Analysis of cell lineage in two- and four-cell mouse embryos
Toshihiko Fujimori1,*,
,
Yoko Kurotaki1,
Jun-ichi Miyazaki2 and
Yo-ichi Nabeshima1
1 Department of Pathology and Tumor Biology, Graduate School of Medicine, Kyoto
University, Yoshida-Konoe Cho, Sakyo-ku, Kyoto 606-8501, Japan
2 Division of Stem Cell Regulation Research, Osaka University Medical School,
2-2 Yamadaoka, Suita, Osaka 565-0871, Japan
Author for correspondence (e-mail:
fujimori{at}lmls.med.kyoto-u.ac.jp)
Accepted 30 June 2003
 |
SUMMARY
|
---|
Compared with other animals, the embryos of mammals are considered to have
a highly regulative mode of development. However, recent studies have provided
a strong correlation between the first cleavage plane and the future axis of
the blastocyst, but it is still unclear how the early axes of the
preimplantation embryo reflect the future body axes that emerge after
implantation. We have carried out lineage tracing during mouse embryogenesis
using the Cre-loxP system, which allowed us to analyze cell fates over a long
period of development. We used a transgenic mouse strain, CAG-CAT-Z as a
reporter line. The descendants of the manipulated blastomere heritably express
ß-galactosidase. We examined the distribution of descendants of a single
blastomere in the 8.5-day embryo after labeling at the two-cell and four-cell
stages. The derivatives of one blastomere in the two-cell embryo randomly mix
with cells originating from the second blastomere in all cell layers examined.
Thus we find cells from different blastomeres intermingled and localized
randomly along the body axis. The results of labeling experiments performed in
the four-cell stage embryo fall into three categories. In the first, the
labeled cells were intermingled with non-labeled cells in a manner similar to
that seen after labeling at the two-cell stage. In the second, labeled cells
were distributed only in the extra-embryonic ectoderm layers. Finally in the
third category, labeled cells were seen only in the embryo proper and the
extra-embryonic mesoderm. Manipulated embryos analyzed at the blastocyst stage
showed localized distribution of the descendants of a single blastomere. These
results suggest that incoherent clonal growth and drastic cell mixing occurs
in the early mouse embryo after the blastocyst stage. The first cell
specification event, i.e., partitioning cell fate between the inner cell mass
and trophectoderm, can occur between the two-cell and four-cell stage, yet the
cell fate is not determined.
Key words: Mouse, Cell lineage, Cre recombinase
 |
Introduction
|
---|
It is well known that the future embryonic body axes are specified prior to
the first cleavage in a variety of animals. In the case of the mammalian
embryo, however, the timing of the earliest specification events that control
the future body axes are still unclear
(Beddington and Robertson,
1999
; Tam et al.,
2001
; Zernicka-Goetz,
2002
). Two groups have reported that the earliest specification in
the mouse embryo might occur before the first cleavage event
(Gardner, 2001
;
Piotrowska and Zernicka-Goetz,
2001
; Piotrowska et al.,
2001
). Studies showed that, the orientation of the first cleavage
plane is strongly related to the axes of the blastocyst. The relationship
between the axes of the blastocyst and the axes at the egg cylinder stage has
also been investigated (Weber et al.,
1999
). It was concluded that descendants of cells located near the
polar body at the blastocyst stage tend to contribute to the distal visceral
endoderm in the egg cylinder. The relationship between this axis (near the
polar body - away from the polar body) to the distal-proximal axis at egg
cylinder stages was also examined. From these investigations, a possible
relationship between the earliest axis manifestation and the final embryonic
body axes at later stages was proposed. However, as yet, continuous
observation from the fertilized egg to the post-implantation period has not
been performed. An important issue remains, namely how each cell changes its
relative spatial position in an embryo. A second problem is how cell fates are
specified during early stages of development. To ascertain this it is
important to observe embryos without disturbing the original developmental
program, because the mammalian embryo possesses a high potential to regulate
its development after experimental manipulation. Our aim was to investigate
the inherent developmental program in early stages of normal embryogenesis
with minimal disturbance to the embryonic program.
A major reason for the paucity of experiments attempting to follow cell
lineage in the early mouse embryo is one of technical difficulty. A number of
approaches have been used to mark cells. Individual cells have been labeled
with oil drops (Wilson et al.,
1972
), dyes (Lawson et al.,
1986
; Gardner,
2000
; Tam and Beddington,
1992
), reporter enzymes such as horseradish peroxidase
(Balakier and Pedersen, 1982
),
or by mRNA injection (Zernicka-Goetz et
al., 1997
). However, it has been difficult to follow the lineage
over a long period because these lineage tracers are easily diluted or
alternatively may disturb normal development. We have traced cell lineages in
the early mouse embryo using a Cre-loxP system
(Araki et al., 1995
). After the
activation of Cre recombinase, cells express the bacterially derived
ß-galactosidase enzyme. Activation of the reporter expression is
irreversible and heritable via the changes of genomic organization. This
system allows us to follow the fate of a cell and its progeny for a long
period after activation of Cre recombinase.
