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Formation of ectopic neurepithelium in chick blastodermsAge-related capacities for induction and self-differentiation following transplantation of quail Hensen's nodes.

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THE ANATOMICAL RECORD 229:437-448 (1990)
Formation of Ectopic Neurepithelium
in Chick Blastoderms: Age-Related Capacities
for Induction and Self-Differentiation
Following Transplantation of
Quail Hensen’s Nodes
MARK S.DIAS AND GARY C. SCHOENWOLF
Department of Anatomy, University of Utah, School of Medicine,
Salt Lake City, Utah 84132
ABSTRACT
Hensen’s node, regarded as the avian and mammalian homologue
of Spemann’s neural inducer (i.e., the amphibian dorsal blastoporal lip), has been
transplanted in many previous studies to the germinal crescent of avian blastoderms to examine ectopic neural induction. All these studies have suffered from
one or more major shortcomings, the most significant of which has been the lack of
a reliable cell marker to determine the contributions of graft cells to ectopic embryos. In the absence of such marker, induced (i.e., derived from the host) and
self-differentiated (i.e., derived from the graft) neurepithelium cannot be distinguished from one another with certainty. We have transplanted quail Hensen’s
nodes to chick host blastoderms and have subsequently used the quail nucleolar
heterochromatin marker to identify graft cells unequivocally. We systematically
varied both donor and host ages (i.e., stages 3-8 and 3-5, respectively) to examine
the effects of age on ectopic neural induction and self-differentiation. Our results
demonstrate that the age of the donor is more critical than that of the host over the
stages examined. With advancing donor age, the frequency of host induction decreases, while the frequency of graft self-differentiation increases. Previous studies
not using cell markers have concluded that the craniocaudal level of the induced
neuraxis is determined by the age of the donor, that is, young donors induce cranial
neuraxial levels, whereas old donors induce caudal levels. By contrast, we found
that with grafts from older donors, neurepithelium was more commonly self-differentiated rather than induced and that progressively more caudal levels of the
neuraxis self-differentiated with advancing donor age. Induction of caudal neuraxial levels never occurred in the absence of induced cranial levels. The frequency of
neural induction was inversely correlated with the age of the donor and directly
correlated with the quantity of graft endodermal cells contributed to the ectopic
embryo, supporting a previous assertion that in avian embryos, the earliest and
principal source of neural inducer lies within the endoderm rather than mesoderm.
From our results, we propose that the role of neural induction is to produce neurepithelium of unspecified regional character, and that the formation of regional
character depends on subsequent morphogenetic events.
One of the key processes in vertebrate morphogenesis is induction, wherein the fate of a particular tissue
is determined through interaction with another, adjacent tissue. One of the most striking inductions-neural induction-occurs during gastrulation. As a result
of neural induction, a region of the ectoderm becomes
determined to form neurepithelium. In amphibians,
the dorsal lip of the blastopore is responsible for induction of neurepithelium. To demonstrate this, Spemann
and Mangold (1924) transplanted tissue from the dorsal blastoporal lip to the ventral region of a host embryo, which would not ordinarily become neurepithelium. An ectopic embryo resulted, which was composed
0 1990 WILEY-LISS, INC.
of neurepithelium, notochord, endoderm, and (variably) somites and surrounding mesenchyme.
Each of the tissues of the ectopic embryo could arise
from the host, through induction; from the graft,
through self-differentiation; or from both host and
graft, through a combination of induction and self-dif-
Received March 29, 1990; accepted June 18, 1990.
Address reprint requests to Dr. Gary C. Schoenwolf, Department of
Anatomy, University of Utah, School of Medicine, Salt Lake City, UT
84132.
438
M.S.DIAS AND G.C. SCHOENWOLF
ferentiation. To distinguish among these, Spemann
and Mangold transplanted grafts from Triton cristatus
to embryos of T. taeniatus; these two species of newt
differ in the amount of intracellular pigment granules.
The use of such heteroplastic transplants revealed that
the mesenchyme, notochord, somites and a portion of
the endoderm were graft derived, having arisen
through self-differentiation, whereas the neurepithelium was mostly host derived, having arisen chiefly
through induction. These experiments led to the recognition of the dorsal blastoporal lip as the organizer of
the amphibian embryo.
Hensen’s node has been regarded as the avian and
mammalian homologue of the amphibian dorsal blastoporal lip (Hara, 1978). Numerous investigators have
examined neural induction in birds by transplanting
Hensen’s node or other portions of the primitive streak
beneath the epiblast of avian blastoderms (Waddington, 1932,1933; Waddington and Schmidt, 1933; Woodside, 1937; Grabowski, 1957,1962; Sher-Pu et al., 1963,
1965; Vakaet, 1964, 1965, 1981, 1984; Gallera and
Ivanov, 1964; I-Lan et al., 1965; Shu-Dung et al., 1965;
Gallera and Nicolet, 1969; Gallera, 1970a,b, 1971; McCallion and Shinde, 1973; Cuevas and Orts Llorca,
1974; Hornbruch et al., 1979; Sanders and Prasad,
1986). However, all these studies suffer from one or
more major shortcomings. Earlier studies were neither
consistent in what was transplanted nor thorough in
analyzing the results of such transplants; in many
(Waddington, 1932, 1933; Waddington and Schmidt,
1933; Woodside, 1937), donor tissues were obtained
from various regions of the blastoderm, including a
variety of primitive streak levels (with or without Hensen’s node) and portions of the head process region (containing neurepithelium, notochord, paraxial mesoderm, and underlying endoderm), or even from entire
blastoderms. In these and other studies (Grabowski,
1957,1962), the ages of either donors or hosts, or both,
either were not stated or their effects were not analyzed in a systematic fashion.
