. 5A). These findings reveal
high variability of Cdx2 levels in the nuclei of blastomeres after
compaction.
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;
Kurimoto et al., 2006). This
mosaicism has been shown to be independent of position within the ICM
(Chazaud et al., 2006). Because
we observed strong variability in Nanog expression even at 10 hp-c
(n=15 embryos), we tested for a possible relationship between Nanog
expression in each blastomere and its position in the morula, i.e. inside
versus outside. However, Nanog variability appeared to be independent of the
position of the individual blastomeres within the conceptus
(Fig. 3C 10-35 hp-c, green and
white arrowheads). Furthermore, when the mosaic is established, Nanog is not
restricted to the ICM. Until 35 hp-c (bIII), cells on the outside strongly
expressed Nanog (Fig. 3C 25 and
35 hp-c, Fig. 5A 25 and 35 hp-c
and see Fig. S1C in the supplementary material), and at 45 hp-c, a subset of
TE cells was Nanog+ (in n=10/13 embryos,
Fig. 5A). Only at 55 hp-c was a
high Nanog expression restricted to a cluster of cells within the ICM,
presumably the epiblast (Fig.
3C, Fig. 5A,
n=5 embryos).
Cdx2 levels initially show no correlation to Oct4
It has been proposed that reciprocal inhibition between Oct4 and Cdx2
segregates TE and ICM fate (Niwa et al.,
2005). Analysis of Oct4 and Cdx2 expression within the same
embryo, however (Fig. 4A; see
Movie 1 in the supplementary material) revealed strong Oct4 expression in
nuclei of all blastomeres (Fig.
4A, 15-45 hp-c), independent of high variability in Cdx2
expression (Fig. 4A, 15 and 20
hp-c). Moreover, comparison of normalized mean fluorescence intensities of
Oct4 and Cdx2 within individual nuclei at different times post-compaction
(Fig. 4B; see Materials and
methods) revealed slow nuclear Cdx2 accumulation above cytoplasmic levels
[(t)=15 hours] but no evidence of a reciprocal relationship between
Cdx2 and Oct4 levels at 15 hp-c (note the `irregular cloud' in the scatter
plot for this time; n=3). At 25 hp-c, a separation of Cdx2+ and
Cdx2-populations was detectable, but both populations retained high levels of
Oct4 (n=4). At 30 hp-c, Cdx2+ cells had slightly lower levels of Oct4
than did the Cdx2-population (n=4; n=5 at 35 hp-c). However,
most Cdx2+ cells had a strong Oct4 signal until 45 hp-c (n=2). By 55
hp-c, Oct4 expression was strongly reduced in Cdx2+ cells (n=4).
These data demonstrate that Cdx2 accumulates and is maintained despite high
levels of Oct4, and that Cdx2 variability is established independent of Oct4
levels. Oct4 is weakly suppressed in the TE 45 hours after the onset of Cdx2
expression.
Cdx2 and Nanog variabilities show no correlation
Because of the variability of Cdx2 and Nanog expression, we investigated
the possibility of a relationship between their expression patterns. Analysis
of Cdx2 and Nanog within the same embryo
(Fig. 5A; see Movie 2 in the
supplementary material) revealed some nuclei that exhibited reciprocal levels,
whereas others showed high levels of both proteins. Comparison of normalized
mean fluorescence intensities of Nanog and Cdx2 within nuclei at different
times post-compaction (Fig. 5B)
revealed increased nuclear Cdx2 above cytoplasmic levels by10 hp-c
(n=4), in contrast to the much slower increase in Cdx2 in the
Oct4/Cdx2 plot (Fig. 4B). The
difference in times until detectable Cdx2 had accumulated probably reflects
the different sensitivities of the Cdx2 antibodies used (rabbit polyclonal
antibody for the analyses in Fig.
3B 0, 10, 20, 25 and 45 hp-c, and
Fig. 4; and mouse monoclonal
antibody for those in Fig. 3B
15, 35 and 55 hp-c, and Fig. 5;
see Materials and methods). By 15 hp-c, the Cdx2/Nanog plot
(Fig. 5B) formed an `irregular
cloud' (n=6); by 25 hp-c, separation of Cdx2+ and Cdx2-populations
was observed, independent of high Nanog levels in some cells of both
populations (n=5, n=4 at 35 hp-c, n=3 at 45 hp-c,
n=5 at 55 hp-c). These findings indicate the mutual independence of
Cdx2 and Nanog expression variability, and show the co-existence of Cdx2 and
Nanog within the same nucleus until the late blastocyst
(Fig. 5B).
