Download The development of animal cap cells in Xenopus: the effects of

Development 101, 23-32 (1987)
Printed in Great Britain © The Company of Biologists Limited 1987
23
The development of animal cap cells in Xenopus: the effects of
environment on the differentiation and the migration of grafted
ectodermal cells
E. A. JONES and H. R. WOODLAND
MRC Animal Development Group, Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, UK
Summary
We have used blastocoel and vegetal pole grafts to
investigate the effect of environment on differentiation and movement of animal pole cells of Xenopus.
In the blastocoel of embryos earlier than stage 10,
fragments of animal pole primarily form mesoderm.
The cells are either integrated into normal host tissues
or they organize a secondary posterior dorsal axis. If
either host or graft is later than stage 9 the graft
forms ectoderm and its cells all migrate into the host
ectoderm. Inner layer animal cells form sensorial
layer; outer cells move to the epidermis. Thus considerable powers of appropriate movement are seen.
In the vegetal pole no movement occurs. If the graft is
stage 9 or earlier, or the host is stage 1(H or earlier,
the graft forms mesoderm, including striated muscle
in the gut. This shows that muscle can develop in
wholly the wrong environment, it suggests that the
dorsal inductive signal from mesoderm is rather
general in the vegetal mass and suggests that dorsal
mesoderm development involves little subsequent
adjustability. If the host is stage 11 or later, or the
graft later than stage 9, the graft forms epidermis in
the gut. This shows that the epidermal pathway of
development is also insensitive to environment.
Introduction
isolated animal hemispheres only form epidermis
(Holtfreter & Hamburger, 1955; Asashima & Grunz,
1983; Slack, 1984; Jones & Woodland, 1986). It is
believed that this mesoderm is formed by the inducing action of cells in the presumptive endoderm
on competent ectoderm, the latter being reported to
be able to respond to this induction up to gastrulation
(Dale, Smith & Slack, 1985). In experimental tissue
combinations, at least, the presumptive ectoderm
may also form pharyngeal endoderm (Sudarwati &
Nieuwkoop, 1971). Thus, in atypical sites, presumptive epidermis might be expected to form mesoderm
and anterior gut, in addition to the epidermis or
nervous system that it normally forms.
When single cells are placed in the blastocoel of a
host embryo their descendants appear in a variety of
tissues and the cells concerned apparently conform to
the differentiated state of their surroundings (Wylie,
Smith, Snape & Heasman, 1985; Wylie, Snape, Heasman & Smith, 1987; Snape, Wylie, Smith & Heasman, 1987). Do they differentiate in accordance with
In this paper, we describe experiments using monoclonal antibodies to epidermis and muscle-specific
epitopes to investigate the migration, development
and subsequent differentiation of animal cap cells of
Xenopus embryos when they are transplanted into
unusual positions in the embryo.
In Xenopus, the ectoderm is primarily derived from
the pigmented half of the embryo (Keller, 1975;
Cooke & Webber, 1985; Dale & Slack, 1987) though
even vegetal pole cells of the 32-cell embryo give
rise to a little ectoderm at high frequency (Heasman, Wylie, Hausen & Smith, 1984). The ectoderm
eventually produces two main components, epidermis and nervous system, a process involving a number
of steps of commitment, first to ectoderm rather than
mesoderm and subsequently to either epidermis or
nervous system. Recent fate mapping shows that the
animal cap region also forms much of the mesoderm
(Cooke & Webber, 1985; Dale & Slack, 1987) though
Key words: Xenopus, animal cap cells, migration, graft.
24
E. A. Jones and H. R. Woodland
their surroundings or do they settle on a differentiation pathway and then move to the appropriate
site? Indeed, how much are migratory abilities of cells
responsible for maintaining and achieving the three
germ layer structure of the embryo? In this paper, we
show that ectodermal cells have considerable ability
to migrate to their appropriate location in an embryo,
but that this location is not necessary for them to form
epidermis. Similarly, muscle can develop in completely unusual surroundings, although mesbderm
cells probably also have migratory abilities around
the general blastocoel region. The picture that
emerges is first that the structure of the embryo is
probably maintained by sophisticated migratory abilities in its constituent cells. Second, it seems that once
certain major choices in differentiation pathways are
made, cells differentiate autonomously.
