Download Hensen`s node induces neural tissue in Xenopus

Development 113, 1495-1505 (1991)
Printed in Great Britain © The Company of Biologists Limited 1991
1495
Hensen's node induces neural tissue in Xenopus ectoderm. Implications for
the action of the organizer in neural induction
C. R. KINTNER1 and J. DODD 2
^Molecular Neurobiology Laboratory, The Salk Institute, San Diego, CA 92186, USA
Department of Physiology and Cellular Biophysics, Center for Neurobiology and Behavior, Columbia University, New York, NY 10032,
USA
2
Summary
The development of the vertebrate nervous system is
initiated in amphibia by inductive interactions between
ectoderm and a region of the embryo called the
organizer. The organizer tissue in the dorsal lip of the
blastopore of Xenopus and Hensen's node in chick
embryos have similar neural inducing properties when
transplanted into ectopic sites in then" respective embryos. To begin to determine the nature of the inducing
signals of the organizer and whether they are conserved
across species we have examined the ability of Hensen's
node to induce neural tissue in Xenopus ectoderm. We
show that Hensen's node induces large amounts of
neural tissue in Xenopus ectoderm. Neural induction
proceeds in the absence of mesodermal differentiation
and is accompanied by tissue movements which may
reflect notoplate induction. The competence of the
ectoderm to respond to Hensen's node extends much
later in development than that to activin-A or to
induction by vegetal cells, and parallels the extended
competence to neural induction by axial mesoderm. The
actions of activin-A and Hensen's node are further
distinguished by their effects on lithium-treated ectoderm. These results suggest that neural induction can
occur efficiently in response to inducing signals from
organizer tissue arrested at a stage prior to gastrulation,
and that such early interactions in the blastula may be an
important component of neural induction in vertebrate
embryos.
Introduction
tissue when transplanted to ectopic sites in chick or
amphibian embryos, respectively. In addition, both
tissues pattern the induced neural tissue along the
anterior-posterior axis (Spemann, 1931; Tsung et al.,
1965; see Hara, 1978). The similarities in the properties
of the organizer in different vertebrate embryos
suggests that the inducing signals underlying neural
induction may be conserved. To examine this idea, we
have asked whether Xenopus ectoderm responds to the
organizer taken from the chick embryo as it does to its
own organizer. Chick Hensen's node was isolated and
cultured with Xenopus ectoderm in a standard animal
cap assay. The assay was carried out at room
temperature (approximately 22°C) such that the ectoderm can respond to inducing signals while the chick
tissue is developmentally static. Thus tissue differentiation and morphogenetic movements observed represent the response of Xenopus ectoderm, while the
inducing signals detected in the assay can be attributed
to Hensen's node tissue as it existed at the stage of
isolation from the embryo.
We show that Xenopus ectoderm responds to
Hensen's node by forming large amounts of neural
tissue without detectably forming dorsal mesodermal
Vertebrate neural development begins when ectoderm
on the dorsal side of the embryo is induced to form
neural tissue. The neural inducer is a region of tissue,
known as the organizer, whose properties have been
defined by transplantation studies in several vertebrate
species. When the organizer is transplanted from one
blastula to the ventral side of another, the host embryo
forms a second dorsal axis (Spemann and Mangold,
1924). The nervous system in this second axis is derived
from host ectoderm which, in the absence of the graft,
would have formed ventral epidermis. This result
indicates that ectoderm can be induced to form neural
tissue upon appropriate interaction with the organizer,
an event that normally occurs only on the dorsal side of
the embryo. In amphibia, the organizer maps to the
dorsal lip of the blastopore (DLB), a region of the
marginal zone of the blastula just above the first site of
invagination at gastrulation. In birds, the organizer
maps to the anterior end of the primitive streak, a
region called Hensen's node (Waddington and Schmidt,
1933; see Hara, 1978).
Both Hensen's node and the DLB induce neural
Key words: neural induction, organizer, activin-A,
Hensen's node, notoplate.
1496
C. R. Kintner and J. Dodd
derivatives. The inducing activity of Hensen's node is
different from the potent embryonic inducer, activin-A
(Asashima et al., 1990; Eijnden-Van Raaij et al., 1990;
Smith et al., 1990; Thompsen et al., 1990). In addition,
Hensen's node induces ectoderm to undergo morphological movements resulting in axial elongation in the
absence of mesoderm. These results are discussed in the
light of the model in which an early interaction between
ectoderm and organizer tissue in the blastula is an
important component of neural induction in vertebrate
embryos.
Methods
Animals and reagents
Embryos were obtained from Xenopus laevis adult frogs
(NASCO and Xenopus 1) by hormone-induced egg-laying
and in vitro fertilization using standard protocols. Animal
caps were dissected from appropriately staged (Nieuwkoop
and Faber, 1967) embryos in 0.5x MMR containing penicillin
and streptomycin, as previously described (Dixon and
Kintner, 1989). For lithium treatment, embryos were incubated at the 64 cell stage with 0.25 M LiCl for 10 minutes and
then washed extensively with 0.5 x MMR. To measure
competence, ectodermal caps were isolated from embryos at
appropriate times after the start of gastrulation by removing
ectoderm that was not yet contacted by involuting tissue.