Patterning of in vivo developed embryos
To examine whether culturing embryos ex vivo affects the patterning process
we analyzed total cell numbers and the expression of Oct4, Cdx2 and Nanog in
freshly collected embryos at three timepoints
(Fig. 2 and see Fig. S1 in the
supplementary material). Seventy hp-hCG represents morula stage (n=9
embryos for Cdx2/Nanog, n=5 embryos for Oct4), 90 hp-hCG early
blastocyst (n=4 embryos for Cdx2/Nanog, n=3 embryos for
Oct4) and 115 hp-hCG late blastocyst (n=6 embryos for Cdx2/Nanog).
According to our synchronization scheme we compared the condition of 70 hp-hCG
to 10 hp-c, 90 hp-hCG to 30 hp-c and 115 hp-hCG to 55 hp-c. At 70 hp-hCG
embryos consisted of 16 cells (n=2 embryos), at 90 hp-hCG of
68±10.75 (n=7 embryos) and at 115 hp-hCG of 155±12.60
cells (n=6 embryos), suggesting that in vivo embryos develop slightly
faster than in culture (Fig.
2). At 70 hp-hCG Oct4 was expressed at high levels by all cells
(see Fig. S1A in the supplementary material), while Cdx2 and Nanog levels were
highly variable among blastomeres (see Fig. S1B,C in the supplementary
material). High levels of Cdx2 were found preferentially in outside cells, but
in a few cases inside cells had higher levels than outside cells
(n=2/9 embryos; see Fig. S1B in the supplementary material). Nanog
was independent of position in the morula. At 90 hp-hCG Oct4 was expressed in
both the TE and ICM (see Fig. S1A in the supplementary material). In some
embryos Oct4 was slightly weaker in the TE than in the ICM (n=2/3
embryos). Cdx2 was restricted to outside cells (see Fig. S1B in the
supplementary material, 90 hp-hCG). Nanog was highly variable, with some cells
expressing high levels and others very low levels. Highly Nanog+ cells were
present in both the ICM and TE (see Fig. S1C, 90 hp-hCG in the supplementary
material). At 115 hp-hCG Cdx2 was restricted to the TE (see Fig. S1B in the
supplementary material). Nanog was expressed at high levels by a population
located within the ICM, presumably the EPI. In some embryos a small number of
TE cells had detectable Nanog levels (n=5/6; see Fig. S1C in the
supplementary material, 115 hp-hCG).
These data show that in vitro culture, despite delaying late blastocyst development, does not alter the patterning process. Both in vivo and ex vivo developing embryos exhibit the same characteristic patterning features.
Asymmetric cell divisions at the 8-cell stage may underlie Cdx2 variability
The late morula comprises an inside cell population with low Cdx2 levels
and a strongly Cdx2+ outside TE that initiates cavitation. These morphogenetic
processes are preceded by cellular polarization at the 8-cell stage. Cellular
polarization is known to be dependent on two pathways: cell-cell adhesion via
E-cadherin (Kemler et al.,
1977; Yoshida and Takeichi,
1982; Larue et al.,
1994) and reorganization of the microtubule network
(Houliston et al., 1989;
Maro et al., 1991). Owing to
the increasing cell-cell adhesion, the time point of cellular polarization is
called compaction. To understand the emergence of variability in Cdx2 and
Nanog protein levels among blastomeres we tested a possible role of cellular
polarization. For this we dissociated 8-cell embryos before compaction
(Fig. 6). Each 1/8 blastomere
was then allowed to develop (Tarkowski and
Wroblewska, 1967; Graham and
Lehtonen, 1979; Johnson and
Ziomek, 1981) until 70 hp-hCG (n=5 embryos, 40
blastomeres; Fig. 6A), 78
hp-hCG (n=11 embryos, 87 blastomeres;
Fig. 6B) or 90 hp-hCG
(n=5 embryos, 40 blastomeres; Fig.
6C).