A more significant shortcoming of many previous
studies is that the origin of cells contributing to various
tissues of ectopic embryos could not be determined with
certainty because a reliable cell marker was not used
(Waddington, 1932, 1933; Waddington and Schmidt,
1933; Woodside, 1937; Grabowski, 1957,1962; Sher-Pu
et al., 1963, 1965; Vakaet, 1964, 1965; Gallera and
Ivanov, 1964; I-Lan et al., 1965; Shu-Dung et al., 1965;
Gallera, 1970a,b, 1971). Instead, assumptions were
made regarding the origin of ectopic structures based
exclusively on morphological criteria. For example, ectopic neurepithelium in continuity with or directly subjacent to the host ectoderm has been assumed to be of
host origin, whereas that in continuity with or subjacent to the host endoderm has been assumed to be of
graft origin. The use of such criteria may or may not
permit valid interpretations.
Some recent studies have used reliable cell markers
to examine avian neural induction. Gallera and Nicolet
(1969) used primitive streak grafts labelled with
[3H]thymidine; however, only a single donor age was
examined, and Hensen’s node was not grafted. Hornbruch et al. (1979), Sanders and Prasad (1986), McCallion and Shinde (1973), Cuevas and Orts Llorca (1974),
and Vakaet (1981, 1984) have employed heteroplastic
transplants of Hensen’s node, constructing quaillchick
chimeras, to demonstrate that ectopic neurepithelium
can form both by induction and by self-differentiation.
However, these studies fall short of examining neural
induction in a comprehensive fashion; all have used
donors and hosts of similar age and have examined
only one or two developmental stages, and none has
examined the independent effects of donor and host age
in a systematic manner.
Deficiencies of previous studies on avian ectopic induction were addressed in the present study by transplanting tissue of constant type and quantity, by systematically analyzing the effects of both donor and host
age on the frequency and regional character of induction, and by using a reliable cell marker to determine
the host or donor origin of cells constituting the various
tissues of ectopic embryos. Heteroplastic transplants of
quail Hensen’s nodes were made to chick host blastoderms at several donor and host ages; the quail nucleolar heterochromatin marker was used subsequently to
identify graft-derived cells in sections through ectopic
embryos (Le Douarin, 1973). Hensen’s node was chosen
for several reasons: (1)it is the structure that has been
transplanted most frequently in the past and its transplantation reliably yields ectopic embryos containing
neurepithelium (reviewed by Waddington, 1952; Gallera, 1971); (2) it is considered to be homologous to the
neural inducer of amphibians (Hara, 1978); (3) it has
well-defined boundaries and can be readily identified
in donor blastoderms; and (4) it contains, a t various
times, both presumptive endoderm and presumptive
chordamesoderm (Nicolet, 1970, 1971), both of which
have been strongly implicated in neural induction
(Hara, 1961, 1978; Gallera and Nicolet, 1969).
The use of the quail cell marker provided important
new information. First, it demonstrated definitively
the origin of ectopic tissues and suggested that assumptions of previous investigators regarding the origin of
some of these may have been erroneous. For example,
ectopic neurepithelium adjacent to host ectoderm did
not always arise through induction as had been assumed; in some instances, it arose through self-differentiation. Second, the incorporation of graft cells into
normal host tissues, such as the gut, the endodermal
layer of the proamnion and the neural tube, could be
recognized; this was previously impossible with homoplastic transplants.
The present study addresses several questions regarding the interaction of tissues during the formation
of ectopic embryos:
1. During which stages is the host ectoderm competent to form a neurepithelium under the inductive influence of grafted Hensen’s node?
2. During which stages is Hensen’s node capable of
inducing neurepithelium from the host ectoderm?
3. During which stages is Hensen’s node capable of
self-differentiating neurepithelium?
4. What effect, if any, do host tissues have on the
graft’s ability to self-differentiate neurepithelium?
5. To what extent does induced ectopic neurepithelium undergo the characteristic morphogenetic events
of normal neurulation?
6. Is the induced or self-differentiated neurepithelium sufficiently organized to permit identification of
NEURAL INDUCTION AND SELF-DIFFERENTIATION
specific craniocaudal levels of the neuraxis, and is either the donor or host capable of influencing which
levels are formed?
7. Which graft-derived tissues are formed, and
which of these are capable of inducing neurepithelium?
Our results demonstrate that both neural induction
and self-differentiation occur in ectopic embryos; the
extent to which each occurs is dependent on the age of
both the donor and host at transplantation. Induction
of neurepithelium is most vigorous when grafts from
young donors are transplanted to young hosts, whereas
self-differentiation is most robust when grafts from
older donors are transplanted to older hosts. Neither
donor nor host age has any effect in specifying craniocaudal level of the induced neuraxis, but with advancing donor age, progressively more caudal regions of the
neuraxis self-differentiate from the graft. These results
provide a clearer understanding of Spemann’s organizer in avian embryos and establish baseline data for
future studies of neural induction.