We have recently isolated two monoclonal antibodies that react specifically with the epidermis and
striated muscle of the amphibian embryo. The epidermal marker reacts with all of the surface epidermal
cells of the neurula, even though these cells may be as
different as the cement gland and ciliated cells (Jones,
1985; Jones & Woodland, 1986). This antigen, which
first appears in the stage-12^ late gastrula, is a major
secreted molecule with a protein component, present
in all superficial cells of the early neurula, except the
future central nervous system. It can be used as a
marker of the appearance of the epidermal phenotype, even when cells do not gain the morphological
characteristics of epidermis. For example, the marker
subsequently appears when cell division is blocked at
the mid-blastula stage, even though the cells become
multicellular and disorganized (Jones & Woodland,
1986). The muscle-specific marker is a monoclonal
antibody (5A3.B4) raised by immunizing Balb/c mice
with a homogenate of adult Xenopus muscle. It stains
striated muscle from stage 20 onwards and reacts with
no other tissue type (Fig. 1). We also used a further
muscle-specific antibody (Kinter & Brockes, 1985).
All the antibodies used in this study stain X. laevis and
X. borealis in an identical way.
Methods
Embryo culture, manipulations and histology
Embryos were cultured and explants made as described by
Jones & Woodland (1986). Ectodermal sandwich experiments were made with ectodermal explants from X.
borealis sandwiched between complete animal caps derived
from two X. laevis stage-9 blastula and incubated. They
were fixed when embryos synchronous with the implant had
reached stage 19. X. borealis cells were recognized by
staining with quinacrine. They exhibit intensely fluorescent
chromatin granules, which are absent from X. laevis
(Fig. 1A; Thi^baud, 1983).
Blastocoel-grafted embryos were made by inserting rhodamine-labelled or X. borealis donor ectoderm into a small
slit at the animal pole of demembranated embryos. Pieces
of ectoderm were approximately one eighth of an animal
cap in size. They were cultured in MBS [88mM-NaCl;
lmM-KCl; 24mM-NaHCO3; 15mM-Tris-HCl; 0-33 mMCa(NO3)2; lmM-MgSO4; lmM-NaHCO,; 2mM-sodium
phosphate pH7-4; and 0-1 mM-Na2EDTA (Gurdon, 1977)]
to heal and then transferred into 1/10 MBS to gastrulate.
Vegetal pole grafts were achieved by grafting similar
explants into gaps teased between vegetal pole cells or into
the holes left after removing whole vegetal pole blastomeres. All grafted embryos were healed in MBS. They
were either maintained in this medium to produce exogastrulae or transferred to 1/10 MBS to gastrulate normally.
Fixation, embedding, sectioning and staining with antibodies or simpler chemicals were as described by Jones &
Woodland (1986). Fig. 1B,C shows the normal staining
pattern of the epidermal and muscle-specific antibodies on
stage-46 X. laevis embryos.
Results
Migration and differentiation of epidermal cells in
ectodermal sandwiches
In a normal embryo, the cells that form epidermis,
the outermost layer of which binds 2F7.C7, bound the
embryo. The same is very largely true when a morula
or blastula explant of animal cap cells is cultured in
saline, although in this case there is also a scattering
of somewhat more lightly stained cells within the solid
ball of 'atypical epidermis' which forms (Jones &
Woodland, 1986). Can highly pigmented ectodermal
cells differentiate into the strongly positive phenotype in an internal position or is it essential that they
migrate to the cell surface before differentiating in
this way?
To find if this was so, we made a sandwich of two
animal caps from stage-9 blastulae and placed a
smaller piece of animal cap from embryos of various
stages in the centre (Fig. 2A). Implants were taken
from embryos between stage 3 and stage 9 (8-cell to
late blastula) and placed in stage-9 tissue. In every
case, greater than 20 in total, a high proportion of the
implant, and all of its heavily pigmented cells, bound
2F7.C7 strongly. However, the great majority did so
without moving to the surface (Fig. 2B-D). An
outside position is thus not necessary to form epidermis and migration to the surface of the explant does
not occur. However, the overall environment of the
explants is still ectodermal.