We labeled individual cells during the pre-implantation period and analyzed
the distribution of labeled descendants in the post-implantation embryo around
day 8.5. By performing multiple experiments, we have established the
generalized behavior of cells in the embryo after labeling between the
two-cell and four-cell stages of development.
 |
Materials and methods
|
---|
Mouse strains and collection of embryos
CAG-CAT-Z (Araki et al.,
1995
) mice were maintained as a homozygous transgenic colony. To
identify homozygous males, individual animals were crossed with wild-type
females, embryos were collected and subjected to PCR genotyping. Males were
confirmed as homozygous transgenic animals when all embryos were
heterozygotes. For the lineage tracing experiments, wild-type BDF1 females
were used. BDF1 females (Charles River Japan, Inc.) were super-ovulated by
injection of 5 IU pregnant mares serum (PMS; Teikoku Zouki, Japan) followed by
5 IU human chorionic gonadotropin (hCG; Teikoku Zouki, Japan) as described
previously (Hogan et al.,
1994
). Primed females were mated with homozygous CAG-CAT-Z
transgenic males. To obtain one-cell stage embryos, swollen ampullae were
flushed with M2 medium (Specialty Media) containing 0.5 mg/ml of hyaluronidase
(Sigma, H3506). The embryos were washed several times in M2 medium to remove
the cumulus cells. To obtain embryos at the two-cell or four-cell stages,
embryos were flushed from the oviduct.
Activation of Cre recombinase
To activate Cre recombinase in a specific blastomere, we used one of three
methods. One was the injection of an expression vector pBS185 (Gibco-BRL),
which drives expression of bacteriophage P1-encoded Cre recombinase under the
control of human cytomeglovirus early promoter. The second method was
injection of the expression vector CAG-Cre, in which Cre expression is
controlled by a CAG promoter cassette. This plasmid was constructed using the
Cre cDNA fragment from pBS185 subcloned into the pCAG-GS expression vector
(Araki et al., 1995
) to give
pCAG-Cre. Third was the direct injection of Cre recombinase protein
(Clontech). For all of these methods, injections were performed into
blastomere nuclei using conventional injection techniques. Following
injection, embryos were cultured in KSOM medium (Specialty Media) containing
0.1% BSA in an incubator at 37°C, 5% CO2. Healthy embryos were
selected and transferred into the oviduct of pseudo-pregnant females.
Embryo culture
For direct observation at the blastocyst stage, embryos were cultured at
37°C in a drop of KSOM covered with light mineral oil. Alginate gel was
used to maintain the position of embryos in culture
(Gardner, 2001
). Embryos were
put into a drop of KSOM (MR-106-D, Specialty Media) containing 0.7% alginic
acid sodium salt (A-2158, Sigma) and covered with paraffin oil. The alginate
was induced to gel by application of a small amount of solution containing
1.5% CaCl2 and 0.9% NaCl. After 1 minute, residual KSOM containing
alginate was removed. The alginate drop was rinsed more than five times with
alginate-free KSOM, and the alginate gel was covered with KSOM under paraffin
oil. Embryos were cultured in a 37°C, 5% CO2 incubator.
Photographs of embryos in culture were taken with an inverted microscope
(IX70, Olympus) equipped with a 35 mm film camera.
X-gal staining and histology
Manipulated embryos were collected from uteri and fixed with 2%
paraformaldehyde, 0.2% glutalaldehyde in PBS at 4°C for 1 hour. Fixed
embryos were washed with PBS containing 0.1% Triton X-100 at 4°C
overnight, and ß-gal was visualized with X-gal staining buffer (1 µM
MgCl2, 3 mM K4[Fe(CN)6], 3 mM
K3[Fe(CN)6], 0.1% Triton X-100 and 0.05%
5-bromo-4-chloro-3-indolyl-b-D-galactopyranoside (X-gal) in PBS). For the
X-gal staining of embryos embedded in alginate gel, buffer without Triton
X-100 was used. Embryos were incubated in the staining buffer at 37°C
overnight and post-fixed in PBS containing 4% paraformaldehyde and 50 mM EDTA.
Whole-mount images were captured with a M-420 Leica microscope equipped with
CCD camera (Progress 1012, Zeiss Vision). Embryos were dehydrated through 70%
ethanol, 90% ethanol, 100% ethanol and chloroform, and then embedded in
paraffin wax. 7.5 µm serial sections were stained with Haematoxylin and
Eosin. These sections were observed under the microscope (DMRB, Leica), and
photographic images were taken with a CCD camera (Progress 1012, Zeiss
Vision). Images were processed with Adobe Photoshop.
 |
Results
|
---|
The CAG-CAT-Z transgenic mouse is an appropriate model for tracing
cell lineage in early development
We used the transgenic CAG-CAT-Z (Araki
et al., 1995
) mouse line for our experiments. These mice were
designed so that ß-galactosidase becomes active in a cell-autonomous
fashion following activation by Cre recombinase. Control experiments were
performed to ensure that the cell marking system worked appropriately. Mice
homozygous for the CAG-CAT-Z transgene were shown to develop normally and give
rise to viable fertile animals. We crossed homozygous transgenic males to
super-ovulated wild-type BDF1 females to obtain heterozygous embryos.
Following induced activation of Cre recombinase in a specific blastomere,
embryos were transferred into the oviducts of pseudo-pregnant females.
Manipulated embryos were recovered from uteri at approximately day 8.5 of
development and subjected to X-gal staining. We tested several methods for
activating Cre recombinase in the embryo
(Table 1) and compared the
efficiency of labeling. No positive staining was obtained in embryos directly
injected with Cre recombinase protein. We tested two Cre recombinase
expression vectors. The first was pBS185, which drives expression of Cre under
control of the CMV promoter. This induced recombination but the X-gal-positive
embryos was less than 10% of those injected. Next we tested pCAG-Cre, which
utilizes the regulatory sequence of CMV and chick ß-actin and a rabbit
ß-globin poly(A). Injection of this plasmid into the nuclei of specific
blastomeres gave the most consistent results: 30% and 11.2% X-gal-positive
embryos following injection at the 1-cell stage, and at the two-cell stage
respectively. Accordingly we decided to use this method to mark individual
cells.