MATERIALS AND METHODS
To prepare host embryo cultures, White Leghorn
chicken eggs were incubated in humidified incubators
for 12-15 h; blastoderms, together with large portions
of their adjacent vitelline membranes, were removed
from the eggs and prepared for modified New culture
(New, 1955). Each blastoderm, still attached to its vitelline membrane, was oriented ventral side up and
placed in a 35-mm Petri dish on an agar-albumen substrate consisting of 0.6% Bactoagar (Difco Laboratories, Detroit) in 123 mM NaCl mixed 1 : 1with albumen obtained from fresh, fertilized eggs. A glass ring
was then positioned around each blastoderm, and the
vitelline membrane was draped over the ring. Such
cultured chick blastoderms a t stages 3-5 (Hamburger
and Hamilton, 1951) were used as hosts. Those at Hamburger and Hamilton’s stage 3 were further subdivided
into four substages, designated 3a, 3b, 3c, and 3d, as
described by Schoenwolf (1988).
Quail blastoderms were used as donors. These were
removed from their vitelline membranes and placed
dorsal side up in Spratt culture (Spratt, 1947), using
the culture medium just described. Hensen’s node
grafts were obtained from these blastoderms at stages
3-8 (Hamburger and Hamilton, 1951), with subdivisions of stage 3 as described above. Each donor Hensen’s node was excised with a cactus needle mounted on
a wooden handle, cutting along the cranial and lateral
contours of the node. A small portion of the postnodal
primitive streak (<lo0 pm in length) was included
with the graft. The grafts so excised were approximately bullet shaped, facilitating identification of their
cranial and caudal ends. Dorsal and ventral sides were
also identifiable because the grafts typically curled
ventrally.
Each excised graft was transferred with a micropipette from the donor blastoderm and placed atop the
host blastoderm. A small hole was made in the hypoblast of the host’s germinal crescent, and the graft was
gingerly inserted between the hypoblast and ectoderm
near the cranial margin of the area pellucida. Grafts
were oriented with their cranial edge directed toward
the caudal end of the host blastoderm. This allowed
439
regression of the grafted node toward the area opaca, so
that growth of the ectopic embryo would be less encumbered by growth of the host embryo. Most grafts were
placed with their ectodermal side adjacent to the host
ectoderm, mimicking the dorsoventral orientation of
the host. Some grafts were placed with their hypoblast
side adjacent to the host ectoderm; that is, the grafts
were placed upside down in relationship to the host.
The results obtained with the latter grafts were similar
to those obtained with the former; therefore, data were
combined. Excess saline was removed from the hosts
with a micropipette, and the cultures were returned to
the incubator for an additional 24-30 h.
Both donor and host blastoderms were videotaped
during the grafting procedure to document the stages
of each and to record the extent of graft excision and its
position of placement in the host. All blastoderms were
videotaped again when the cultures were terminated;
selected ones were also photographed.
Of 140 blastoderms with Hensen’s node grafts, 96
formed ectopic embryos; these were prepared for histological study. Each was fixed overnight in a mixture of
glacial acetic acid, 37% formaldehyde solution, and
100% ethanol (1:2:7, v:v:v), processed for paraffin embedding and serially sectioned in the plane transverse
to the longitudinal axis of the ectopic embryo. Sections
were processed according to the Feulgen-Rossenbeck
procedure described by Lillie (1965).
Donors and hosts were grouped according to stage
into one of three categories. For donors, the three categories encompassed stages 3a-3c (hereafter designated as young grafts), 3d-4 + (intermediate grafts)
and 5-8 (old grafts). For hosts, the three categories
encompassed stages 3b-3c (young hosts), 3d-4 (intermediate hosts), and 4 -5 (old hosts). Each transplantation thus involved a combination of graft and host of
specified stages. Results were recorded in 9-quadrant
grids, with each cell representing a combination of donor and host ages. We were therefore able to discern
the independent effects of donor and host age on both
neural induction and self-differentiation.
+
RESULTS
Utility of the Quail Nucleolar Marker
Ectopic embryos often had a gross appearance very
similar to that of host embryos (Fig. 1). In serial sections through ectopic embryos, the quail marker was
used to ascertain the origin of cells contained within
various tissues (Figs. 2-4). Ectopic embryos were chimeric, consisting of some tissues derived exclusively
from graft cells (e.g., notochord and somites), others
derived exclusively from host cells (e.g., surface ectoderm) and still others derived from both graft and host
cells (e.g., mesenchyme and endoderm). Ninety of the
96 ectopic embryos contained neurepithelium. In some
cases, the neurepithelium arose by self-differentiation
(Fig. 2a,b) and in others by induction (Fig. 3a,b). Graft
cells incorporated with tissues of the host embryo
proper; in all cases, donor cells incorporated with the
host’s gut (Fig. 4a) and/or endodermal layer of the
proamnion, and in one case, donor cells incorporated
with the host’s neural tube (Fig. 4b). The contributions
of donor cells to these structures would not have been
recognized if a cell marker had not been used.
440
M.S. DIAS AND G.C. SCHOENWOLF
Figs. 1-4.
NEURAL INDUCTION AND SELF-DIFFERENTIATION
441
Fig. 5. Transverse sections from ectopic embryos collected approximately 24 h after transplantation of a quail Hensen’s node. a: Neurallike vesicle (arrow), which formed from quail cells (Hensen’s node
transplanted at stage 4 to a stage 3c host). Note graft cells in the
endodermal layer of the host’s proamnion (arrowheads). b Gutlike
vesicle (arrow), which formed from quail cells (Hensen’s node transplanted at stage 4 to a stage 3c host). Arrowheads, induced neural
plate (on right) continuous with host’s neural plate (on left). Asterisk
(*) indicates the transition between the induced and host’s neural
plate. a: ~ 3 0 0b; ~ 2 1 0 .