Migration of ectodermal cells in blastocoel grafts
Since ectodermal cells do not migrate in the wholly
ectodermal environment of an ectodermal sandwich,
Development of animal cap cells in Xenopus
25
we have tested the ability of ectodermal cells to
migrate after grafting into different regions of the
whole embryo. Initially, ectoderm was grafted into
the blastocoel. Classically this operation was used as a
test of the ability of the dorsal mesoderm to induce a
secondary CNS, that is as a modification of the
original Spemann and Mangold graft (Spemann &
Mangold, 1924). As pointed out by Slack (1983), this
kind of experiment introduces the graft into variable
situations, with complex results, at least in terms of
the overall tissue organization of the embryo. However, we have grafted animal cap and not dorsal
mesoderm, and primarily ask three very simple questions: do the grafted cells migrate and, if so, what
kind of tissue do the grafted cells enter and what
differentiated phenotype do they display? Similar
approaches have been used in Xenopus with single
ectodermal and endodermal cells (Heasman et al.
1984; Wylie et al. 1985) and in axolotls with multiple
disaggregated cells from a number of germ layers
(Boucaut, 1974a,b). In mammals, the analogous technique is to inject cells into the blastocyst (Gardner,
1985).
When ectoderm from pregastrula embryos is
grafted into the blastocoel of similar, but not
necessarily identical, stages of embryos, secondary
embryonic axes were often formed. Embryos with
both normal and secondary embryonic axes were
serially sectioned and the location of grafted cells and
their differentiated phenotype recorded.
Tissue distribution of grafted cells
Fig. 1. Characterization of cellular and differentiation
markers on cryostat sections. (A) Grafted Xenopus
borealis cells (arrowed) in the somites and nervous system
of a host Xenopus laevis embryo showing the
characteristic punctate nuclear pattern when stained with
quinacrine. Host cellsfluorescediffusely.
(B) Cryostat section through a stage-19 Xenopus laevis
embryo stained with the epidermal marker 2F7.C7. Only
the outer layer of the epidermis is stained.
(C) Longitudinal cryostat section through a stage-46
Xenopus laevis tadpole tail stained with the muscle
marker 5A3.B4. The somites only are stained, showing a
characteristic striated pattern. Abbreviations: a,
archenteron; ep, epidermal ectoderm; g, graft; ns,
nervous system; m, notochord; s, somitic muscle. Bar:
(A,C)75^m; (B) 150^m.
When ectoderm, either from X. borealis or rhodamine-labelled X. laevis donor embryos, was taken
prior to stage 10 and grafted into the blastocoel of
X. laevis host embryos earlier than stage 10, it was
mainly found in the mesoderm of the host, although
some cells were also found in the ectoderm (Fig. 3A).
No cells remained in the blastocoel. The grafted cells
were usually integrated into all parts of the somitic
and lateral plate mesoderm, though none were found
in the notochord. No grafted cells were ever seen in
the endoderm, though experiments of Heasman et al.
(1984), using single cells, show that ectodermal cells
from these stages can be found in this location.
Grafted cells had the morphology typical of their
surroundings, which suggested that they had adopted
the appropriate differentiated phenotype. They were
tested with respect to two cell types, striated muscle
and epidermis. As in control embryos, 2F7.C7 bound
only to the outer ectodermal layer of the embryo, and
this included some of the grafted cells. We identified
muscle cells using the two monoclonal reagents described in the methods. Grafted cells found in the
myotomes reacted appropriately with these antibodies, even in cases where the embryos were quite
26
E. A. Jones and H. R. Woodland
abnormal, proving that they had become differentiated as muscle (Fig. 3A).
The tissue distribution in double-axis embryos was
the same as in morphologically normal embryos,
except that in single-axis embryos grafted cells were
more frequently seen in lateral plate mesoderm or
epidermis. In double-axis embryos, grafted cells were
found more exclusively in the dorsal mesoderm.
In this kind of experiment, the competence of the
animal cap cells to form mesoderm was lost at about
stage 10 and 10i. The capacity of the host to induce
mesoderm formation also disappeared by stage 10
2A
(Table 1). Thus, one can test the migratory capacity
of cells that will later form ectoderm either by
grafting stage-10 animal cap cells into a blastocoel of
hosts at any stage, or by grafting pre-stage-10 cells
into hosts at stage 10 or later.