Initially we examined the efficiency of reporter enzyme activation in
embryos. Fig. 1 shows the
representative embryos recovered following injection of pCAG-Cre into
fertilized eggs. The manipulated embryos developed normally regardless of
expression of the ß-galactosidase reporter. From whole-mount observation,
both embryonic and extra-embryonic tissues were stained, with the exception of
the extra-embryonic membranes (Fig.
1A,E). Next, we performed a detailed examination of serial
sections (Fig. 1B-D,F-H).
Almost all cells in every germ layer were uniformly positive for ß-gal
activity. Variation in strength of staining between layers reflects the
promoter activity in these cell types. Two cell populations persistently
failed to stain; they were the primitive endoderm derivatives including the
parietal endoderm and visceral endoderm. Few X-gal-positive cells were
detected in parietal endoderm and no stained cells were observed in visceral
yolk sac endoderm (Fig. 1H,
arrows). Although all the cells were stained, variation of staining intensity
was apparent in the trophoblast giant cells (data not shown). Most of the
giant cells were clearly stained, however, punctuate staining within the
cytoplasm was seen in some cells. Similar variation in staining was also
observed in chorionic ectoderm. We also stained embryos obtained from
wild-type females crossed with males carrying the reporter cassette in which
the floxed CAT gene had been excised (Fig.
2). The same staining pattern, namely lack of activity in the
extra-embryonic endoderm, was observed
(Fig. 2C, arrow and arrowhead),
suggesting a lack of CAG promoter activity in this cell lineage. Accordingly,
we were unable to trace the lineage of extra-embryonic endoderm derivatives in
this system.

View larger version (97K):
[in this window]
[in a new window]
|
Fig. 1. Expression of ß-galactosidase in embryos injected at the one-cell
stage. Two representative embryos, 1-5 (A-D) and 1-4 (E-H) are shown. (A,E)
Whole mounts and (B-D,F-H) Hematoxylin and Eosin-stained sections. Almost all
parts of the embryos were positive for X-gal staining (A,B,E-F). (C,D,G,H)
Higher magnifications of B and F respectively. (H) Derivatives of primitive
endoderm (arrows) did not show X-gal staining. Scale bars: 300 µm (A,E),
150 µm (B,F), 30 µm (C,G).
|
|

View larger version (69K):
[in this window]
[in a new window]
|
Fig. 2. Distribution of X-gal-positive cells in an embryo derived from a mating
with a F1 male lacking the floxed CAT gene. (A) Whole-mount embryo
in the deciduum. All embryonic cells expressed ß-gal except for the
layers of primitive endoderm origin. (B,C) Sections stained with Hematoxylin
and Eosin. The staining pattern was similar to that of embryos after injection
at the one-cell stage. Almost all cells were stained except for the parietal
endoderm (arrowhead) and visceral yolk sac endoderm (arrow). Scale bars, 300
µm (A), 150 µm (B), 30 µm (C).
|
|
Injection at the two-cell stage resulted in random distribution of
labeled cells
Two-cell stage embryos were collected from oviducts and the expression
vector pCAG-Cre was injected into the nucleus of one blastomere. The embryos
were then transferred into oviducts of pseudo-pregnant females. Eight days
after transfer they were isolated from the uteri and subjected to X-gal
staining. Representative embryos are shown in
Fig. 3. While overtly similar
to the embryos injected at the one-cell stage, staining was more uneven, with
a mixture of X-gal-positive (blue) cells and white cells within a single
embryo. These embryos were serially sectioned and the distribution of stained
cells was examined in detail. We analyzed 37 embryos that were positive for
X-gal staining and morphologically normal. One embryo showed a restricted
distribution of the X-gal-positive cells only in extra-embryonic ectoderm,
while in a second embryo there were no positive cells in the extra-embryonic
ectoderm. In other embryos, the X-gal-positive and X-gal-negative cells were
distributed uniformly throughout all lineages except for the primitive
endoderm derivatives. There was no clear predominance of positive cells along
these body axes. To obtain more quantitative data, we counted the number of
X-gal-positive and -negative cells in nine independent embryos after
Haematoxylin staining. Positive cells were identified by their blue
cytoplasmic X-gal staining. The sections that were counted are shown in low
magnification in Fig. 3B,G,J.
Because we could not see clear preferential distribution, we selected a
representative section of each embryo where we could observe the most tissues.
The number of cells in several different cell layers are shown in the tables
and histogram of Fig. 3E,H,K.
The contribution of X-gal-positive cells in each layer varied between embryos.
An exceptional case, was embryo 2-2 in which the amnion was completely
X-gal-positive (Fig. 3G,H). In
other embryos, the overall number of X-gal-positive cells was in the range of
20-80%. Positive cells made clusters. The contribution of X-gal-positive cells
to the trophoblast giant cells was lower than in other tissues, which may
reflect the coherent growth of this lineage.

View larger version (89K):
[in this window]
[in a new window]
|
Fig. 3. Embryos injected at the two-cell stage showed random distribution of
labeled cells. Embryos showed spotted distribution of X-gal-positive cells,
and this pattern was seen evenly throughout the body axis (A,B) without regard
to any embryonic axes. C and D are higher magnifications of B. In all cell
layers, X-gal-positive and -negative cells were randomly intermingled. (E)
Quantitative analysis. Classification of cell layers, and the result from one
embryo, 2-8, are shown. Left panels indicate the origin of the each layer at
3.5 days and 4.5 days based on classic studies, and the name and abbreviation
of the each tissue is also shown. The actual numbers of positive and negative
cells are given and also represented by the histograms. F,G,H and I,J,K show
the results of embryos 2-2 and 2-34, respectively. In the amnion of embryo
2-2, all the cells were positive for X-gal, however, this was an exceptional
case because positive cells were intermingled with negative cells in other
embryos. Scale bars: 300 µm (A) 150 µm (B), 30 µm (C).
|
|
Injection at the four-cell stage resulted in three types of
distribution of labeled cells
Next we labeled single blastomeres of four-cell stage embryos by nuclear
injection of the expression vector. Embryos were recovered from the uteri 8
days after transplantation and subjected to X-gal staining. Whole-mount images
of the embryo are shown in Figs
4 and
5. A total of 54 positive
embryos were recovered that developed normally. Three classes of staining
patterns were observed.