Frequency of Neural Induction and Self-differentiation
in Ectopic Embryos
In addition, in 41 ectopic embryos, the grafts formed
vesicles of indefinite character. On the basis of morphology and spatial relationships to other tissues,
these vesicles were tentatively classified as neural tissue in 12 embryos (Fig. 5a) and as gut epithelium in
the remaining 29 (Fig. 5b). To avoid error, these structures were excluded from the data in the analysis described below.
Ectopic neurepithelium was induced most frequently
(93% of cases) when grafts from young donors were
transplanted to young hosts (Fig. 6a). Induction was
common (frequency >50%) when young or intermediate grafts and hosts were combined, when young grafts
and old hosts were combined or when old grafts and
young hosts were combined. Induction was less frequent with both advancing graft and host age; it declined by an average of 36% when graft age was increased from young to old, and by an average of 41%
when host age was increased from young to old (see
Fig. 6a legend for details of averaging). These effects
were additive; when old grafts were combined with old
hosts, only 14%of the ectopic embryos formed induced
neurepithelium.
By contrast, neurepithelium self-differentiated least
frequently when grafts from young or intermediate donors were transplanted to young or intermediate hosts
(Fig. 6b). Self-differentiation occurred frequently when
old grafts were combined with hosts of any age, or
when old hosts were combined with grafts of any age.
In ectopic embryos, neurepithelium formed solely by
self-differentiation of graft cells in 38 (42%) of cases
and solely by induction of host cells in 27 (30%);a further 25 ectopic embryos (28%of cases) each contained
more than one portion of neurepithelium, with oneformed by induction and one or more formed by selfdifferentiation. In total, neurepithelium was induced
in 63 embryos and it self-differentiated in 52.
Fig. 1. Dorsal view of an ectopic embryo (asterisk) approximately 24
h after transplantation of a quail Hensen’s node to the germinal crescent. Note that its morphology is similar to that of the host’s. FB,
forebrain. x 40.
Fig. 2. Transverse section from an ectopic embryo collected approximately 24 h after transplantation of a stage 8 quail Hensen’s node to
a stage 4 host. b is a n enlargement of part of a. Neural tube (hindbraidspinal cord; NT), notochord (N), somites (S), lateral plate (LP)
and midline gut endodermal cells (E) have formed from the graft;
surface ectoderm (SE) and lateral gut endodermal cells have formed
from the host. Note the quail cell marker in b (indicated in the midline endoderm by arrows). a: X 215; b X 560.
Fig. 3. Transverse section from an ectopic embryo collected approximately 24 h after transplantation of a stage 4 quail Hensen’s node to
a stage 4 host. b is a n enlargement of part of a. Notochord, somites,
lateral plate and gut endodermal cells have formed from the graft.
Note the quail cell marker in b (examples indicated in the notochord
by arrows). a: X 215; b X 710.
Fig. 4. Transverse sections from ectopic embryos collected approximately 24 h after transplantation of quail Hensen’s nodes. a: Graft
cells have incorporated with the floor (asterisk) of the host’s foregut
(Hensen’s node transplanted at stage 3a to a stage 3d host). b: Graft
cells (arrow) have incorporated with the host’s neural tube (forebrain/
midbrain) (Hensen’s node transplanted at stage 3c to a stage 3c host).
x 275.
Regional Character of the Ectopic Neuraxis
Neurulation of induced neurepithelium in ectopic
embryos closely mimicked that occurring in host embryos (Figs. 1, 7). The induced neural plate underwent
characteristic shaping (i.e., apicobasal thickening,
transverse narrowing, and longitudinal lengthening)
442
M.S. DIAS AND G.C. SCHOENWOLF
FREOUENCY OF NEURAL INDUCTION
HOST AGE
Young
Intermediate
Old
(3b - 3c)
(3d - 41
(4+ - 5 )
80% (12/15)
76% (13/17)
42% (5/12)
14% (1D)
FREOUENCY OF NEURAL SELF-DIFFlERENTZATION
HOST AGE
Young
Intermediate
b
I
Intermediate
29% (4/14)
42% (5/12)
27% (4/15)
35% (6/17)
I
Old
80% (4/5)
I
I
Fig. 6. Tables showing the frequency of neural induction (a)and
self-differentiation (b)after transplantation of quail Hensen’s nodes.
Numbers in parentheses indicate number of positive cases/total number of ectopic embryos. Tables are broken up into groups with similar
results. To determine the percentage decrease of induction as a function of advancing donor age, the difference between the “young” and
“old” percentages for each column were averaged. Similarly, to determine the percentage decrease of induction as a function of advancing
host age, the difference between the “young”and “old”percentages for
each row were averaged.
and bending (i.e., formation of median and dorsolateral
hinge points and elevation and convergence of neural
folds in proper relationship to the hinge points) (for
details of these processes in normal embryos, see
Schoenwolf, 1982, 1985; Schoenwolf and Smith, 1990).
Moreover, closure of the neural groove occurred on
schedule at each craniocaudal level, and neural crest
cells left the roof of the neural tube and began their
migration. Finally, typical spatial relationships formed
among the neural tube, notochord, somites, surface ectoderm, and gut endoderm, even though these structures were collectively derived from both graft and host
cells.