When the implanted donor ectoderm was derived
from embryos at stage 10 or later, the final positions
of the grafted cells were quite different. They all
moved to the surface of the embryo and became
incorporated into the ectoderm, where they formed
either epidermis, including both the outer 2F7.C7positive epidermal layer and the negative, inner,
sensorial layer, or else they formed nervous system
(Fig. 3B). Often these grafts formed a blister, fold or
pouch in the skin, usually ventrally, possibly simply
because there were larger numbers of epidermal cells
than usual in this region (Fig. 3C). However, the
morphology of these regions was always typical of
larval epidermis.
These experiments suggest two conclusions. First,
migration to the correct site (the outside of the
embryo) is part of the ectodermal phenotype. Since
the cells that form mesodermal tissues also become
incorporated into normal/mesodermal structures,
this conclusion also seems to be true of mesoderm,
although the very frequent occurrence of secondary
mesodermal axes suggests that this ability is limited.
Second, animal cap cells are determined to form
ectoderm by the early gastrula stage and have lost the
ability to form mesoderm. However, this conclusion
applies only to tissue fragment grafts into the blastocoel.
We also tested the migration of the inner and outer
ectodermal layers from donor embryos in blastocoel
grafts. During normal development of Xenopus,
the outer layer in non-neural regions predominantly
forms epidermis, the inner layer forming the so-called
Fig. 2. Epidermal differentiation in ectodermal
sandwiches. Ectodermal fragments were dissected from
three different stage-9 embryos and the sandwich
constructed as in A. The internal fragment was taken
from X. borealis and the outer caps from X. laevis.
Sandwiches were incubated until control embryos were
stage 19, fixed and processed, as described in the
methods. Sections were stained with 2F7.C7 and
rhodamine-conjugated FITC RAM IgG as the second
step antibody. Grafted X. borealis cells were identified by
quinacrine staining. (B) Cryostat section through a
representative graft containing region of a sandwich made
from stage-9 embryos, illuminated to show antibody
binding. The arrow indicates grafted 2F7.C7-positive
cells. The arrowed region is enlarged to show antibody
binding (C) and the punctate quinacrine staining
diagnostic of graft-derived X. borealis cells. Abbreviations
as in Fig. 1. Bar (B) 180fim; (C,D) 36^m.
Development of animal cap cells in Xenopus
'sensorial layer'. In neural regions, both contribute to
the developing nervous system. When grafted separately into stage-9 hosts, both layers from stage-9
27
embryos were incorporated into mesoderm, epidermis and CNS. Both layers from stage-10 embryos
entered only epidermis and CNS, predominantly
Fig. 3. Tissue distribution of ectodermal cells grafted into the blastocoel. Small pieces of A", borealis ectoderm were
inserted into the blastocoel of X. laevis host embryos and the grafted embryos allowed to develop to stage 25-28. The
embryos were then fixed and processed as described in the Methods and stained with the muscle marker, 5A3.B4, or
the epidermal marker 2F7.C7, and these antibodies were revealed with rhodamine RAM IgG and counterstained with
quinacrine to identify the grafted X. borealis cells. Cells from stage-9 X. borealis grafted into stage-9 X. laevis hosts (A)
were found mainly in the mesoderm, in this case the somites (arrowed), and stained strongly with the muscle marker.
When ectoderm from stage-9 X. borealis was grafted into stage-10 hosts (B) grafted cells were found exclusively in the
ectoderm and stained with 2F7.C7, when in the outer epidermal layer. Occasionally double epidermal folds or blisters
were formed from this latter graft (C) which also showed the normal epidermal staining with 2F7.C7. D-F show
quinacrine staining of the same regions, identifying X. borealis-derived grafted cells. Abbreviations as in Fig. 1. Bar:
(A,B,D) 75^m; (C,E,F) 150^m.