View larger version (83K):
[in this window]
[in a new window]
|
Fig. 4. Embryos with one type of X-gal distribution after activation of Cre
recombinase in a blastomere at the four-cell stage. (A-G) The results of
embryo 4-7. In this embryo, X-gal-positive cells were distributed randomly
following labeling at the four-cell stage. (A) Whole mount, (B-F) sections
showing that the stained cells were in all cell layers. The numbers of
positive cells were less than in those embryos labeled at the two-cell stage.
(C) In the extra-embryonic ectoderm of this embryo, clusters of relatively
large numbers of X-gal-positive cells were observed. (G) Quantitative
analysis. X-gal-positive cells accounted for between 5% and 31% of the total
cells depending on the tissue. (H-O) The results of another example, embryo
4-6, which showed a similar pattern of distribution of labeled cells. (L,N)
Sections at different positions of the same embryo. (I,J,K) High
magnifications of L. Histograms show that the contribution of labeled cells is
between 5% and 41%, however localization to any specific tissue or position in
the body was not observed. Scale bars: 300 µm (A), 150 µm (B), 30 µm
(C).
|
|

View larger version (89K):
[in this window]
[in a new window]
|
Fig. 5. Two other types of labeled cell distribution after injection at the
four-cell stage. (A-D) In the second type of distribution, only
extra-embryonic ectoderm contained X-gal-positive cells. (A,C) Whole mounts;
(B,D) sections of the embryos in A and C, respectively. In both embryos, a
portion of trophoblast giant cells (arrows) and a part of the ectoplacental
cone is for X-gal-positive. Note that embryo 4-59 (C,D, arrows) has labeled
cells localized to the trophoblast giant cell layer. (E,F) The last type of
cell distribution. In this embryo, there were no X-gal-positive cells in the
extra-embryonic ectoderm region. In other regions, X-gal-positive and
-negative cells were intermingled. Scale bars: 300 µm (A), 150 µm
(B).
|
|
In the first type, the staining patterns were similar to those of the
embryos injected at the two-cell stage
(Fig. 4). 34 embryos were
placed in this category. The X-gal-positive cells were distributed both in
embryonic tissue and extra-embryonic ectoderm. In these cases, X-gal-positive
cells were distributed randomly and there was no clear localization of these
cells along the body axes. We quantified the results by counting the cell
numbers (Fig. 4G,M,O). In all
embryos in this category, the numbers of X-gal-positive cells in all germ
layers were less than that seen after labeling of a blastomere at the two-cell
stage. The ratio of positive to negative cells was variable between the germ
layers. However, no embryo had more than 50% blue cells, and no specific germ
layer showed preferential accumulation of X-gal-positive cells. We could
observe relatively large clusters of positive cells in the extra-embryonic
ectoderm (Fig. 4C), which may
suggest coherent clonal growth in this lineage. In other tissues, such as the
neural ectoderm (Fig. 4D,J),
embryonic ectoderm (Fig. 4E,K)
and embryonic mesoderm (Fig.
4D-F,K), the number of labeled cells in a cluster of
X-gal-positive cells were relatively small and similar to that seen after
injection at the two-cell stage. We also compared the distribution of labeled
cells in different sections of the same embryo
(Fig. 4L-O). We could not
detect any clear preference of localization, which may also suggest random
cell mixing in this embryonic population.
The second and third classes of distribution patterns
(Fig. 5) were different from
the pattern seen after labeling at the two-cell stage. In the second type,
X-gal-positive cells were seen only in the extra-embryonic region
(Fig. 5A-D). The trophoblast
giant cells and some cells in the ectoplacental cone were stained blue. No
positive cells were found in the embryo proper or in the extra-embryonic
mesoderm layers in these embryos, while both X-gal-positive and -negative
cells were found in the extra-embryonic ectoderm. As shown in
Fig. 5D, more labeled giant
cells were seen in one part than the other. In the final class, cells of the
extra-embryonic ectoderm were completely unlabeled, although positive cells
were found in other tissues (Fig.
5E,F). The numbers of embryos obtained from the experiments are
summarized in Tables 2 and
3.
Distribution of labeled cells in the blastocyst
Piotrowska et al. (Piotrowska et al.,
2001
) previously reported that the descendants of one blastomere
in the two-cell stage embryo have a localized distribution in the blastocyst.
Our results from analyzing cell distribution at later stages, however, showed
that the descendants of a single blastomere of two-cell stage embryos
intermingled completely in the embryo. To understand the timing of this cell
mixing, we next examined the distribution of labeled cells in the blastocyst
following labeling of a blastomere of the 2 or the four-cell stage embryo.
This allowed us to address when the change in relative position of cells
occurs; i.e. prior to or following the blastocyst stage. A blastomere of
two-cell stage embryos was injected with the Cre expression vector and embryos
were cultured in vitro until the blastocyst stage. The results are shown in
Fig. 6A-F and in
Table 4. We obtained similar
results to those described by Piotrowska et al.