Embryos with induced ectopic neurepithelium were
examined to determine whether the neuraxis had welldefined regional character (that is, typical forebrain,
midbrain, hindbrain, and spinal cord levels). Regional
character was identified using specific criteria. Forebrain was defined by its broad width and long neural
folds, the presence of optic vesicles, and the absence of
neural crest cells (Fig. 7a); the lack of an underlying
notochord was not, by itself, an identifying feature of
forebrain in the absence of other criteria. Midbrain was
defined by its oval shape with abundant neural crest
cells (Fig. 7b). Hindbrain was defined by its narrower
shape and the presence of either otic placodes or
somites (Fig. 7c). Spinal cord was defined by its characteristic tall, narrow shape with a small, slitlike central canal, and by the presence of somites (Fig. 7d).
Neurepithelium that did not fit the above criteria was
labelled only as neural tube/plate.
Regional character developed in 45 of the 63 ectopic
embryos (71%) with induced neurepithelium (Fig. 8a).
Most frequently (40 cases) only rostra1 levels of the
neuraxis (forebrain with or without midbrain) were induced. Less frequently (5 cases), the induced neuraxis
began rostrally with forebrain and continued caudally
to hindbrain or spinal cord levels. Caudal levels of the
induced neuraxis were never formed in the absence of
more cranial levels. In one case, a small neural structure was induced (Fig. 9). This structure might have
been mistaken for a spinal cord in a few sections; however, serial sections revealed that it was a vesicle
(<lo0 pm in length) rather than a tube. Therefore, it
was classified as induced neuraxis lacking regional
character.
Recall that neural induction was most frequent with
combinations of young donors and hosts (Fig. 6a); similarly, the neuraxis from such combinations frequently
developed regional character (77% of cases) (Fig. 8a).
Moreover, neuraxes with regional character were induced frequently (250%of cases) when young or intermediate grafts were combined with any age host. The
percentage of induced neuraxes with regional character declined by an average of 71% when comparing
young and intermediate grafts with old grafts, and regional identity developed in only 1 of 9 cases (4%) in
which old grafts were used. By contrast, host age did
not affect whether the induced neuraxis developed regional character.
Self-differentiated neurepithelium developed regional character less frequently than did induced neurepithelium. Usually, self differentiated neurepithelium completely lacked regional character (Fig. 8b).
Recall that neural self-differentiation was least frequent with combinations of young donors and hosts
(Fig. 6b); similarly, neurepithelium from such combinations lacked regional character. By contrast to the
lack of an effect of donor age on the level of the
neuraxis that was induced, donor age had a pronounced
effect on the level of the neuraxis that self-differentiated. With young grafts, only 1of 16 embryos (6%)had
a self-differentiated neuraxis with regional character;
it exhibited cranial (forebraidmidbrain) morphology.
In contrast, with old nodes, 12 of 22 cases (55%)had a
self-differentiated neuraxis with regional character; of
these, all but one exhibited caudal (hindbraidspinal
cord) morphology. With intermediate nodes, the response was in between that obtained with young and
old nodes; 6 of 14 cases (43%) had a self-differentiated
neuraxis with regional character: 4 exhibited cranial
morphology and 2 exhibited caudal morphology. Host
age had no consistent effect on the regional character of
the self-differentiated neuraxis.
Formation of Non-neural Structures in Ectopic Embryos
Although neurepithelium was the most conspicuous
structure formed in ectopic embryos, several nonneural structures also were present in many specimens. Surface ectoderm, otic placodes and, in part, en-
NEURAL INDUCTION AND SELF-DIFFERENTIATION
443
Fig. 7. Transverse sections from ectopic embryos collected approximatley 24 h after transplantation of quail Hensen’s nodes. a: Induced
forebrain (Hensen’s node transplanted a t stage 4 to a stage 4 host).
Arrows, quail endodermal and mesodermal cells; arrowhead, induced
lens placode. b Induced midbrain (Hensen’s node transplanted a t
stage 4 to a stage 4 host). Arrow, quail notochord. c: Induced hind-
brain (Hensen’s node transplanted a t stage 4 to a stage 4 host). Arrows, quail notochord and somite; arrowhead, induced otic placode. d,
Induced spinal cord (Hensen’s node transplanted at stage 4 to a stage
3c host). Arrows, quail notochord (flexed) and lateral plate (somatic
and splanchnic layers). x 270.
doderdgut were formed from host tissues in ectopic
embryos. Non-neural structures of graft origin included endoderm in 96 cases (100% of the embryos),
mesenchyme and/or lateral plate in 83 cases (86%),notochords in 66 cases (69%), and somites in 26 cases
(27%).
Notochords formed least frequently with young
grafts, and their frequency generally increased with
advancing graft age (Fig. 10a). In a manner reminiscent of that seen with self-differentiated neural tubes
(cf. Figs. 10a and 6b), notochords frequently developed
following combinations of intermediate or old grafts
with any age host, or of young grafts with old hosts.