Hensen's node tissue was dissected from chick embryos
(White Leghorns from Spafas, CT, Truslow Farms, VA, and
Mclntyre Poultry, CA) at stages 3-10 (Hamburger and
Hamilton, 1951) in ice cold L15 medium. The chick tissue was
placed between two pieces of animal cap tissue as shown in
Fig. 1, and the resulting recombinants maintained on agarosecoated dishes in 0.5 x MMR with penicillin and streptomycin
at room temperature (approximately 22°C). Recombinants
were harvested for RNA analysis after 20 hours or were
maintained for 2.5 days with frequent changes of the culture
medium for observation and histological analysis.
A highly purified preparation of porcine activin-A was
kindly provided by Drs W. Vale and J. Vaughan in the Peptide
Biology Laboratory at the Salk Institute for Biological
Studies.
RNA analysis
RNA was isolated from embryos or explants and assayed for
the expression of specific RNA transcripts using an RNAase
protection assay described previously (Melton et al., 1984;
Kintner and Melton, 1987). The hybridization probes used to
detect NCAM, NF3, muscle-specific actin and EF-1 alpha
RNA are described elsewhere (Kintner and Melton, 1987;
Dixon and Kintner, 1989). The hybridization probe for
XlHbox6 RNA is a fragment of the XlHbox6 cDNA (Sharpe
et al., 1987) kindly provided by Drs C. Wright and E.
DeRobertis. The hybridization probe for en-2 RNA is a
portion of the Xenopus engrailed-2 cDNA that was isolated
and provided for us by Drs R. Harland and A. HemmatiBrivanlou (Hemmati-Brivanlou et al., 1991). The amounts of
RNA in each sample were normalized by monitoring the
levels of EF-1 alpha RNA (Krieg et al., 1989). RNA samples
prepared from 4 recombinants or embryos were assayed
simultaneously with all probes except for the muscle-specific
actin probe, in which case 10% of the RNA sample was
assayed separately. The Xenopus hybridization probes were
fully protected by transcripts isolated from frog tissue but
were not protected by transcripts isolated from chick
embryos. In every assay the size of undigested probes for each
transcript used was measured on the same gel as the
experimental samples. This permitted us to determine
whether probe escaped RNAase protection in any experimental samples.
Histology and immunohistochemistry
Recombinants were fixed in 3% trichloracetic acid at 4°C for
30 min. After extensive washing, they were dehydrated
through an ethanol series and embedded in paraffin using
standard procedures (Kintner, 1988). Tissue was sectioned at
10 fsm on a rotary microtome and mounted on gelatin-subbed
slides. For histology, sections were stained with haematoxylin
and eosin or Giemsa's stain. For immunohistochemistry, the
sections were dewaxed, labelled with monoclonal antibodies,
using a fluorescein-labelled second antibody and counterstained with Hoechst dye. Monoclonal antibodies used were
as follows: anti-NCAM (gift of Drs K. Sakaguchi and W.
Harris, UCSD), 12/101 (Kintner and Brockes, 1985) and Not1 (Placzek et al., 1990; Yamada et al., 1991).
Results
Hensen's node induces neural tissue
Fate mapping and transplantation studies have led to
the idea that Hensen's node in the chick is functionally
equivalent to the dorsal lip of the blastopore (DLB) in
amphibia (see Hara, 1978). We therefore examined
whether Hensen's node could substitute for DLB tissue
in a Xenopus animal cap assay by combining Hensen's
nodes, dissected from stage 3.5 and stage 4 chick
embryos, with pairs of ectodermal caps dissected from
stage 9 Xenopus laevis embryos (Fig. 1). The recombinants were allowed to develop for 18-24 hours at room
temperature and then assayed for the expression of
tissue-specific RNA transcripts.
The recombinants were examined for the presence of
neural tissue by measuring the expression of two
neural-specific transcripts, NCAM and NF-3. NCAM is
expressed in Xenopus ectoderm soon after neural
induction and is restricted to neural tissue (Jacobson
and Rutishauser, 1986; Kintner and Melton, 1987)
providing an early general marker of neural differentiation. NP-3 encodes a neuronal intermediate filament
protein (Charnas et al., 1987) that is expressed in
neurons beginning about 4 hours after neural tube
closure (Dixon and Kintner, 1989). All transcripts were
detected with an RNAase protection assay in which the
Xenopus tissue but not the chick tissue generated fulllength protected fragments.
Recombinants formed between Xenopus ectoderm
and Hensen's node expressed both NCAM and NF-3
transcripts (Fig. 2, lane 4). The levels of the two
transcripts expressed in the ectodermal caps after
induction by Hensen's node were approximately equivalent to the levels observed in stage-matched control
embryos normalized for the expression of EF-1 alpha
RNA (Fig. 2, lane 6). Control explants of ectoderm
cultured alone expressed neither NCAM nor NF-3 (Fig.
2, lane 2). Of the tissue in the chick blastula, only
Hensen's node was able to induce large amounts of
neural tissue in Xenopus ectoderm, although very low
Neural induction in Xenopus laevis 1497
ectoderm
Chicken
embryo
levels of NCAM RNA were induced by primitive streak
immediately posterior to the node (Fig. 2, lane 5).
Equal-sized pieces of tissue dissected from lateral or
anterior epiblast (see Fig. 1) or caudal primitive streak
did not induce neural transcripts in isolated frog
ectoderm (Fig. 2, lane 3). Thus, neural inducing
capacity was restricted to the region of the chick that is
thought to be equivalent to the amphibian DLB (Hara,
1978).