(Piotrowska et al., 2001
) who
used an alternative labeling method. In 73% of the blastocysts, the labeled
cells stayed in clusters and did not intermingle completely. In
Fig. 6 the boundary between the
embryonic and the abembryonic parts is indicated where visible. This boundary
was clear in 84 out of 147 embryos. Labeled cells were found in the embryonic
part in 35 of those embryos and in the abembryonic part in 26 embryos. These
results indicate that the descendants of a two-cell blastomere could be
located either in the embryonic or the abembryonic part of the blastocyst in
73% of embryos, supporting the idea that the first cleavage plane is nearly
orthogonal to the embryonic-abembryonic axis of the blastocyst. Similar
results were obtained after labeling a single blastomere at the four-cell
stage (Fig. 6G-I and
Table 4), although the number
of X-gal-positive cells was lower than that seen following labeling at the
two-cell stage. To further examine the relationship between the first cleavage
plane and embryonic-abembryonic axis of the blastocyst, we cultured embryos in
alginate gel to maintain the position. Two-cell stage embryos with zona
pellucida were embedded in alginate gel, and cultured in a CO2
incubator. Several embryos were cultured in a drop of alginate gel covered
with alginate-free KSOM. In these drops, the relative positions of embryos
were maintained, and this relative position was used to identify the
orientation of each embryo. At the two-cell stage, polar bodies were located
on or very close to the plane of the first cleavage. Although embryos had zona
pellucidas, the positions of polar bodies did not changed while they were
visible, until the eight-cell stage (data not shown). Because it was not
always easy to reliably distinguish the second polar body from the first polar
body in the living embryo without damage as Gardner described
(Gardner, 2002
), the plane of
the first cleavage was used as a landmark in the two-cell stage embryo.
Photographs of embryos at the two-cell, three-cell and blastocyst stages were
taken and compared. We measured the angle between the plane of the first
cleavage and the embryonic-abembryonic axis of the blastocyst
(Fig. 7A). Those embryos in
which two blastomeres were positioned side by side in a drop of gel were used
to measure the angle of cleavage. Embryos in which the first cleavage plane
was not orthogonal to the bottom of the culture plate were not counted. We
also analyzed the division order of blastomeres at second cleavage in cases
where the angle between first cleavage plane and the embryonic-abembryonic
axis was more than 50°. First, we examined intact embryos without
injection. The angle was analyzed in 123 embryos
(Fig. 7B). Although we observed
a wide variety of distribution, more embryos in which the angle was greater
than 50° (dark red columns in Fig.
7B) were mapped. Out of 123 embryo analyzed, 90 embryos (73%) were
mapped in this category.

View larger version (157K):
[in this window]
[in a new window]
|
Fig. 6. Distribution of labeled cells at the blastocyst stage. Embryos were
cultured in vitro following injection of Cre recombinase into a blastomere at
the two-cell (A-F) or at the 4-cell (G-I) stage. Labeled cells were seen as
clusters in these embryos. The boundary between the embryonic and abembryonic
parts is indicated by the dashed line.
|
|
View this table:
[in this window]
[in a new window]
|
Table 4. Distribution of X-gal-positive cells in blastocysts after labeling
single blastomeres of two-cell or four-cell stage embryos
|
|

View larger version (59K):
[in this window]
[in a new window]
|
Fig. 7. Analysis of alginate-embedded embryos. Normal embryos without injection or
embryos labeled with Cre recombinase were cultured in alginate gel to maintain
their position. (A) The angle (a) between the first cleavage plane (P) and the
embryonic-abembryonic axis (E-Ab) of the blastocyst was measured by comparing
photographs. Only the embryos in which the first cleavage plane was orthogonal
to the bottom of culture plate were used for the analysis. (B) The angle was
found to be 80° in 28 out of 123 of the normal uninjected embryos. The
angle was greater than 50° in 90 embryos (73%) (columns are colored in
dark red). (C) In the case of DNA-injected embryos, similar distribution in
angles were observed after analyzing 71 embryos. (D-I) Examples of uninjected
embryos used for analysis. In the embryo in D-F, the angle was 70°, and
the first blastomere to divide contributed mainly to the embryonic part of the
embryo. In another normal embryo shown in G-I the angle was 80°, and the
early-dividing blastomere contributed to the abembryonic part of the embryo.
(J-Q) Two examples of labeled embryos. (J-M) The injected blastomere divided
later than the other blastomere and contributed to the embryonic part in the
embryo as can be seen from the X-gal staining in this region (M). In the
embryo shown in N-Q, the injected blastomere also divided late. The labeled
cells were seen in the trophectoderm of the abembryonic part and around the
boundary zone. Scale bar: 30 µm.
|
|
This result was consistent with our results of the labeling experiments at
the two-cell stage, and the results reported by two other groups
(Gardner, 2001
;
Piotrowska et al., 2001
). We
then analyzed the order of cell division of two-cell blastomeres. It is known
that mammalian two-cell blastomeres divide asynchronously. Piotrowska et al.,
(Piotrowska et al., 2001
)
suggested that the cell that is first to divide preferentially contributes to
the embryonic part of the blastocyst. We examined this possibility using an
alternative method. Embryos embedded in alginate gel were observed and
photographed every 30 minutes until most of the embryos had developed to the
three-cell stage, and then they were allowed to develop to the blastocyst
stage in a 37°C CO2 incubator. Out of 90 embryos in which the
angle between the first cleavage plane and the embryonic-abembryonic axis of
the blastocyst was greater than 50°, we could capture images of the
three-cell stage in 67 embryos. The first blastomere to divide was seen in the
position corresponding to the future embryonic part of the blastocyst in 31
embryos (46%), as is shown in Fig.