E. A. Jones and H. R. Woodland
28
Table 1. Final destination of ectodermal cells grafted into the blastocoels of host embryos
Graft present in
Donor stage
Host stage
Number
analysed
Ectoderm
Mesoderm
Endoderm
Secondary
axis
6i
9
9 inner
9 outer
9
9
3
3
3
0
2
9
9
9
9
3
3
9
3
3
8
3
3
0
0
0
6
2
1
10
5
5
0
0
0
9
9
9
2
3
3
2
3
3
0
0
0
0
0
0
0
0
0
10
10 inner
10 outer
returning to their original inner or outer locations in
the host embryos.
Migration is absent in cells grafted into the vegetal
pole, but epidermal and muscle differentiation is
unaffected
Perhaps the most unusual position for ectoderm to be
grafted is into the vegetal pole region of host embryos. This region normally forms the internal border
of the gut. Can ectodermal cells migrate from such a
position and do they differentiate? The results of
these experiments are summarized in Table 2A,B.
The grafted embryos were stained sequentially with
the epidermal marker and then the muscle marker,
since one of the likely outcomes of such a graft might
be its induction to form mesoderm. Table 2A shows
the summary of results of grafting donor ectoderm
from stage 6- to-12 embryos into the vegetal poles of
hosts at stage 9 or 10. Grafted embryos gastrulated
normally resulting in the graft being internalized into
the gut region. The grafts were either found as
coherent tissue masses completely surrounded by
endoderm, or in regions bordering the lumen of the
developing gut. No cells migrated away. All grafts
from stage 6- to-9 embryos were found to express the
muscle marker, indicating that the ectoderm had
been induced to form muscle (Fig. 4). None of these
grafts expressed the epidermal marker, though some
of the grafted cells were negative with both antibodies. The differentiation of grafted cells into notochord, assessed purely by morphological criteria, was
not detected. In all grafts from stage 10 or later, the
graft always expressed the epidermal marker (Fig. 5)
and did not express any muscle-specific determinants.
These results show that epidermal differentiation can
take place in an environment as unusual as the centre
of the gut. They also show that the extreme vegetal
pole of an embryo is capable of inducing dorsal
mesoderm in competent ectoderm and that stage-10
ectoderm is no longer competent to respond to this
signal under the conditions of this graft.
Table 2B shows the results of a similar series of
experiments in which the stage of competent donor
ectoderm was kept relatively constant, but the graft
Table 2. The differentiation of ectoderm in vegetal pole grafts
Donor stage
Host stage
Total number
of embryos
(A) The effect of varying donor stage
6
7
9
10
10-5
11
12
10
10
9
9
9
9
9
1
2
3
6
4
6
4
(B) The effect of varying host stage
9
9
9
7
8
9
7-5
8
9
10
10-5
11
2
2
2
2
2
2
No with grafted cells
positive for epidermal marker
No. with grafted cells
positive for muscle marker
Development of animal cap cells in Xenopus
29
Fig. 4. Determination of the ectoderm tested by grafting ectoderm into the vegetal pole of host blastulae; epidermal
differentiation. Small pieces of donor X. laevis ectoderm were inserted into the vegetal pole region of host X. borealis
embryos and allowed to develop until stage 25. Grafted embryos were fixed, embedded and stained as described before
with 2F7.C7, rhodamine RAM IgG and quinacrine. A and B show the expression of the epidermal marker on cryostat
sections of grafted cells on the archenteron wall following grafting from stage-9 and -10 donors, respectively, into
stage-9 hosts. C and D show quinacrine staining of the same sections, identifying the graft, in this case, by the lack of
X. borealis-speafic granules in the nuclei. Abbreviations as in Fig. 1. Bar: (A,B) 180jxm; (C,D) 15jxm.
Fig. 5. Determination of the ectoderm tested by grafting
into the vegetal pole of host blastulae. A graft from a
stage-9 X. borealis animal cap was made into the vegetal
pole of a stage-9 X. laevis host embryo. The grafted
embryo was incubated until control embryos reached
stage 30, fixed, embedded and stained with the muscle
marker 5A3.B4. Characteristic striated muscle is seen in
most of the graft. Abbreviations: e, endoderm; ism,
induced somitic muscle.
was made into host embryos varying from stage 7 to
stage 11. All grafts carried out into host embryos from
stages 7 to 10i expressed the muscle marker and did
not express the epidermal antigen. Those grafts
carried out into stage-11 host embryos developed into
balls of epidermis suggesting that the inductive stimulus is no longer present in the vegetal pole of stage-11
embryos. This shows that epidermal development, as
defined by 2F7.C7 binding, can proceed in a wholly
inappropriate environment.