Hensen's node does not appear to differentiate within
the recombinates
An important aspect of these experiments is that the
recombinants were maintained at room temperature
(approximately 22°C) in 0.5x MMR, conditions in
which development of Hensen's node should be
arrested. The inducing signals detected in this assay are
therefore likely to be present in the node at the time of
explantation and unlikely to be generated by tissues
arising from the node during the course of the assay. To
determine whether Hensen's node arrests within the
recombinant, or alone at 22°C in 0.5x MMR, we
examined sections of the node for the expression of the
differentiation antigen, Not-1. Not-1 is expressed by
chick notochord, beginning at stage 5, as Hensen's node
begins to regress and notochord is laid down (Placzek et
al., 1990; Yamada et al., 1991). Hensen's node
incubated in the presence or absence of ectodermal
caps in 0.5 x MMR overnight at room temperature did
not express Not-1. The antigen was expressed by stage 5
or 6 chick notochord before and after incubation of the
tissue in 0.5x MMR overnight, indicating that the
antigen itself is not destroyed by incubation under these
conditions (not shown). Not-1 was expressed in stage 4
Hensen's node that had been incubated at 37°C in
isotonic medium overnight. These results suggest that
Hensen's node is arrested developmentally at room
temperature and that its effects on Xenopus ectodermal
tissue do not depend on differentiation into its
mesodermal derivatives.
The inducing properties of Hensen's node change with
developmental stage
Transplantation of the amphibian DLB taken from
different developmental stages results in the induction
Fig. 1. Pieces of chick epiblast
with underlying hypoblast were
dissected as shown from
Hensen's node (HN), the
primitive streak (PS) and other
regions of the embryo (e).
These were sandwiched
between two animal
ectodermal caps dissected from
stage 9-10 Xenopus blastulae.
Xenopus
blastula
chick tissue and
frog ectoderm
UNPROTECTED
PROBES
muscle
actin
8 « 1
a
O
2
Q
z
f)
Q.
^
8
PROTECTED
PROBES
w
to
muscle
actin
I 8
NF-3
NF-3
NCAM
NCAM
EF-1A
EF-1A
1 2
3
4
6 7
Fig. 2. RNA transcripts expressed in Xenopus ectoderm in
response to Hensen's node. Total RNA samples prepared
from 4 recombinants or control tissues were assayed
simultaneously for the presence of NF-3, NCAM and EF1alpha transcripts by RNAase protection. 10% of each
sample was assayed separately for the presence of musclespecific actin transcripts. Lane 1 shows the position of the
unprotected probes, indicated by arrows. Arrows on the
extreme right show the position of the probes after
protection with the appropriate RNA transcripts. Ectoderm
cultured with stage 4 Hensen's node expresses large
amounts of NCAM and NF-3 transcripts (lane 4) at levels
found in stage-matched control embryos (lane 6).
Ectoderm cultured with anterior primitive streak tissue
expressed NCAM RNA at just detectable levels (lane 5).
Note that ectoderm cultured alone (lane 2) or combined
with chick epiblast (lane 3) expresses EFl-alpha but not
NCAM RNA. None of the probes are protected by tRNA
(lane 7).
1498 C. R. Kintner and J. Dodd
of tissues with different regional characteristics (Spemann, 1931). Dorsal lip tissue taken from an early
gastrula stage induces anterior structures while dorsal
lip taken from later-staged embryos induces posterior
tissues such as spinal cord. A similar phenomenon has
been observed in the chick embryo in that secondary
neural tubes induced by a subepiblast transplant of
Hensen's node became progressively more posterior in
character as donor nodes were taken from later staged
embryos (Tsung et al., 1965; but see Dias and
Schoenwolf, 1990). These observations suggest that the
inducing properties of Hensen's node and the DLB
change similarly with developmental age. We therefore
tested the ability of Hensen's nodes, dissected from
chick over a range of ages, to induce neural tissues of
anterior and posterior character in Xenopus ectoderm.
To do this we measured the expression of two
homeobox-containing genes, en-2 and XlHbox6, that
are expressed at different levels of the A-P axis in
Xenopus ectoderm. En-2 is expressed in the neural
plate at the level of the midbrain (Hemmati-Brivanlou
and Harland, 1989; Hemmati-Brivanlou et al., 1991).
XlHbox6 transcripts are expressed in the posterior
region of the neural plate (Sharpe et al., 1987) and in
lateral plate mesoderm (Wright et al., 1990).
In animal caps induced with stage 4 Hensen's nodes
the en-2 transcript was observed at levels comparable to
those observed in stage-matched control embryos (Fig.
3, lanes 1,4). However, when Hensen's nodes from late
stage 5 to stage 9 were used, the expression of en-2
ceased (Fig. 3, lanes 2,3). Conversely, stage 4 Hensen's
node did not induce expression of XlHbox6 (Fig. 3, lane
1), whereas later-staged Hensen's nodes did induce
XlHbox6 (Fig. 3, lanes 2,3).
The results described above indicate that Hensen's
node tissue mimics at least two characteristics of tissue
in the DLB. First, Hensen's node induces large
amounts of neural tissue in Xenopus ectoderm. This
effect is observed with both stage 3 and stage 4
Hensen's nodes, indicating that Hensen's node contains
neural inducing signals before gastrulation. Second, the
neural tissue induced has different regional characteristics that depend on the age of the embryos from which
Hensen's node is isolated. These results suggest that
Hensen's node can be used as a model to study the
properties of the signals that are present in the DLB.