7D-F. However, in 36 embryos (54%) the first two-cell blastomere
to divide was located in the future abembryonic part. Thus in our experiments,
intact embryos did not show a clear correlation between the division order of
the two-cell blastomeres and the axis of the blastocyst. We then analyzed this
relationship in embryos injected with Cre expression plasmid. These injected
embryos were fixed and subjected to X-gal staining following the same
procedures used for uninjected embryos. A summary of the angles between the
first cleavage plane and the embryonic-abembryonic axis of the blastocysts is
shown in Fig. 7C. Variation in
the angle was similar to that of un-injected embryos. The distribution was
also similar to that of uninjected embryos with the peak of distribution at
around 80°. Out of 71 embryos analyzed, 56 embryos (79%) were in the
category in which the angle was more than 50°
(Fig. 7C, dark red columns).
This result suggests that injection of expression plasmid did not have a clear
influence on the specification of the embryonic-abembryonic axis in the
blastocyst. We also analyzed division order of two-cell blastomeres in 18
embryos. The injected blastomere divided first in four embryos (22%). Out of
these 4 embryos, X-gal-positive cells were found in the embryonic half in two
and in the abembryonic half in the other two. The injected blastomere divided
later in 14 embryos (78%). Nine embryos had X-gal-positive cells in the
embryonic part, and in five embryos they were mainly in the abembryonic part.
These results suggest that injection of DNA may slightly affect the timing of
cell division. However, the labeled cells could contribute to both embryonic
and abembryonic parts of the blastocyst.
 |
Discussion
|
---|
We carried out cell lineage tracing experiments in the early mouse embryo.
We took advantage of a recombination-based method, using expression of Cre
following permanent activation of reporter gene expression to label a specific
cell. This method generates a non-diffusible, cell-autonomous and heritable
marking of cells, and thus is suitable for lineage tracing during extended
periods of embryonic development. Only low levels and transient expression of
Cre recombinase is required to mediate recombination of the target DNA. We
first tested the efficiency of the cell labeling method with Cre recombinase
in heterozygous CAG-CAT-Z (Araki et al.,
1995
) transgenic mouse embryos. Injection of the Cre expression
plasmid (pCAG-Cre) gave the most consistent results for cell labeling. The
manipulated embryos recovered from the uteri were morphologically normal,
indicating that the manipulation did not perturb normal development. After
injection at the one-cell stage, the majority of cells became positive for
X-gal staining with the exception of cells derived from the primitive
endoderm. The lack of X-gal staining in these cells was not due to inefficient
recombination in their cell lineage because no negative cells were found at
the blastocyst stage after labeling at the one-cell stage (data not shown),
and embryos derived from matings with males that carry a transgene without
theCAT gene insertion showed a similar staining pattern. Lack of staining
suggests that the CAG expression cassette inserted in this transgenic mouse
was silent or very weakly expressed in these cells. However, combining the
CAG-CAT-Z transgene and pCAG-Cre provided a very suitable system for analyzing
other cell lineages in developing embryos. An important concern is the effect
of the manipulation on the developmental potential of the cell. For example,
the manipulation may disturb the normal fate of the marked cell or affect the
potential to contribute to a certain type of tissue. This possibility,
however, was ruled out by the results of marking a single blastomere in
two-cell and four-cell stage embryos. Analysis of the cleavage pattern of
two-cell blastomeres following the injection showed that the manipulation had
only a slight effect on the timing of cell division. However, the relationship
between the first cleavage plane and the embryonic-abembryonic axis of the
blastocyst was not affected. The manipulated blastomere could contribute to
both embryonic and abembryonic parts in the blastocyst, suggesting that the
manipulation did not affect the specification of cell fates. We also found the
labeled cells could contribute to the cell types examined in 8.5-day embryos.
Accordingly in this paper we exploited this transgenic mouse to analyze cell
lineage.
After marking a blastomere at the two-cell stage, we observed random
distribution of the labeled cells in 8.5-day embryos. These results suggested
that rearrangement of the relative position of descendant cells from the two
blastomeres occurred in the embryo between the two-cell stage and 8.5-day, the
stage at which we performed the analysis. Although there are experiments
showing indirectly incoherent clonal outgrowth of cells in the epiblast
(Gardner and Cockroft, 1998
;
Weber et al., 1999
), our
report is the first observation to clearly demonstrate this phenomenon by
tracing the cell lineages in normal mouse embryos over a long developmental
period. This feature of cell mixing during early embryogenesis is specific to
the mouse embryo. For example, in Xenopus embryos, a relatively clear
segregation of cells along the left-right body axis is observed after labeling
of a blastomere at the two-cell stage
(Jacobson and Hirose, 1978
).
The first cleavage plane thus reflects the midline of the resulting tadpole.
Our results suggest the program of embryonic development in mouse has inherent
plasticity and that mixing of epiblast cells along the body axis happens
during early stages. Piotrowska et al.,
(Piotrowska et al., 2001
)
showed that descendants of a two-cell stage blastomere analyzed in the
blastocyst tend to be localized and the cells originating from one blastomere
form a distinct boundary with those cells derived from the other blastomere.
Similarly, we showed, with our labeling method, that drastic cell mixing does
not occur until the blastocyst stage (Fig.
6 and Table 4),
which is consistent with previous reports. Our data suggests that extensive
intermingling of cells occurs subsequently to the blastocyst stage. The
coherency of clonal cell growth may vary between cell layers
(Gardner and Cockroft, 1998
;
Weber et al., 1999
). Gardner
and Cockroft and Weber et al. suggested that there is coherent clonal
outgrowth of visceral endoderm and incoherent growth of ICM derivatives.