At 70 hp-hCG many blastomeres had divided once to give rise to a 2/16
doublet (n=36/40, n=5 embryos;
Fig. 6A). Fourteen of 36
doublets appeared to have equally sized sister blastomeres
(Fig. 6A1, empty rhombus),
while the other 22 were unequal in size
(Fig. 6A2, filled circle)
(Johnson and Ziomek, 1983).
After cytokinesis, cell-cell adhesion between the sister blastomeres quickly
increased, as if they were undergoing compaction
(Fig. 6B). At 78 hp-hCG 85/87
blastomeres had divided once; 25 of 85 appeared symmetric
(Fig. 6B1, empty rhombus), 12
were slightly asymmetric (Fig.
6B2, empty circle) and 48 2/16 doublets had formed characteristic
structures in which one cell partially or completely engulfed the sister
blastomere (Fig. 6B3-4, filled
circle) (Johnson and Ziomek,
1983). To compare quantitatively Cdx2 and Nanog levels of two
daughter blastomeres we plotted the pair representing the mean fluorescence
intensity of the bigger one on the y-axis and that of the smaller one
on the x-axis for 70 hp-hCG (Fig.
6C1), or the swallowing blastomere along the y-axis, and
that of the swallowed one on the x-axis for 78 hp-hCG
(Fig. 6C2; symmetric pairs were
randomly attributed to the x- or y-axis). The swallowing and
swallowed blastomeres may well correspond to `outside' and `inside' cells of
an intact embryo, respectively (Johnson
and Ziomek, 1983). Therefore we will refer to them as such from
now on. Each symbol represents one doublet, each color one embryo. Four
doublets were excluded from the analysis at 78 hp-hCG because blastomeres were
mitotic. The red dotted line marks equal fluorescence levels of sister
blastomeres. For Cdx2, the positions of the empty rhombi (symmetric doublets)
are variable with many lying close to the dotted (i.e. equal) line
(n=12/13 at 70 hp-hCG, Fig.
6C1; n=15/25 at 78 hp-hCG,
Fig. 6C2), suggesting that
after symmetric division, Cdx2 levels are similar in sister blastomeres. Empty
circles (slightly asymmetric doublets) show no obvious trend for higher Cdx2
levels in either one of the sister blastomeres (n=12). Most filled
circles (asymmetric doublets) are positioned away from the line toward higher
Cdx2 in the `outside' cell (n=12/22 at 70 hp-hCG; n=40/46 at
78 hp-hCG). These data suggest that following cleavage a substantial number of
sister blastomeres acquired different Cdx2 levels, with bigger cells having
higher, and smaller cells lower, Cdx2 levels
(Fig. 6A2,C1). At 78 hp-hCG the
`outside' blastomere had higher Cdx2 levels than the `inside' one
(Fig. 6B2-4,C2). Variability of
the Nanog levels among sister blastomeres was small and without correlation to
asymmetric cell division from 8- to 16-cell stage
(Fig. 6C1-2), or that of Cdx2
levels (Fig. 6A,B).
|
): type
I, 4/32 quadruplet with three `outside' cells surrounding one `inside' cell
(n=23/40, Fig. 6D1);
type II, 4/32 quadruplet with two `outside' cells surrounding two `inside'
cells (n=6/40, Fig.
6D2); type III, 4/32 quadruplet with four `outside' cells, which
all had strong Cdx2 signal (n=7/40,
Fig. 6D3). Four of 40
quadruplets were not unequivocally attributable to type I, II or III.
`Outside' cells had high Cdx2 levels, whereas the `inside' cells had weak
signals. It is important to note that the composition of the type of
quadruplets derived from different embryos was highly variable
(Fig. 6D4; see Discussion). Taken together, these data suggest that isolated blastomeres attempt to continue blastocyst morphogenesis and embryonic patterning in the same temporal and spatial manner as intact embryos. Intriguingly, the timing of cleavage, compaction, molecular patterning, inside/outside segregation, as well as cavitation was comparable to that observed for non-manipulated embryos (Figs 2, 3, 4 and 5). Thus, most, and of some embryos all, 1/8 blastomeres are able to recapitulate the segregation into a Cdx2-inside and Cdx2+ outside population that is able to initiate cavitation. Furthermore, these data demonstrate a correlation between asymmetric cell divisions at 8-cell stage and the generation of variability in Cdx2, but not in Nanog protein levels.