We have therefore analysed the response of ectoderm
to Hensen's node and compared the responses of
Xenopus ectoderm to Hensen's node and another
potent embryonic inducer, activin-A.
Hensen's node does not induce axial mesoderm
Ectoderm treated with embryonic inducers such as
activin-A and FGF forms mesodermal derivatives such
as notochord and muscle (Slack et al., 1987; Smith,
1987). In order to determine whether ectoderm also
responds to Hensen's node by forming axial mesoderm,
we examined recombinants for the expression of
somitic mesoderm using two assays. We first measured
the expression of muscle-specific actin RNA in an
RNAase protection assay. Hensen's node did not
HENSEN'S
NODE
c
in
o
CO
CO
CO c/D
8
x
en-2
r
NCAM
EF-1A
•HI?
XIHBOX 6
1
2
3
4 5
Fig. 3. Expression of RNA transcripts in Xenopus
ectoderm in response to Hensen's node from embryos at
different developmental stages. Hensen's node was isolated
from chick embryos of different stages and combined with
Xenopus ectoderm from stage 10 embryos. After 24 hours
in culture, total RNA samples were prepared and assayed
simultaneously for en-2, NCAM, XlHbox6 and EFl-alpha
RNA, using RNAase protection. The position of the
protected probe for each transcript is indicated by arrows
on the left. Stage 30 embryos expressed all four transcripts
(lane 4) while ectoderm cultured alone expressed only EF1 alpha RNA (lane 5). Ectoderm combined with stage 4
Hensen's node expressed NCAM and en-2 but not
XlHbox6 transcripts (lane 1). Ectoderm combined with late
stage 5 (lane 2) or later (lane 3) expressed NCAM and
XlHbox6 but not en-2 RNA.
induce the expression of the muscle-specific actin
transcript (Fig. 2, lane 4), indicating that the responding ectoderm did not form somitic mesoderm. In
contrast, whenever neural tissue was induced by
activin-A, it was accompanied by the induction of large
amounts of somitic mesoderm (Fig. 5, lanes 2,3). As an
alternative assay for somitic muscle, recombinants were
examined for the presence of the muscle specific
antigen, 12/101, by immunohistochemical labelling of
tissue sections. While patches of muscle were brilliantly
labelled with this antibody in control explants contain-
Neural induction in Xenopus laevis 1499
Fig. 4. Histological analysis of recombinants. (A,B) Haematoxylin and eosin-stained paraffin sections of recombinants
formed between, Xenopus ectoderm and Xenopus organizer (A) and Xenopus ectoderm and chick Hensen's node (B).
Notochord cells (n) are present in A but not in B. (C) Immunofluorescence micrograph showing labelling of neural tissue
with anti-NCAM antibodies. The neural tissue (large arrowhead) is found adjacent to the chick tissue (small arrowheads)
which can be identified by Hoechst staining in serial sections (D) (small arrowheads).
ing Xenopus DLB tissue, no labelling with the 12/101
antibody was observed in recombinants between
ectoderm and Hensen's node (not shown).
Although muscle-specific actin and 12/101-reactive
tissue were not induced by Hensen's node, it is possible
that notochord tissue was induced in the absence of
somitic mesoderm. To determine whether chordamesoderm was present we examined sections of recombinants for the presence of notochord and other
mesodermal derivatives. Notochordal tissue, recognizable by the presence of large vacuolated cells in
recombinants containing DLB tissue (Fig. 4A), could
not be detected in recombinates containing Hensen's
node (Fig. 4B). The lack of detectable mesoderm was
independent of the stage of embryo (3-10) from which
Hensen's node was taken (not shown). While this
analysis does not exclude the presence of a few isolated
notochordal cells, it provides strong evidence that the
formation of axial mesoderm is not a major response of
Xenopus ectoderm to stage 4 Hensen's node.
Recombinants containing stage 4 Hensen's node
were also examined immunohistologically for the
formation of neural tissue, using a monoclonal antibody
against NCAM. Recombinants contained large
amounts of NCAM-immunoreactive tissue (Fig. 4C),
confirming the results obtained measuring neural
transcripts. NCAM labelling was primarily associated
with tissue in direct contact with Hensen's node (Fig.
4C,D), suggesting that the ectoderm formed neural
tissue in the absence of intervening cells or tissues.
Together, these results indicate that Hensen's node
can induce large amounts of neural tissue to form from
Xenopus ectoderm and that this induction occurs in the
absence of axial mesodermal derivatives such as
notochord and somitic muscle. In addition, since
Hensen's node induces neural tissue in the absence of
axial mesoderm, its properties appear to be different
from those of factors identified as embryonic mesodermal inducers, such as activin-A and FGF.
Hensen's node inducing activity is distinct from
activin-A activity
Several studies indicate that different parameters can
alter the responsiveness of ectoderm to induction by
polypeptide growth factors. In the case of activin-A, the
ectodermal response can change as a function of the age
of the embryo from which the ectoderm is isolated and
by treatment of embryos with a dorsalizing agent,
lithium. To characterize the properties of the inducing
signals in Hensen's node further, we examined the
effect of these parameters on responses of ectodermal
caps to Hensen's node.