Unfortunately, we could not examine the coherency of growth in cells
originating from the primitive endoderm because our reporter system failed to
mark primitive endoderm derivatives. The previous work of Gardner and a
colleague (Gardner and Cockroft,
1998
) showed cell mixing in the epiblast of chimaeric embryos
after transplantation of cells into the blastocyst. They also showed two
daughter cells located in non-adjacent positions following cell division in a
non-chimeric 7-day epiblast, suggesting possible incoherent clonal growth in
this cell layer. We could clearly show incoherent clonal growth of cells
during development of the non-chimeric embryo, showing that cell mixing occurs
in the post-implantation embryo. In contrast, we found larger patches of
labeled cells in the trophoblast giant cell population
(Fig. 3J and
Fig. 5), indicating relatively
coherent clonal growth of cells in this tissue.
Recently, the relation between the sperm entry point, the first cleavage
plane, and the embryonic-abembryonic axis of the blastocyst has been discussed
(Gardner, 2001
;
Piotrowska and Zernicka-Goetz,
2001
; Davies and Gardner,
2002
). Piotrowska et al.
(Piotrowska et al., 2001
)
proposed a model in which the first cleavage plane is determined by the
position of the polar body and the sperm entry point. However, other groups
claimed that the first cleavage plane is not related to the sperm entry point
(Davies and Gardner, 2002
). It
is also suggested that the first cleavage plane is nearly orthogonal to the
embryonic-abembryonic axis of the blastocyst
(Gardner, 2001
;
Piotrowska et al., 2001
).
Kelly et al. (Kelly et al.,
1978
) suggested that the descendants of the first cell to divide
to produce a three-cell stage embryo divide ahead of those cells derived from
its slower partner. They also showed that the first pair of cells to reach the
eight-cell stage contribute more descendants to the ICM than the last cell to
produce the eight-cell stage. Piotrowska et al.
(Piotrowska et al., 2001
)
suggested that a two-cell blastomere that divides early tends to contribute to
embryonic parts of the blastocyst. However, their data revealed that neither
of the two blastomeres contributes exclusively to the cells of the ICM. We
also analyzed the relationship between the first cleavage plane, the division
order of two-cell blastomeres and the orientation of embryo-abembryonic axis
of the blastocyst. The labeling of the two-cell stage blastomere resulted in
labeled cells in either the embryonic or in abembryonic part of blastocyst in
73% of embryos. This suggests a strong relationship between the first cleavage
plane and the axis of the blastocyst. We observed a similar relationship by
analyzing the embryos embedded in the alginate gel. It seemed that the
variation of the angle of the first cleavage plane and the embryonic axis was
greater in our results than those reported before. This may be because of the
difference in mouse strains and methods. Gardner showed that the angle differs
somewhat between mouse strains (Gardner,
2001
). Our results support the idea that the first cleavage plane
is close to orthogonal to the embryonic-abembryonic axis of the blastocyst.
However, our results of the relationship between the cell division order and
the allocation of the ICM in the blastocyst were different from that
previously reported (Piotrowska et al.,
2001
). We observed that the first cell to divide contributed to
the embryonic part in 46% of uninjected normal embryos in which the angle
between the first cleavage plane and the axis of the blastocyst was greater
than 50°. The early dividing blastomere was located to the abembryonic
part of the blastocyst in the remaining 54% of embryos
(Fig. 7G-I). We could only
examine the topological relationship in this study although the position of
cells in embryos were basically maintained in zona pellucida, and we did not
trace actual cell lineages from three-cell to blastocyst stages because of
technical limitations. As we noticed that the DNA injection affected division
order of two-cell stage blastomeres, alternative methods are needed to further
examine this relationship. In the manipulated embryos, labeled blastomeres
contributed to both the embryonic part and the abembryonic part. The variation
in angle between the first cleavage plane and the axis of the blastocyst was
similar to that of normal embryos. These suggest that the DNA injection has
little effect on the specification of cell fate, and division order does not
necessarily determine the fates of blastomeres. Our results after the labeling
at the two-cell stage showed that the descendants of either blastomere
contribute to embryonic tissues at every axial level of the resulting
embryonic body in 8.5-day embryos. These suggest that incoherent clonal growth
and drastic cell mixing occurs later than the blastocyst stage, and that the
actual body axis may be specified during later development.
The earliest segregation of cell fate is believed to occur between
trophectoderm and inner cell mass during formation of the blastocyst. Our data
corroborate this. We classified the pattern of labeled cell distribution.
Labeling at the four-cell stage resulted in three types of distribution
patterns of X-gal-positive cells in 8.5-day embryos. The first pattern type
was similar to the results of labeling at the two-cell stage, namely that the
X-gal-positive cells were distributed randomly in all germ layers. As
expected, the number of positive cells was less than that seen in the embryos
labeled at the two-cell stage, and there were no specific biases in
contribution. In the second type, only trophoblast giant cells and part of the
ectoplacental cone were positive. This suggests that one blastomere of the
four-cell stage embryo contributed exclusively to extra-embryonic tissue.
However we note that in these embryos, X-gal-negative cells were also present
in these extra-embryonic layers suggesting that descendants of other
blastomere also contributed to this lineage. The reciprocal patterns of
staining were also seen where X-gal-positive cells were absent from
extra-embryonic ectoderm, while contributing to the embryo proper. These
results also ruled out the possibility that the manipulation affected the
potential of the cell to differentiate into certain cell lineages. From these
data, we can propose two models for the segregation of cell fate. The first
possible explanation is that the specification of cells starts at the
four-cell stage and one blastomere can contribute only to the extra-embryonic
tissue, another one only contributes to other cell types, and the remaining
two retain the potential to contribute to both tissues. However, because we
could not test the variation between embryos, the possibility remains that
this segregation of future cell fates is not a common event observed in all
the embryos. We should also pay attention to the loss of cells during
development. The limitation of this type of experiment is that we can analyze
only surviving cells and not the pattern of cell death that is routinely seen
during development. An alternative explanation for the segregation of fate is
that it depends on the cleavage pattern and the position of blastomeres in an
embryo as proposed by Wilson et al.