The role of cell-cell contacts and cell cycle progression in embryonic patterning
To test the role of cell-cell contacts on the patterning process we
re-separated 2/16 doublets immediately following cleavage so that the
blastomeres were cultured essentially in the absence of cell-to-cell contacts
until fixation at 78 hp-hCG (n=3 embryos,
Fig. 6E). The 1/16 blastomeres
were variable in size and 23/31 cells apparently had polar characteristics
(Fig. 6E, filled triangle),
with one of the poles having more intense actin staining (i.e. microvilli of
the apical membrane) and the nucleus at the opposite pole
(Fig. 6E, arrowhead and
asterisk, respectively). The remaining eight blastomeres appeared apolar
(Fig. 6E, cross). The
normalized mean fluorescence intensity of Cdx2 was significantly higher in
polar cells (11.6±3.6 arbitrary units (a.u.);
Fig. 6F, black line), compared
to apolar cells (6.9±2.5 a.u.; Student's t-test
P=0.002; Fig. 6F,
black dotted line). Nanog levels were similar in both populations
(5.0±1.7 a.u. for polar cells, black line, 4.5±1.2 a.u. for
apolar cells, black dotted line; Student's t-test P=0.437;
Fig. 6F). This suggests that
Cdx2 levels of polar and apolar cells remain different when blastomeres are
kept isolated until the 16-cell stage and that continuous cell-cell contact is
not necessary to maintain variability.
Compaction can also be reversibly inhibited by antibodies directed against
the extracellular domain of E-cadherin (see Fig. S2A in the supplementary
material; see Materials and methods for details)
(Johnson et al., 1979;
Damsky et al., 1983;
Richa et al., 1985;
Vestweber and Kemler, 1985).
Indeed, embryos treated with those antibodies for a certain period were able
to form hatching blastocysts in vitro (see Fig. S2A and Movie 3 in the
supplementary material; see Materials and methods) and to give rise to
live-born mice after transfer into a foster mother (n=6/49). The
number of blastomeres in embryos inhibited until 90 hp-hCG in 1-1.5 µg/ml
was about 16 (16±1, n=14; 49±4 for control,
n=2). This suggests that culturing embryos in the presence of
anti-E-cadherin antibodies delayed cell-cycle progression. Furthermore, levels
of Cdx2 and Nanog protein remained low when compaction was inhibited with 1.5
µg/µl DECMA-1 for 15 hours (until 80 hp-hCG, n=3; control
n=2; see Fig. S2B in the supplementary material), suggesting that
cell cycle progression and/or E-cadherin mediated cell-to-cell contacts are
necessary for upregulation of Cdx2 and Nanog expression.
|
DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The mode of Oct4 restriction to a subset of cells differs significantly
from that of Cdx2 and Nanog. Oct4 never exhibits the phase of variability that
characterizes Cdx2 and Nanog before becoming restricted to the ICM. Thus, Oct4
does not reflect the patterning phases seen through Cdx2 and Nanog. Since Oct4
is necessary to impart pluripotency in the early embryo
(Nichols et al., 1998), this
protein might be present in all blastomeres until the first lineages are
irreversibly determined at blastocyst stage
(Rossant, 1975;
Rossant, 2004). This notion
contrasts with the recent proposal that reciprocal inhibition between Oct4 and
Cdx2 enhances blastomere differences and determines TE
(Niwa et al., 2005). Oct4
protein is present in all blastomeres long after (more than 20 hours) Cdx2 is
restricted, and it is not until then that Oct4 expression is low in Cdx2+
cells. Thus, Oct4 levels do not affect Cdx2 accumulation and maintenance,
suggesting that even if reciprocal inhibition between Oct4 and Cdx2 functions
during in vivo patterning, additional regulatory mechanisms must be involved
in its regulation (Tolkunova et al.,
2006). Nevertheless, the proposed reciprocal inhibition pathway
could function in the downregulation of Oct4 in Cdx2+ TE after the actual
patterning process.
Cdx2 variability reflects formation of an inside population
Cdx2 and Nanog exhibit two distinct patterning phases that eventually lead
to their restricted expression. In the first phase, molecular differences
between blastomeres are generated, and in the second phase, a relationship
between molecular signature and position within the embryo is established.
|
Descendants of isolated 1/8 blastomeres undergo a compaction-like event, many differentiate into an inside and outside population, and all initiate cavitation. In these blastocyst-like structures Cdx2 signal is high in the outside and low in the inside population, whereas Nanog levels appeared random. Importantly, all 1/8 blastomeres of at least some embryos are able to undergo these processes. Thus, the descendants of 1/8 blastomeres self-organize into blastocyst-like structures.