1500 C. R. Kintner and J. Dodd
(i) Effects of ectodermal competence on the
response to Hensen's node
The ability of the ectoderm to respond to mesoderminducing signals is known to decline at the beginning of
gastrulation and to be lost by midgastrula stages. We
therefore tested the response of ectoderm, taken from
progressively older Xenopus embryos, to activin-A and
Hensen's node. In accordance with the results of others
(Green et al., 1990), ectoderm taken from embryos
before gastrulation (stages 8.5-10) responded to activinA (0.07 nM) by expressing both NCAM and musclespecific actin transcripts (Fig. 5A, lanes 1,2). No
response was observed at 7 pM activin-A (not shown).
By midgastrulation (stage 11), ectoderm no longer
expressed NCAM RNA in response to activin-A (Fig.
5A, lanes 3,4) but continued to express muscle-specific
actin in response to high concentrations of activin-A
(1.4 nM) (Fig. 5A, lane 4). Ectoderm taken from
embryos older than stage 11 did not express NCAM or
muscle-specific actin in response to activin-A (not
shown). Surprisingly, ectoderm continued to respond to
activin-A by expressing XlHbox6 even when its ability
to form neural tissue was lost. The XlHbox6 signal in
ectoderm treated with activin-A after stage 11 may
reflect the induction of posterior lateral mesoderm that
is known to express XlHbox6 in embryos (Wright et al.,
1990).
B
k
The response of ectoderm to Hensen's node also
declined with developmental age, but the decline
occurred at later stages than that observed with activinA. Ectoderm continued to respond to stage 4 Hensen's
node by expressing large amounts of NCAM RNA until
late gastrula stages (Fig. 5B, lanes 1-3), considerably
later than the developmental stages at which activin-A
is still able to induce NCAM expression. Moreover,
even when the ability of the ectoderm to respond to
Hensen's node declined, Hensen's node did not induce
the expression of muscle-specific actin (Fig. 5B, lane 3).
Thus the competence of ectoderm to form neural
tissue in response to Hensen's node persisted for a
longer period after gastrulation than that to activin-A.
At stage 11, the ectoderm has lost competence to form
neural tissue in response to activin-A despite the fact
that muscle-specific actin RNA was expressed. In
contrast, Hensen's node continues to induce neural
tissue at late stages of gastrulation.
(ii) Effects of lithium on the responses of ectoderm
to Hensen's node
The response to activin-A is enhanced in ectoderm
taken from embryos treated with lithium at the 64 cell
stage. The ratio of dorsal to ventral induced mesoderm
increases and the competence of the ectoderm to
respond is extended (Cooke et al., 1989; Kao and
Fig. 5. Effects of competence
on the ectodermal response to
Ect + Act (nM)
activin-A and stage 4 Hensen's
Ect + HN
node. Animal caps were
8
isolated from embryos at
O
i-r^
o
different ages (stages 9-11.5)
muscle
muscle
and exposed either to activinactin
actin
A (A) or to Hensen's node
(B). Total RNA samples were
prepared from 8 animal caps, 4
NCAM
NCAM
recombinants or 4 control
tissues, and assayed by
RNAase protection for the
expression of NCAM, en-2,
XlHbox6 and EFl-alpha RNA.
EF-1A
EF-1A
10% of each sample was
assayed separately for the
presence of muscle-specific
actin transcripts. The position
of the protected probe for
each transcript is shown to the
XIHBOX 6
XIHBOX 6
left of each panel. Stage 30
m
embryos (St 30 con) express all
en-2
en-2
*"
Av
-•-• ~~* ^m
"" ^* """'
^
five
transcripts (lane 5 in A;
1 2 3 4
5
1 2 3 4
' a n e 4 in B). (A) Ectoderm
from stage 10 embryos
expressed muscle actin,
NCAM and XlHbox6 RNA in response to both 0.07 nM (lane 1) and 1.4 nM (lane 2) activin-A. When ectoderm was
isolated from embryos at a later stage (stage 11, early gastrulation), the response to activin-A was greatly diminished.
Stage 11 ectoderm did not express NCAM RNA in response to 0.07 nM (lane 3) or 1.4 nM (lane 4) activin-A. Stage 11
ectoderm could be induced to express muscle-specific actin RNA but in response only to high concentrations of activin-A
(1.4 nM, lane 4). The low levels of en-2 signal in lane 4 represent undigested probe (see methods). (B) Recombinants of
Hensen's node and ectoderm from stage 10 (lane 1), stage 11 (lane 2) and stage 11.5 (lane 3) embryos express NCAM and
en-2 RNA but not muscle-specific actin or XlHbox6 RNA. The levels of NCAM and en-2 transcripts are approximately the
same in ectoderm from stage 10 and stage 11 embryos, but decrease in ectoderm from stage 11.5. Although the stage 11.5
recombinant has less NCAM RNA expression, it does not express muscle-specific actin or XlHbox6 RNA-
I
Neural induction in Xenopus laevis 1501
B
Fig. 6. Effect of lithium on the
response of ectoderm to
activin-A and Hensen's node.
Animal caps were isolated
from stage 9-11.5 lithiummuscle
muscle
8
8
treated or control embryos and
actin
actin
were treated with activin-A or
combined with stage 4
Hensen's nodes. Total RNA
NCAM
NCAM
samples were prepared from 8
animal caps, 4 recombinants or
4 control tissues and assayed
by RNAase protection for the
expression of NCAM, en-2,
XlHbox6 and EFl-alpha RNA.