The frequency of notochord formation was unaffected
by host age. Somites also formed least frequently with
young grafts and generally increased in frequency with
444
M.S. DIAS AND G.C. SCHOENWOLF
REGIONAL CHARACTER OF INDUCED NEURAXIS
HOST AGE
Young
Intermediate
Old
77% (10/13)
23% (3/13)
89% (8/9)
11% (1/9)
80% (4/5)
20% (1/5)
I
Wl
I
50% (1/2)
0%
50% (1/2)
85% (11/13)
25% (3/12)
25% (3/12)
7.5% (1/13)
7.5% (1/13)
I
I
I
I
I
I
0%
100% (1/1)
20% (1/5)
80% (4/5)
I
8a
I
I
I
REGIONAL CHARACTER OF SELF-DFFERENTIA’IED NEURAXIS
HOST AGE
F/M
T
Young
Intermediate
0%
20% (1/5)
I
Intermediate
2
wc
T
8
-8
100% (7/7)
16.5% (1/6)
16.5% (1/6)
67% (4/6)
50% (2/4)
0%
50% (2/4)
‘I
r
9% (1/11)
Old
WC
b
I
1
1
33% (2/6)
67% (4/6)
55% (6/11)
36% (4/11)
60% (3/5)
40% (2/5)
T
I
I
I
25% (1/4)
25% (1/4)
50% (2/4)
F/M
0%
80% (4/5)
100% (4/4)
!?I
Old
J’
I
Fig. 8. Tables showing the frequency with which regional subdivisions formed in induced (a)and
self-differentiated (b)ectopic neuraxes. F/M, forebraidmidbrain; F/C, forebraidspinal cord; H/C, hindbraidspinal cord; TIP, neural tubelplate.
advancing graft age (Fig. lob). Somites frequently developed with old grafts (average of 73% of cases). The
frequency of somite formation was unaffected by host
age.
The quantity of graft-derived endoderm varied with
the graft age. Young grafts typically gave rise to a
large quantity of endoderm, some of which formed vesicles (see Fig. 3b), some of which incorporated with the
host’s gut (see Fig. 4a) and/or endodermal layer of the
proamnion and some of which appeared as lateral
evaginations of the host’s gut (Fig. 11). By contrast, old
grafts contributed only a few endodermal cells, which
typically incorporated with the midline roof of the
host’s gut (see Fig. 2b).
DISCUSSION
There are four major conclusions of this study. First,
neural induction is optimal in combinations of younger
grafts and hosts, whereas neural self-differentiation is
optimal in combinations of older grafts and hosts. Second, in contrast to previous studies, our results show
that irrespective of graft age, neural induction always
results in formation of cranial neuraxis with or without
more caudal neuraxis, provided that typical regional
character is established in the induced neuraxis. Thus,
graft age does not specify whether cranial or caudal
neuraxis is induced. Third, neurulation of induced ectopic neurepithelium closely mimics that occurring
during normal development. Therefore, this paradigm
can serve as a useful model for studying events underlying neurulation. Fourth, there is a strong correlation
between the quantity of graft endodermal cells incorporated with the host’s tissues and the vigor of the
induction response. From this we infer that prospective
endodermal cells of Hensen’s node likely have a paramount role in neural induction in avian embryos. Collectively, our results suggest that neural induction
produces neurepithelium of unspecified regional character, and that the formation of regional character results from subsequent morphogenetic events.
NEURAL INDUCTION AND SELF-DIFFERENTIATION
Fig. 9. Transverse section from an ectopic embryo collected approximately 24 h after transplantation of a quail Hensen’s node (Hensen’s
node transplanted at stage 5 to a stage 3c host). A small neural vesicle
(asterisk) has been induced from host epiblast. Arrows, quail notochord, somites and endoderm. X 325.
FORMATION OF ECTOPIC NOTOCHORDS
445
Fig. 11. Transverse section from an ectopic embryo collected approximately 24 h after transplantation of a quail Hensen’s node (Hensen’s
node transplanted at stage 4 to a stage 3c host). Arrow, induced neural plate; arrowheads, host neural plate; double asterisk, quail gut
vesicle; single asterisk, lateral extent of the host’s foregut. x 210.
young hosts and decline with advancing host age
(Woodside, 1937; Grabowski, 1962; Gallera and Ivanov,
HOST AGE
1964), although none of these studies used a cell
marker to differentiate between graft and host cells.
Young
Intermediate
Old
Woodside (1937) suggested that competence of the host
43% (6/14)
33% (4/12)
ectoderm is greatest in stage 3 blastoderms, declines
steadily with advancing age and is lost completely beyond stage 6. However, neither donor age nor graft
composition was held constant in that study. Grabowski (1962) reported that induction occurred frequently (70% of cases) after transplantation of stage 4
Hensen’s nodes to stage 4 hosts, whereas it occurred
infrequently (30% of cases) after their transplantation
to stage 5 hosts. Gallera and Ivanov (1964) concluded
that host ectoderm was competent to form ectopic neuFORMATION OF ECTOPIC SOMITES
ral tubes from stages 2-4, but it could form only “neuroidal placodes” between stages 4 and 6, and it comHOST AGE
pletely lost neural competence beyond stage 6. The
frequency of induction of ectopic neurepithelium as a
Young
Intermediate
Old
function of donor age has been virtually ignored. There
W
Young
7%(1/14)
0% (0/12)
11% (1/9)
is only one study suggesting that ectopic neural induc0
tion is more frequent with grafts from younger donors
d
and declines with advancing graft age (Gallera, 1970a).
z
Taken together with other results, our present data,
0
based on the use of a reliable cell marker, demonstrate
9
Old
80%(4/5)
83% (10/12)
57% (4/7)
I
I
I
I
that the frequency of induction depends on both the
b
graft’s inductive ability (a function of donor age) and
the competence of the host ectoderm (a function of host
Fig. 10. Tables showing the frequency of notochord (a)and somite
age) (Fig. 6a). Host competence is present a t the earli(b)formation in ectopic embryos.
est stages examined and declines rapidly after stage 4.