Our data from isolated blastomeres further suggest that differences in Cdx2
levels among blastomeres at morula stage may be generated by asymmetric cell
divisions of the 8- to 16-cell stage transition. The mechanism controlling the
symmetric or asymmetric division is unclear. Interestingly, we found that the
number of asymmetric divisions in isolated blastomeres varies from embryo to
embryo. Similarly, the number of inside cells of intact 16-cell embryos is
highly variable. Asymmetric divisions might be regulated by extrinsic cues,
such that, for example, cleavage patterns during the 1- to 8-cell stages
influence cleavage orientation of the subsequent divisions. But it is unclear
whether this occurs in an organized way as previously suggested
(Piotrowska-Nitsche and Zernicka-Goetz,
2005; Zernicka-Goetz,
2005), since cleavage plane specification during the first cell
division is not predetermined (Hiiragi and
Solter, 2004) and mitotic spindle orientation of the second cell
division appears random (Louvet-Vallee et
al., 2005). Cleavage orientation of the third division may depend
simply on mechanical constraints imposed on the dividing cells, thus resulting
in a stochastic cleavage pattern that may explain the variability in the
number of asymmetric divisions among embryos and the presence of Cdx2-cells on
the outside of the early morula.
Stochastic establishment of Nanog variability
The mechanism leading to Nanog variability and a complete expression mosaic
is unclear. As may be the case for Cdx2, a cleavage pattern could be involved
in its generation. Within the ICM, the cleavage pattern has been proposed to
generate two cell populations, one cytokeratin filament-positive and the
other, negative (Chisholm and Houliston,
1987; Yamanaka et al.,
2006). Apolar cells that are generated during the 8- to 16-cell
transition, but not those generated in the 16- to 32-cell transition, lack
filaments. EPI versus PE segregation within the ICM has been shown to involve
enhancement of Gata6 (Koutsourakis et al.,
1999) and repression of Nanog signaling through the Grb2-Mapk
pathway (Chazaud et al., 2006).
These mechanisms may cooperate to establish the Nanog mosaic.
Another study proposed that Nanog expression is regulated by histone
arginine methylation (Torres-Padilla et
al., 2007). This study suggested that 4-cell-stage embryos that
follow a specific second cleavage pattern, exhibit differential H3R26
methylation among the blastomeres. Furthermore, they concluded that
blastomeres with higher H3R26 methylation levels predominantly contribute to
the ICM. Our data, however, show that molecular differences in Nanog levels
are established independently of blastomere position in the morula. It will,
therefore, be important to analyze H3R26me levels at the morula stage, to see
whether they correlate to position or to Nanog levels.
The functional role of Nanog is not fully understood, although it is
required to maintain pluripotency in the epiblast lineage
(Chambers et al., 2003;
Mitsui et al., 2003). Thus, it
was an unexpected finding that Nanog is expressed not only in the ICM of the
blastocyst, but also in some cells of the TE. Furthermore, Nanog levels
neither show a relationship with asymmetric cell divisions from the 8- to
16-cell stage, nor with Cdx2 levels. Although it is possible that an
epigenetic mechanism as described above or as yet unknown factors direct
lineage segregation in an ordered fashion, the position-independence, as well
as the initial independence from Cdx2 levels and from a first round of
asymmetric cell division favor stochastic generation of variability in Nanog
levels (Novick and Weiner,
1957; Spudich and Koshland,
Jr, 1976; Ko,
1992; McAdams and Arkin,
1997; Sigal et al.,
2006).