EF-1A
EF-1A
10% of each sample was used
to assay for the muscle-specific
actin transcript. The position
of the protected probe for
each transcript is marked to
the left of each panel with
XIHBOX 6
XIHBOX 6
arrows. Stage 30 embryos
expressed all five transcripts
(lane 5 in A; lane 5 in B). (A)
en-2
__ „ _
Ectoderm from stage 11
embryos does not express
NCAM RNA in response to
t -a
3 4
activin-A and requires high
concentrations of activin-A (1.4 nM) to express muscle-specific actin and XlHbox6 transcripts (lane 2). The low levels of
en-2 in lane 2 represent unprotected probe. In contrast, stage 11 ectoderm from lithium-treated embryos responds to
activin-A at concentrations of 0.07 nM and 1.4 nM by expressing NCAM, muscle-specific actin and XlHbox6 transcripts.
Note that en-2 transcripts are not expressed in ectodenn treated with activin-A even when isolated from lithium-treated
embryos. (B) The response of stage 11 (lanes 1,2) and stage 11.5 ectoderm (lanes 3,4) to Hensen's node was the same in
animal caps isolated from lithium-treated (lanes 2,4) and untreated (lanes 1,3) embryos. The band in the muscle-specific
actin assay (lane 4) represents unprotected probe (see methods).
St11 Ect + Act (nM)
st 11 Ect st 11.5 Ect
+ HN
+HN
c
o
o
o
co
I
Elinson, 1989). We therefore compared the effects of
lithium treatment on the responses of ectodenn to
Hensen's node and activin-A. As described above,
ectoderm from control embryos older than stage 11
does not express NCAM in response to activin-A and
expresses muscle-specific actm transcripts only in
response to high concentrations of activin-A (1.4 nM)
(Fig. 6A, lanes 1,2). In marked contrast, lithiumtreated stage 11 ectoderm expressed both NCAM and
muscle-specific actin RNA (Fig. 6A, lanes 3,4) even
when exposed to low concentrations of activin-A (0.07
nM). Thus, as previously reported (Cooke et al., 1989),
lithium treatment potentiates the response of ectodenn
to activin-A.
The generation of neural tissue by ectoderm in
response to stage 4 Hensen's node was not altered by
lithium treatment (Fig. 6B, lanes 1-4). Both the levels
of NCAM transcripts and the competence of the
ectoderm to respond to Hensen's node were the same in
lithium-treated and control embryos. Because the
Hensen's node signal cannot be titred, we do not know
whether it was already optimized in these experiments.
Thus we cannot definitively conclude that lithium is not
capable of potentiating the response of early ectoderm
to Hensen's node. However, the results do show that
lithium treatment is not able to extend the competence
of older ectoderm to respond to Hensen's node (Fig.
6B, lanes 1-4), in contrast to its effect on the activin-A
response.
The effects of competence and lithium treatment on
the expression of region specific markers, en-2 and
XlHbox6 in Xenopus ectoderm in response to
Hensen's node
Stage 4 Hensen's node induces the expression of en-2
but not of XlHbox6 in stage 10 Xenopus ectoderm, as
described above. Conversely, activin-A induces
XlHbox6 but not en-2 in stage 10 ectoderm. The
differential expression of en-2 and XlHbox6 RNA in
response to activin-A and Hensen's node was unaffected by lithium treatment or by the age of the ectoderm.
As in control ectoderm, en-2 RNA was not induced by
activin-A in lithium-treated ectodenn (Fig. 6A, lanes 14) but was expressed at control levels in response to
Hensen's node (Fig. 6B, lanes 1-4). XlHbox6 RNA was
not expressed in response to stage 4 Hensen's node but
was expressed in ectoderm that responded to activin-A
(Fig. 6A, lanes 2-4).
Extension movements are induced by Hensen's node
In the course of these experiments, we consistently
observed that ectodenn underwent extensive morphological movements in response to Hensen's node. In
some cases these movements resulted in extreme axial
1502 C. R. Kintner and J. Dodd
A
B
neurulation (Fig. 7 B-D). Movements were not
observed in isolated Hensen's node placed alone in
0.5x MMR at room temperature (not shown) or in
isolated ectoderm from Xenopus embryos, indicating
that they were initiated in the ectoderm in response to
Hensen's node. It is unlikely that ectoderm induced the
movements in Hensen's node, since in sections of the
recombinants the chick cells appeared to have remained
in a local cluster. Elongation movements of the
magnitude observed in the Hensen's node recombinants are known to occur in Xenopus embryos only in
cells of the involuting marginal zone at the DLB as they
form the midline of the prospective dorsal mesoderm,
or in cells of the noninvoluting marginal zone as they
form the midline of the neural plate (Keller, 1985).
Since the recombinants do not form mesodermal
derivatives, the observed movements are likely to be
associated with the formation of neural tissue.
Discussion
Both Hensen's node of the chick and the DLB of
amphibia induce host ectoderm, in chick and Xenopus
respectively, to form neural tissue upon transplantation
to an ectopic site (Spemann and Mangold, 1924;
Waddington and Schmidt, 1933; Smith and Slack, 1983;
Vakaet, 1965; McCallion and Shinde, 1973; Hara, 1978;
Hornbruch et al., 1989; Dias and Schoenwolf, 1990).