(Wilson et al., 1972
). In
their model, the cortical cytoplasm contributed to the trophectoderm and the
deep cytoplasm mostly contributed to the ICM. Considering this, it might be
possible that if the cleavage pattern and the allocation of blastomeres are
relatively fixed, developmental programs of each blastomere are also regulated
via these factors. To resolve this issue, we need to examine embryos in
further detail and obtain statistical data. In any case, our data do not show
that the cell fate is determined in this stage. As Kelly showed clearly
(Kelly, 1977
), all of the
isolated four-cell stage blastomeres still posses the potential to contribute
to all cell types of the body. In conclusion, our experiments show that the
first segregation of cell fates is between ICM and trophectoderm, and that it
may happen between the two-cell and four-cell stages.
 |
ACKNOWLEDGMENTS
|
---|
We thank Dr Elizabeth J. Robertson for critical reading and discussion of
this manuscript, Drs Masatoshi Takeichi and Goro Eguchi for continuous
encouragement and helpful discussions, and Dr Ruth Yu for comments on this
manuscript. We acknowledge the excellent technical assistance of Tsutomu Obata
and the help of people in the animal facility in this department.
 |
Footnotes
|
---|
* PRESTO, Japan Science and Technology Corporation (JST) 
 |
REFERENCES
|
---|
Araki, K., Araki, M., Miyazaki, J. and Vassalli, P.
(1995). Site-specific recombination of a transgene in fertilized
eggs by transient expression of Cre recombinase. Proc. Natl. Acad.
Sci. USA 92,160
-164.[Abstract]
Balakier, H. and Pedersen, R. A. (1982).
Allocation of cells to inner cell mass and trophectoderm lineages in
preimplantation mouse embryos. Dev. Biol.
90,352
-362.[Medline]
Beddington, R. S. and Robertson, E. J. (1999).
Axis development and early asymmetry in mammals. Cell
96,195
-209.[Medline]
Davies, T. J. and Gardner, R. L. (2002). The
plane of first cleavage is not related to the distribution of sperm components
in the mouse. Hum. Reprod.
17,2368
-2379.[Abstract/Free Full Text]
Gardner, R. L. (2000). Flow of cells from polar
to mural trophectoderm is polarized in the mouse blastocyst. Hum.
Reprod. 15,694
-701.[Abstract/Free Full Text]
Gardner, R. L. (2001). Specification of
embryonic axes begins before cleavage in normal mouse development.
Development 128,839
-847.[Abstract/Free Full Text]
Gardner, R. L. (2002). Experimental analysis of
second cleavage in the mouse. Hum. Reprod.
17,3178
-3189.[Abstract/Free Full Text]
Gardner, R. L. and Cockroft, D. L. (1998).
Complete dissipation of coherent clonal growth occurs before gastrulation in
mouse epiblast. Development
125,2397
-2402.[Abstract/Free Full Text]
Hogan, B., Beddington, R., Costantini, F. and Lacy, E.
(1994). Manipulating the Mouse Embryo: A Laboratory
Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory
Press.
Jacobson, M. and Hirose, G. (1978). Origin of
the retina from both sides of the embryonic brain: a contribution to the
problem of crossing at the optic chiasma. Science
202,637
-639.[Medline]
Kelly, S. J. (1977). Studies of the
developmental potential of 4- and 8-cell stage mouse blastomeres.
J. Exp. Zool. 200,365
-376.[Medline]
Kelly, S. J., Mulnard, J. G. and Graham, C. F.
(1978). Cell division and cell allocation in early mouse
development. J. Embryol. Exp. Morphol.
48, 37-51.[Medline]
Lawson, K. A., Meneses, J. J. and Pedersen, R. A.
(1986). Cell fate and cell lineage in the endoderm of the
presomite mouse embryo, studied with an intracellular tracer. Dev.
Biol. 115,325
-339.[Medline]
Piotrowska, K., Wianny, F., Pedersen, R. A. and Zernicka-Goetz,
M. (2001). Blastomeres arising from the first cleavage
division have distinguishable fates in normal mouse development.
Development 128,3739
-3748.[Abstract/Free Full Text]
Piotrowska, K. and Zernicka-Goetz, M. (2001).
Role for sperm in spatial patterning of the early mouse embryo.
Nature 409,517
-521.[CrossRef][Medline]
Tam, P. P. and Beddington, R. S. (1992).
Establishment and organization of germ layers in the gastrulating mouse
embryo. Ciba Found. Symp.
165, 27-49.[Medline]
Tam, P. P., Gad, J. M., Kinder, S. J., Tsang, T. E. and
Behringer, R. R. (2001). Morphogenetic tissue movement and
the establishment of body plan during development from blastocyst to gastrula
in the mouse. BioEssays
23,508
-517.[CrossRef][Medline]
Weber, R. J., Pedersen, R. A., Wianny, F., Evans, M. J. and
Zernicka-Goetz, M. (1999). Polarity of the mouse embryo is
anticipated before implantation. Development
126,5591
-5598.[Abstract/Free Full Text]
Wilson, I. B., Bolton, E. and Cuttler, R. H.
(1972). Preimplantation differentiation in the mouse egg as
revealed by microinjection of vital markers. J. Embryol. Exp.
Morphol. 27,467
-469.[Medline]
Zernicka-Goetz, M. (2002). Patterning of the
embryo: the first spatial decisions in the life of a mouse.
Development 129,815
-829.[Medline]
Zernicka-Goetz, M., Pines, J., McLean Hunter, S., Dixon, J. P.,
Siemering, K. R., Haseloff, J. and Evans, M. J. (1997).
Following cell fate in the living mouse embryo.
Development 124,1133
-1137.[Abstract/Free Full Text]