Frequency of Neural Induction Declines With Advancing
Hensen’s nodes from young donors are the most capaHost and Donor Age
ble of inducing neurepithelium from the overlying ecThere is general agreement that both the frequency toderm and this capability is diminished progressively
and the quality of ectopic neural induction are best in with advancing graft age. The effects of advancing
1
446
M.S.DIAS AND G.C. SCHOENWOLF
graft and host age are additive; although an old graft
can still elicit a frequent inductive response with a
young host, or an old host with a young graft, induction
occurs rarely when both graft and host are old.
The quality of the inductive response also declines a t
least with advancing host age; well-formed neural
tubes are induced when both graft and host are young,
whereas nonspecific neuroidal structures are induced
more frequently with advancing host age (Woodside,
1937; Gallera and Ivanov, 1964). The quality of induction also depends on the length of exposure to the inducing tissue; only neural plates are induced after 6 h
of contact between Hensen’s node and the area opaca of
a competent host epiblast, whereas well-formed neural
tubes with regional character develop after 8Yz h of
contact (reviewed by Gallera, 1971). These data are
compatible with models (Saxen, 1980; Gurdon, 1987)
based on interactions between an inducer substance
and a receptor in the competent ectoderm: advancing
donor andior host age could result in a loss of inductive
interaction through a loss of inducer substance or of
host receptors.
Frequency of Neural Self-differentiation Increases With
Advancing Host and Donor Age
In contrast to neural induction, which declines with
advancing graft and/or host age, neural self-differentiation increases with increasing graft or host age or
both. Previous studies have lar ely ignored the effects
of graft or host age on neural se f-differentiation. Only
one has examined graft age (Veini and Hara, 1975);
this showed that when older nodes were transplanted
to the coelomic cavity, the frequency of neural selfdifferentiation increased. None has examined the effects of host age on neural self-differentiation.
Why do graft and host age affect the frequency of
neural self-differentiation? There are three likely possibilities: (1)there are fewer prospective neurepithelial
cells in younger nodes than in older ones; (2) prospective neurepithelial cells are more likely to be lost from
younger nodes than from older ones; and (3)the differentiation of prospective neurepithelial cells is favored
with older nodes, and/or older hosts provide a more
conducive environment for neural differentiation. The
first possibility could explain only the effect of graft
age on neural self-differentiation. Unfortunately, there
is neither evidence for nor against it, so a conclusion
cannot be drawn at present. The second possibility addresses the effect of both graft and host age. Prospective neurepithelial cells could be lost in two ways: by
death, which might be greater after transplantation of
younger nodes (but no evidence exists on this point); or
by dispersion and incorporation with host tissues. In
the latter case, prospective neurepithelial cell dispersion might be greater in younger nodes, containing
principally prospective endodermal cells, than in older
nodes, containing abundant prospective mesodermal
cells (Nicolet, 1970; 1971; Veini and Hara, 1975). Perhaps such prospective mesodermal cells anchor the prospective neurepithelial cells and allow them to undergo
neural differentiation and morphogenesis. The age of
the host also could play a role in the amount of dispersion and incorporation that occur; namely, tissues of
older hosts might inhibit graft cell dispersion and
might be more refractory to the invasion of graft cells
K
than are tissues of younger hosts. The third possibility
also could explain the effect of both graft and host age.
It would be expected that with older grafts, prospective
neurepithelial cells would have had longer exposure to
neural inducer, or would have had more time to initiate
differentiation, before their removal from the donor
and their transplantation to the host. Moreover, with
older hosts it would be expected that more (or perhaps
a “better”) extracellular matrix might have been synthesized in the germinal crescent region, thereby potentially providing a more optimal site for neural differentiation. Obviously, our understanding of neural
induction in avian embryos is far from complete and
further studies will be required to ascertain which of
the above possibilities play the most significant roles in
neural self-differentiation following transplantation of
Hensen’s node.
In addition to the well-formed neural tubes that selfdifferentiated from a large percentage of the grafts,
many vesicles of indefinite character self-differentiated; some of these were more characteristic of neurepithelium and others were more characteristic of gut
epithelium. Both types of vesicles were present in similar locations (often underlying the induced neurepithelium) and many were connected with either the underlying endodermal layer of the proamnion or host
gut. All but two of these vesicles self-differentiated
from grafts encompassing stages 3b-4, when endoderm
is known to be present within Hensen’s node (Nicolet,
1970,1971; Veini and Hara, 1975). Although these vesicles may have appeared to be either more neural or
more endodermal in morphology, depending on the individual case, their tissue of origin remains uncertain
in the absence of either an endodermal or neural-specific marker. Therefore, depending on how these vesicles were classified in previous studies, one could
achieve a higher rate of either homeogenetic induction
(if vesicles were regarded as neural) or nonhomeogenetic induction (if all vesicles were regarded as endodermal).
Regional Identity of the Self-differentiated, But not Induced,
Neuraxis Depends on Graft Age
The concept of an embryonic organizer requires not
only that the organizer be capable of inducing axial
structures, such as neurepithelium, but that it also be
capable of organizing the expression of regional character in these tissues (Grabowski, 1957). Taken one
step further, the assumed homology of Hensen’s node
with the amphibian organizer (i.e., the dorsal blastoporal lip) implies that Hensen’s node is capable of specifying the levels of neuraxis formed from induced tissues. Accordingly, it would be predicted that younger
nodes would induce cranial levels of the neuraxis,
whereas older nodes would induce caudal levels. The
ability of Hensen’s node to control regional organization of the neuraxis was seemingly demonstrated in
avian embryos by Vakaet (1965) and Gallera (1970a);
only caudal levels of the neuraxis were reported to be
induced following transplantation of older grafts,
whereas either cranial levels (Gallera, 1970a) or both
cranial and caudal levels (Vakaet, 1965) were reported
t o be induced following transplantation of younger
nodes.