These experiments show that ectoderm grafted into
the vegetal pole region of host embryos cannot
migrate from this position, but differentiates within
the endoderm. The final differentiated cell type
depends on the stage of development of both host and
donor tissue. If the ectoderm is still responsive to
mesodermal induction and the endoderm still capable
of producing the signal, then the graft differentiates
as mesoderm and often quite a large proportion of the
graft can be identified as apparently normal striated
muscle. If either of these conditions is not fulfilled
then the grafted cells differentiate into epidermis.
30
E. A. Jones and H. R. Woodland
Discussion
Migratory abilities of embryonic ectodermal cells
By placing the cells of the future ectoderm into the
blastocoel of another embryo, we have been able to
test their migratory behaviour in later development.
The results are clearest where the timing is arranged
so that the animal cap forms only ectoderm, rather
than mesoderm. This is achieved by grafting animal
cap cells of any stages into an embryo at stage 10 or
later, when mesodermal induction does not ensue
(Table 1). Alternatively, unresponsive ectoderm
from stage 10 or later may be placed in a blastocoel
at any stage (Table 1). By the tailbud tadpole stage,
the grafted cells are to be found in the epidermis, or
to a lesser extent the nervous system, of the host.
Moreover, transplanted cells from the inner or outer
layers of the animal cap are predominantly found,
respectively, in the inner sensorial layer or in the
outer, epidermal layer, just as they are in normal
development. The appearance of a minority of inner
cells in the epidermal layer is consistent with the view
that the scattered ciliated cells of the epidermis
originate in the inner layer (Billet & Courtenay, 1973;
Steinman, 1968; our unpublished observations).
Thus ectodermal cells do find their way to their
correct location in the embryo. Indeed, after grafting
ectoderm into the blastocoel we never see cells
expressing the epidermal marker except in the epidermis. Since epidermis can differentiate in the endoderm (see below), it seems that cells following this
epidermal pathway find their correct location from
the blastocoel. In addition, we do not see negative
cells inside the sensorial layer.
These results contrast with the failure of the grafted
animal cells to move when surrounded by animal
tissue as in a Holtfreter sandwich. In contrast, an
explant that contains mesoderm shows proper organization of the ectoderm (data not shown). This
suggests that the normal inner components of the
embryo provide something necessary for the migration of the cells. This could be extracellular
matrix, positional cues as to the location of the graft,
or a disaggregating environment, or a combination of
these factors.
What is the role of this migration in normal
development? The progeny of lineage-labelled animal cells show very considerable scattering after
gastrulation (Moody, 1987; Dale & Slack, 1987). This
shows that the cells are naturally very mobile within
their germ layer. Our results suggest that the integrity
of the layer is also actively maintained, to the extent
that a cell which becomes displaced as far as the
blastocoel can still regain its appropriate location.
Somewhat similar migratory abilities of the ectoderm
were demonstrated by Boucaut in Pleurodeles waltl
(Boucaut, 1974a,b). They are also an intrinsic part of
the single-cell transfers of late-stage animal cap cells
of Heasman et al. (1984), although in these experiments the differentiated state of the cells was not
always tested with cell-type markers.