Patterning and morphogenesis: sorting mechanisms
In a second patterning phase, a definite relationship with blastocyst
morphology is established. This could be achieved in two ways. First, the
emerging molecular signature defines the properties of the cells and enables
them to change position (Smith,
2005) with respect to internal and external cues. Cells would then
find their final position based on characteristics that might involve, for
example, specific cellular adhesion properties
(Kimber et al., 1982). Such a
sorting mechanism circumvents the necessity for readjusting molecular
signatures. Subtle changes in cell-to-cell contacts could relocate a
blastomere to an inside or an outside position. Movement of outside
blastomeres to the inside has been reported
(Fleming, 1987). At 10 hp-c
(about 16 cells) we detected the first cell that is truly buried within the
conceptus (Graham and Lehtonen,
1979) and several blastomeres with only a small portion of their
membrane in contact with the outside. The latter cells could be either
completely engulfed internally or extend their outside contact area. A
previous study found that two to seven blastomeres (on average 5.22 cells)
were located on the inside of 16-cell stage embryos
(Fleming, 1987). The
discrepancy with respect to our study may be due to different experimental
approaches and definition used in identifying the positions of the cells. The
requirement for only small changes in cell-to-cell contacts to completely
change the position of a cell with respect to inside versus outside might
explain why such a movement has not been observed thus far
(Kelly, 1979). However, recent
analysis by nucleus tracing in 3D time-lapse recordings revealed a highly
dynamic behavior of blastomeres, especially associated with cell divisions,
within the developing conceptus (Kurotaki
et al., 2007). We speculate that such a sorting through positional
change may depend on global differences in the molecular signature of a cell,
including adhesion properties, which could be reflected by Cdx2 levels. During
the process of blastomere engulfment in the morula interior, cells with high
Cdx2 would then tend to remain on the outside, whereas those with lower Cdx2
would move to the inside.
Secondly, cell position may dictate its fate
(Tarkowski and Wroblewska,
1967). In that case, readjustment of molecular characteristics
with respect to positional influence would be required. Alternatively,
expression of Cdx2 changes over time and is stabilized in cells in the correct
position.
In the case of EPI versus PE divergence, it seems that positional change
establishes the relationship between position and molecular signature. Based
on lineage tracing analysis, Chazaud et al.
(Chazaud et al., 2006) have
suggested that cells of the E3.5 ICM are determined as EPI or PE, independent
of position. The same study provided support for the sorting hypothesis from
time-lapse observations, which revealed dynamic behavior of cells within the
early embryo (Chazaud et al.,
2006). Another study consistently identified the presence of two
populations within the E3.5 ICM that are distinct on a global molecular level
(Kurimoto et al., 2006). These
data imply that these cells adopt the correct position after acquiring
molecular differences in the case of EPI and PE sorting. Currently, the change
in position, the change in gene expression, or both mechanisms in cooperation
remain plausible to explain the second patterning phase by sorting. Definite
proof awaits the availability of reporter lines that allow time-lapse-based
lineage tracing during the patterning process.
Autonomous emergence of asymmetries in the early mouse embryo
Based on molecular data on TE versus ICM and EPI versus PE segregation, as
well as experimental manipulation of pre-implantation development, we propose
a two-phase model of patterning of the blastocyst
(Fig. 7). In a first phase,
molecular differences between blastomeres are established. In the case of
Cdx2, variability may be generated through asymmetric cell division of polar
8-cell blastomeres. Since the number of asymmetric divisions varied from
embryo to embryo under manipulated conditions, it will be interesting to
understand in greater detail what controls asymmetric divisions at that stage.
The mechanism generating Nanog variability remains unclear. Interestingly,
anterior-posterior axis asymmetry of the post-implantation embryo may be
established stochastically at the blastocyst stage
(Takaoka et al., 2006).
Expression of the TGF-ß family member Lefty1
(Meno et al., 1996) was
detected in a few cells of the ICM at E3.5, but no correlation between
Lefty1-expressing cells and their position within the blastocyst was
identified. Definite pattern formation in relation to embryo morphology is
achieved in a second phase in which a sorting mechanism establishes a
relationship between the molecular signature of a blastomere and its position
within the conceptus. This model is consistent with the highly regulative
capacity of the early mouse embryo until late blastocyst stage
(Rossant and Lis, 1979;
Rossant and Vijh, 1980): the
fate of blastomeres is not determined irreversibly until blastocyst
morphogenesis is completed. Thus, we conclude that the first lineages may
emerge after the 8-cell stage, possibly due to stochastic processes and/or
asymmetric cell divisions. The morula-stage mouse embryo would be highly
heterogeneous with respect to the molecular profile, and regularities in
relation to morphology emerge later. From an evolutionary standpoint, these
features may be unique to the pre-implantation stage of mammalian species
(O'Farrell et al., 2004;
Motosugi et al., 2005).
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/134/23/4219/DC1
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
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