The results reported here show that Xenopus ectoderm
also responds in vitro to Hensen's node. The ectoderm
forms neural tissue at levels similar to those formed in
normal embryos and expresses distinct regional neural
markers in response to nodes taken from embryos at
different developmental stages. The induction of neural
tissue by Hensen's node is not accompanied by the
formation of any detectable mesodermal derivatives
and occurs under conditions in which development of
the node itself is arrested. Thus organizer tissue appears
to be able to act as a potent neural inducer before it
gastrulates and forms mesodermal derivatives.
Fig. 7. Xenopus ectoderm/Hensen's node recombinants
show gastrulation-like movements. (A) Several
recombinants showing extreme elongation and
concentration of pigment are shown 18 hours after the
tissues were placed together. (B-D) A recombinant at 2
(B), 6 (C) and 18 (D) hours after sandwiching of the
Hensen's node with Xenopus ectoderm.
elongation of the recombinants (Fig. 7A), during which
the inner ectodermal cells were extruded. In other
cases, cell movements were extensive and resulted in a
complex pigmentation patterns, but seemed not to be
oriented and the explants did not elongate. The timing
of the movements occurred over the time course of axial
elongation in embryos undergoing gastrulation and
The organizer as an early inducer of neural tissue
The finding that avian organizer tissue is a potent
inducer before gastrulation and need not develop into
mesodermal derivatives to act as an effective neural
inducer suggests that neural induction in vivo may occur
in the blastula. This induction may be due to an
interaction between the organizer and ectoderm across
the boundary they share in the blastula and may be
carried by signals travelling in the plane of the
ectoderm. This implies that organizer tissue need not
involute beneath the ectoderm for neural induction to
occur. This idea has already been suggested from results
with Xenopus embryos (exogastrulae) and explants
(Keller sandwiches) in which DLB tissue does not
involute beneath the prospective neural ectoderm
during gastrulation. In this situation in Xenopus the
ectoderm in edgewise contact with the DLB tissue
Neural induction in Xenopus laevis 1503
forms large amounts of neural tissue (Kintner and
Melton, 1987; Keller and Danilchik, 1988; Dixon and
Kintner, 1989).
The idea of an early induction between the organizer
and ectoderm is also supported by the observation that
one of the first responses of ectoderm to Hensen's node
is elongation movements. Elongation movements are
known to occur in only two regions of the Xenopus
embryo. One region corresponds to the cells within the
involuting marginal zone, a region which converges,
involutes, comes to lie beneath the neural plate and
forms notochord. The other region maps to cells in the
noninvoluting marginal zone, called notoplate cells,
which extend within the ectodermal sheet along the
midline of the prospective neural plate, during gastrulation and neurulation (Jacobson and Gordon, 1976;
Gordon and Jacobson, 1978; Keller, 1985; Jessell et al.,
1989). The notoplate arises from the ectoderm that is
closest to the organizer in the blastula fate map and its
movements have been shown to occur in the absence of
underlying mesoderm and when dissected free of
prospective mesoderm after stage 11 (Keller and
Danilchik, 1988). This suggests that the notoplate is
specified by an interaction between the organizer and
adjacent ectoderm or by an inductive signal from
vegetal cells (Jacobson and Sater, 1988) prior to the
onset of gastrulation. Our results support this notion in
that Xenopus ectoderm responds to Hensen's node with
striking movements that can be observed within two
hours of contact. These movements cause the explants
to elongate over a time course similar to that of
gastrulation and neurulation in whole embryos (Symes
and Smith, 1987), and occur in the absence of
detectable mesoderm. The simplest explanation of
these movements is that they reflect the movements of
notoplate cells which have been induced in the
ectoderm by signals from organizer tissue.
Induction of the notoplate by an interaction between
the organizer and ectoderm before gastrulation may be
an important component of neural induction. As they
converge and extend along the anterior-posterior axis,
notoplate cells may be the source of neural inducing
signals that act on surrounding dorsal ectoderm to form
lateral regions of the neural plate. This model predicts
that notoplate extension is necessary for efficient neural
induction and that inducing signals travel from the
midline of the prospective neural plate as proposed by
others (Nieuwkoop et al., 1952; Leussink, 1970;
Gordon and Brodland, 1987). This model may also
explain the formation of neural tissue in cases of
edgewise induction in Keller sandwiches and exogastrulae as a two step process in which the organizer first
induces the notoplate cells in the noninvoluting
marginal zone. Our results, interpreted in the light of
this model, suggest that Hensen's node may owe its
neural inducing properties to its ability to induce a
notoplate. We cannot rule out the alternative model,
however, that induction of the notoplate and the rest of
the neural plate are both mediated directly, and in
parallel, by interactions between the organizer and
ectoderm.
Induction with Hensen's node is distinct from that of
growth factors such as activin-A
To begin to characterize the inducing signals generated
by organizer tissue, we compared induction by Hensen's node to that by another potent embryonic inducer,
activin-A. Three lines of evidence suggest that the
induction of neural tissue by Hensen's node and activinA involve different pathways. First, recognizable
mesodermal derivatives are not observed when neural
tissue is induced by Hensen's node. Second, the
competence of the ectoderm for neural induction
extends much later than that for induction by activin-A.
Third, the response of the ectoderm to activin-A is
enhanced by lithium treatment while induction by
Hensen's node is unaffected.