In contrast to previous studies (Vakaet, 1965;
NEURAL INDUCTION AND SELF-DIFFERENTIATION
Gallera, 1970a), our data do not support the view that
Hensen’s node is responsible for specifying which craniocaudal levels of the neuraxis are induced. Older
grafts in our series never induced caudal neuraxis in
the absence of more cranial neuraxis. However, older
grafts self-differentiated neurepithelium with greater
frequency than they induced neurepithelium, and this
self-differentiated neurepithelium typically had a caudal morphology. We suspect that these self-differentiated neural tubes were mistaken for induced neurepithelium in the previous studies, which did not use a cell
marker. Alternatively, it is possible that neural vesicles (e.g., Fig. 9) were interpreted as spinal cords because of their narrowness. The fact that these structures are spherical rather than tubular argues against
the validity of making such an interpretation.
In addition, our study showed cranial levels of the
neuraxis to be induced more frequently than caudal
levels, regardless of the age of the graft. Moreover,
there is a suggestion that in addition to cranial levels,
caudal levels of the neuraxis were induced more frequently with both advancing graft age and declining
host age (although the number of embryos with induction of both cranial and caudal levels of the neuraxis
was small). The host effect can be explained by our
present results, which clearly reveal that the host’s
neural competence decreases with advancing age.
However, the reason for a donor effect is less clear.
Hensen’s node undergoes progression (i.e., moves cranially) during stages 3a-4 and regression (i.e., moves
caudally) from stage 4 onward (e.g., Nicolet, 1971) and
regional character of the neuraxis is manifested only as
Hensen’s node regresses (e.g.,Schoenwolf,1982;Schoenwolf and Smith, 1990). Hence, older nodes may be able,
by virtue of their stage in development, to organize
induced neurepithelium into its various subdivisions
sooner than would younger nodes. Because cultures
were terminated at similar periods following node
transplantation, regardless of node age, it would be
expected that older nodes would organize more levels of
neuraxis than would younger nodes over the same
length of time.
Our results just discussed are in general agreement
with those of Gallera and Ivanov (1964), although they
differ in that ectopic embryos with both cranial and
caudal levels of the neuraxis formed less frequently in
our study. This disparity may be owing to differences in
the length of culture following transplantation, differences in the culture conditions or to species differences,
as we used heteroplastic rather than homoplastic
grafts.
Ectopic-Induced Neurepithelium Undergoes Typical
Neurulation Events
Neurulation is a complex process involving forces
generated by both the neurepithelium and surrounding
tissues (e.g., Schoenwolf and Smith, 1990). Our results
provide evidence that the induced ectopic neurepithelium undergoes the characteristic morphogenetic
events of neurulation. Of particular importance is the
observation that both median hinge point cells (neurepithelial cells overlying the notochord; MHP cells)
and the more lateral neurepithelial cells (L cells) in
induced ectopic neurepithelium are always derived exclusively from the host ectoderm. Previous experimen-
447
tal evidence suggests that MHP cells are induced by
the subjacent notochord (van Straaten et al., 1988;
Smith and Schoenwolf, 1989), which in our present
study is always derived from graft (quail) cells. This
observation would seemingly suggest that most (or perhaps all) notochordal and MHP cells are not derived
from a common precursor population as has been proposed recently (Jessell et al., 1989).
Endoderm
Is the Most Likely Primary Source of Spemann’s
Neural Inducer in Birds
Graft cells contributed to a variety of non-neural
structures, including notochord, somites, mesenchyme
and endoderm. Although graft-derived notochords and
somites would be identified easily in previous studies
lacking a cell marker, graft contributions to host endoderm, mesenchyme and neural tube would not have
been identified readily without such a marker. The
quail nucleolar marker enabled us to make correlations between the extent to which graft cells contributed to these tissues and the induction of ectopic neurepithelium. No obvious correlations between the
presence of graft-derived notochord, somites, neurepithelium or mesenchyme and the frequency of ectopic
neural induction were found. By contrast, we found a
striking correlation between the degree to which graft
cells contributed to the host endoderm and the frequency of neural induction. These cells not only incorporated with the host’s endoderm but frequently
extended laterally, subjacent to the induced neurepithelium. Ectopic neurepithelium was induced more frequently when young grafts (which contain more endoderm: Nicolet, 1970,1971; Veini and Hara, 1975) were
used, and such grafts contributed a greater quantity of
cells to the host endoderm. Graft cells in the host endoderm are not necessarily exclusively prospective endodermal cells. It is possible that some prospective ectodermal or mesodermal cells may have changed their
fates following transplantation and incorporated with
the host endoderm. Nevertheless, our results support
the assertion of Gallera and Nicolet (1969) that the
source of the neural inducer in birds is the endoderm
and not the mesoderm (see also Hara, 1978, for a
review). Further studies are underway to determine
the precise role of the endoderm in neural induction in
avian embryos.
ACKNOWLEDGMENTS
This research was supported by grant NS18112 from
the National Institutes of Health. We wish to acknowledge the technical assistance of Fahima Rahman and
secretarial assistance of Jennifer Parsons.
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