Migration of mesodermal cells
Boucaut (1974/?) came to the conclusion that disaggregated mesodermal cells when placed into the
blastocoel of a recipient embryo had considerable
ability to organize themselves correctly within the
mesoderm when injected into the blastocoel. Our
experiments would suggest that the same is true in
Xenopus. When fragments of animal caps are introduced into blastocoels under timing regimes where
they can and are induced to form mesoderm, we find
that induced grafted cells are appropriately organized
into mesodermal tissues, although their fully differentiated state can only be positively identified when the
grafts are incorporated into somites and the musclespecific monoclonal 5A3.B4 can be used. This situation mainly occurs in embryos displaying secondary
embryonic axes (61 % of grafts in inductive combinations) when grafted cells are much more strongly
represented in dorsal mesoderm than in normal
grafted embryos when the majority of grafted cells
are in lateral plate and ventral mesoderm. A possible
interpretation of grafted embryos with secondary
embryonic axes might lie in a reduced migratory
potential of dorsal mesoderm. If this were so, dorsal
mesoderm formed would not move to the primary
dorsal region, but instead subverts gastrulation movements and organizes a second embryonic axis from
surrounding tissue. In contrast, ectodermal cells are
both relatively inert at blastula and gastrula stages
and more mobile. They might, consequently, have
longer in which to reach their appropriate positions
before they would upset development. However,
since primary and secondary axes are properly organized, and since mesoderm can differentiate in an
unusual environment (see below), mesodermal cells
can certainly organize themselves in the short range.
The role of the environment in epidermal and
mesodermal differentiation
When animal cap cells are placed in the vegetal pole
under circumstances where they are unresponsive to
mesodermal induction (post stage 104) or the host has
caused mesodermal induction (post stage 10£), they
invariably form epidermis in the walls of the gut or
within its tissue. This indicates first that neural
inductive stimuli do not occur here and, second, that
once either the inductive stimulus or the competence
to respond to mesodermal induction is lost, development into epidermis proceeds in a way that is not at
all upset by the bizarre environment.
Development of animal cap cells in Xenopus
When mesodermal induction can occur, striated
muscle is always formed, even though this is not
normally found in the gut. Moreover, in normal
embryos, this cell interaction occurs with vegetal cells
in an entirely different location, that is at the dorsal
margin between vegetal and animal cells. Our results
indicate that dorsal inductive stimuli are present
generally through the vegetal mass, even at the
extreme vegetal pole, and that once the stimulus to
form muscle has occurred, the cells differentiate
without reference to their environment. This correlates with the fact that blastula cells can differentiate
into muscle when disaggregated (Gurdon, Brennan,
Fairmans & Mohun, 1984; Sargent, Jamrich &
Dawid, 1986). It also fits with the observation that
dorsal mesoderm can change the fate of more vegetal
regions, but is not itself influenced (Slack & Forman,
1980). All of these observations support the idea that
a certain number of major steps in commitment can
be made in early development and that for these
subsequent reference to the environment is not made.
This work was funded by the Medical Research Council.
The authors acknowledge the clerical assistance of Mrs Len
Schofield and the technical assistance of P. Day.
References
M. & GRUNZ, H. (1983). Effects of inducers
on inner and outer gastrula ectoderm layers of Xenopus
laevis. Differentiation 23, 206-212.
BILLET, F. S. & COURTENAY, T. H. (1973). A stereoscan
study of the origin of ciliated cells in the embryonic
epidermis of Arnbystoma mexicanum. J. Embryol. exp.
Morph. 29, 549-558.
BOUCAUT, J.-C. (1974a). Etude autoradiographique de la
distribution de cellules embryonnaires isol6es,
transplanters dans le blastocele chez Pleurodeles waltlii
Micah (Amphibien, Urodele). Annls Embryol. Morph.
7, 7-50.
BOUCAUT, J. C. (19746). Chimeres intergeneriques entre
Pleurodeles waltlii Micah et Ambystoma mexicanum
Shaw (Amphibiens, Urodeles). Annls Embryol. Morph.
7, 119-139.
ASASHIMA,
COOKE, J. & WEBBER, J. A. (1985). Dynamics of the
control of body pattern in Xenopus laevis. I. Timing
and pattern in the development of dorsoanterior and of
posterior blastomere pairs isolated at the 4-cell stage. J.
Embryol. exp. Morph. 88, 85-112.
DALE, L. & SLACK, J. M. W. (1987). Fate map for the 32cell stage of Xenopus laevis. Development 100, 279-2%.
DALE, L., SMITH, J. C. & SLACK, J. M. W. (1985).
Mesoderm induction in Xenopus laevis; a quantitative
study using cell lineage label and tissue specific
antibodies. J. Embryol. exp. Morph. 89, 289-313.