In Xenopus, neural tissue has been shown to
differentiate from animal cap ectoderm exposed to
activin-A. The induction of neural tissue by these
factors is always accompanied by the induction of axial
mesoderm structures, in particular notochordal tissue
and muscle. The neural tissue formed in response to
growth factors such as activin-A may therefore be
generated in response to signals derived from the
induced mesoderm rather than from direct actions of
activin-A on the undifferentiated ectoderm. In support
of this, recent studies have shown that dissociated
animal cap ectodermal cells treated with activin-A do
not themselves acquire neural characteristics but do
have neural inducing ability when assayed by the
response of naive animal cap ectodermal cells (Green
and Smith, 1990).
In contrast, it is unlikely that Hensen's node induces
neural tissue indirectly through the induction of
mesoderm. Axial mesoderm is undetectable in the
recombinants by RNAase protection assays and by
morphological and immunohistochemical analysis. Furthermore, in sections of the recombinants the induced
neural tissue is often found to be in contact with the
node. It is possible that Hensen's node induces
prechordal mesoderm, for which there is no reliable
marker in animal cap conjugates and that this mesodermal cell type is responsible for neural induction. This is
unlikely for two reasons. First, prechordal mesoderm is
relatively ineffective in inducing neural tissue in
Xenopus ectodermal caps (Dixon and Kintner, 1989;
Savage and Phillips, 1989; Sive et al., 1989). Second, the
induction of neural tissue with posterior character by
late stage Hensen's nodes suggests that prechordal
mesoderm is not involved.
The different time courses over which ectoderm
responds to activin-A and Hensen's node provide
compelling evidence that the actions of these two
inducers differ. Hensen's node induced neural tissue in
ectoderm taken from late gastrulation stage embryos.
This time course is similar to the competence of the
ectoderm to form neural tissue in response to axial
mesoderm (Sharpe and Gurdon, 1990) and extends
much later than that for mesodermal induction.
Ectoderm responded to activin-A only until just after
the onset of gastrulation. These observations support
the idea that more than one induction step is required
1504 C. R. Kintner and J. Dodd
for the induction of neural tissue by activin-A while, in
contrast, Hensen's node induces neural tissue more
directly.
The evidence described above shows that the neural
inducing signal in Hensen's node cannot be ascribed
solely to an activin-A-like molecule, suggesting that the
Hensen's node-derived signal may be different from
activin-A. Alternatively, the neural inducing properties
of Hensen's node may result from the presence of an
activin-A-like molecule together with a second agent
that alters the response properties of the ectoderm to
the growth factor. In fact, experiments with lithium
have previously shown that the response properties of
ectodermal cells can be altered (Cooke et al., 1989; Kao
and Elinson, 1989). In addition, a dorso-ventral
difference in the response of ectoderm to activin-A has
been observed: activin-A induces anterior dorsal
structures in dorsal ectoderm and ventral mesodermal
derivatives in ventral ectoderm (Ruiz i Altaba and
Jessell, 1991; Sokol and Melton, 1991). In contrast, the
response of ectoderm to Hensen's node was not altered
by lithium treatment. Furthermore, in preliminary
experiments we have found that Hensen's node induced
equal amounts of neural tissue in stage 10 dorsal and
ventral ectoderm (not shown). Thus, if activin-A were
produced by Hensen's node, a second factor would
have to be invoked that would abolish the dorso-ventral
difference and maximize the response of the ectoderm
to an activin-A-like molecule so that lithium cannot
potentiate the response further.
Regionalization of neural tissue by Hensen's node
Hensen's nodes taken from embryos at different
developmental stages induced neural tissue with different anterior-posterior character. Nodes dissected from
stage 4 and early stage 5 chicks induced neural tissue
with anterior properties (en-2 positive, XlHbox6
negative) while later-staged Hensen's nodes induced
posterior neural tissue (en-2 negative, XlHbox6 positive). Stage-dependent changes in the signals emanating
from Hensen's node may result from a changing cell
population in the node or from a change in the
properties of the same population of cells within the
node over time. Signals emanating from the organizer
may therefore act early to regionalize the prospective
neural ectoderm along the anterior-posterior axis. This
patterning information may also be retained in differentiated mesoderm, including notochord. This possibility
is supported by recent experiments in which the A-P
distribution of en-2 in Xenopus can be altered by
transplanting underlying notochord originating from
different A-P levels (Hemmati-Brivanlou et al., 1990).
Taken together, our results provide evidence that the
inducing signals underlying neural induction and early
axial patterning are conserved in evolution. Furthermore, organizer tissue acts as a potent neural inducer
even when it fails to differentiate into dorsal mesodermal derivatives. These findings support the model that
early interactions between the organizer and animal cap
ectoderm in the blastula are important components of
neural induction.
We thank Cliff Hume, Tom Jesse 11, Nancy Papalopulu,
Marysia Placzek and Ariel Ruiz i Altaba for helpful, if heated,
discussions about the work and comments on the manuscript.
We are also extremely grateful to Drs Ali Hemmati-Brivanlou
and Harland for providing us with an en-2 cDNA before
publication and to Ray Keller for insights into the gastrulation
movements of neural tissue. Eric Hubel provided excellent
photographic assistance and Ira Schieren generatedfigure1.
The work was supported by grants from the NIH, to C.K. and
to J.D., from The McKnight Fund for Neuroscience, to C.K.
and to J.D., and The Esther A. and Joseph Klingenstein
Fund, to J.D.
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