GARDNER, R. L. (1985). Clonal analysis of early
mammalian development. Phil. Trans. R. Soc. Lond. B
313, 163-178.
31
GURDON, J. B. (1977). Methods for nuclear
transplantation in amphibia. Methods Cell Biol. 16,
125-139.
GURDON, J. B., BRENNAN, S., FAIRMANS, S. & MOHUN, T.
J. (1984). Transcription of muscle-specific actin genes
in early Xenopus development; nuclear transplantation
and cell dissociation. Cell 38, 691-700.
HEASMAN, J., WYLIE, C. C , HAUSEN, P. & SMITH, J. C.
(1984). Fates and states of determination of single
vegetal pole blastomers of Xenopus laevis. Cell 37,
185-194.
HOLTFRETER, J. & HAMBURGER, V. (1955). In Analysis of
Development (ed. B. H. Willier, P. A. Weiss & V.
Hamburger), pp. 230-296. New York: Saunders.
JONES, E. A. (1985). Epidermal development in Xenopus
laevis: the definition of a monoclonal antibody to an
epidermal marker. J. Embryol. exp. Morph. 89
Supplement, 155-166.
JONES, E. A. & WOODLAND, H. R. (1986). Development
of the ectoderm in Xenopus: tissue specification and the
role of cell association and division. Cell 44, 345-355.
KELLER, R. E. (1975). Vital dye mapping of the gastrula
and neurula of Xenopus laevis. I. Prospective areas and
morphogenetic movements of the superficial layer. Devi
Biol. 42, 222-241.
KINTER, C. R. & BROCKES, J. P. (1984). Monoclonal
antibodies identify blastemal cells derived from
differentiating muscle in newt limb regeneration.
Nature, Lond. 308, 67-69.
MOODY, S. A. (1987). Fates of the blastomers of the 16cell stage Xenopus embryo. Devi Biol. 119, 560-578.
SARGENT, T. D., JAMRICH, M. & DAWID, I. B. (1986).
Cell interactions and the control of gene activity during
early development of Xenopus laevis. Devi Biol. 114,
238-246.
SLACK, J. M. W. (1983). From Egg to Embryo.
Determinative Events in Early Development. Cambridge,
London: Cambridge University Press.
SLACK, J. M. W. (1984). In vitro development of isolated
ectoderm from axolotl gastrulae. J. Embryol. exp.
Morph. 80, 321-330.
SLACK, J. M. W. & FORMAN, D. (1980). An interaction
between dorsal and ventral regions of the marginal
zone in early amphibian embryos. J. Embryol. exp.
Morph. 56, 283-289.
SNAPE, A., WYLIE, C. C , SMITH, J. C. & HEASMAN, J.
(1987). Changes in states of commitment of single
animal pole blastomeres of Xenopus laevis. Devi Biol.
119, 503-510.
SPEMANN, H. & MANGOLD, H. (1924). Uber Induktion
von Embryonalanlagen durch Implantation artfremder
Organisatoren. Arch. Mikrosk. Anat. EntwMech. Org.
100, 599-638.
STEINMAN, R. M. (1968). An electron microscopic study
of ciliogenesis in the developing epidermis and trachea
in the embryo of Xenopus laevis. Am. J. Anat. 122,
19-56.
32
£. A. Jones and H. R. Woodland
S. & NIEUWKOOP, P. D. (1971). Mesoderm
formation in the anuran Xenopus laevis (Daudin).
Wilhelm Roux Arch. EntwMech. Org,166, 189-204.
THIEBAUD, C. H. (1983). A reliable new cell marker in
Xenopus. Devi Biol. 98, 245-249.
WYLIE, C. C , SMITH, J. C , SNAPE, A. & HEASMAN, J.
(1985). The use of single cell transplantation in the
study of cell commitment in early Xenopus embryos. In
SUDARWATI,
44th Annual Symposium of the Society of Developmental
Biology. Gametogenesis and the Early Embryo. New
York: Alan R. Liss.
WYLIE, C. C , SNAPE, A., HEASMAN, J. & SMITH, J. C.
(1987). Vegetal pole cells and commitment to form
endoderm in Xenopus laevis. Devi Biol. 119, 496-502.
(Accepted 22 May 1987)