The role of actin cables in directing the morphogenesis of the pharyngeal pouches
Robyn Quinlan1,
Paul Martin2,* and
Anthony Graham1,
1 MRC Centre for Developmental Neurobiology, 4th Floor New Hunts House, Guys
Campus, Kings College London, London SE1 1UL, UK
2 Department of Anatomy and Developmental Biology, University College London,
Gower Street, London WC1E 6BT, UK
Author for correspondence (e-mail:
anthony.graham{at}kcl.ac.uk)
Accepted 27 October 2003
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SUMMARY
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The pharyngeal arches are separated by endodermal outpocketings, the
pharyngeal pouches. These are structures of considerable importance; they are
required to segregate the mesenchymal populations of each arch and to induce
the formation of arch components, and they generate specific derivatives,
including the parathyroid and the thymus. The pharyngeal pouches are first
evident as localised sites at which the endoderm contacts the ectoderm, and
they then expand along the proximodistal axis to generate the narrow, tight
morphology of the mature pouch. We currently have no knowledge of the
morphogenetic mechanisms that direct formation of the pharyngeal pouches.
Here, in chick, we show that cells within the pharyngeal pouch endoderm have
an abundance of apically located actin fibres that are networked within the
endodermal sheet, via their insertion into N-cadherin adherens junctions, to
form a web of supra-cellular actin cables. Cytochalasin D disruption of these
actin structures results in the formation of aberrant pouches that fail to
generate their normal slit-like morphology. This suggests that the process of
pharyngeal pouch morphogenesis involves the constraining influence of these
actin cables that direct expansion, within the pouch, along the proximodistal
axis. These results, importantly, provide us with vital insights into how the
pharyngeal pouches form their normal morphology. They also give evidence, for
the first time, of actin cables functioning as constraints during complex
vertebrate morphogenetic episodes.
Key words: Pharyngeal pouches, Pharyngeal endoderm, Actin cables, Morphogenesis, Chick
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Introduction
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The pharyngeal arches are evident as a series of bulges found on the
lateral surface of the head of all vertebrate embryos, and it is within these
that the nerves, muscle, skeleton and epithelia of the pharynx originate and
differentiate. Significantly, the development of this territory is extremely
complex involving interactions between a number of disparate embryonic cell
types: ectoderm, endoderm, mesoderm and neural crest, each of which generates
particular components of the arches, and whose development must be coordinated
to generate the functional adult oro-pharyngeal apparatus
(Graham and Smith, 2001
).
Previous studies have, for a number of reasons, tended to focus on the role
of the neural crest in guiding the development of the pharyngeal arches
(Graham et al., 1996
). We are,
however, gaining a greater understanding of how the pharynx is constructed and
it is now clear that our previous ideas must be reassessed. In particular, it
is becoming apparent that the neural crest plays a less pervasive role than
was previously believed, and that much of the development of the pharynx is
dependent upon cues from the endoderm. Importantly, we have recently shown,
using ablation experiments in the chick, that pharyngeal arches can form, are
regionalised and have a sense of identity in the absence of neural crest
(Veitch et al., 1999
). Indeed,
an event that presages pharyngeal arch formation is the development of the
pharyngeal pouches, and it is likely that it is these structures that play a
central role in arch development. The pharyngeal pouches are first evident as
outpocketings of the endoderm, which contact the ectoderm. At these defined
points, the ectoderm and endoderm remain in intimate contact and expand along
the proximodistal axis (Fig.
1). They thus come to separate the neural crest and mesodermal
cells of the arches and to define the anterior and posterior limits of each
arch. The pouches also act to induce the formation of particular arch
components, such as the epibranchial placodes
(Begbie et al., 1999
), and
themselves generate specific derivatives, including the parathyroid and
thymus. Significantly, in the zebrafish mutant vgo, in which the
pharyngeal pouches fail to form, pharyngeal arch development is severely
perturbed (Piotrowski and
Nusslein-Volhard, 2000
).

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Fig. 1. The pharyngeal pouch endoderm supports a two-dimensional web of
supra-cellular actin cables. Side views of embryos (A-F) where Bmp7
expression (A-C) within the pharyngeal pouch endoderm has been used to
highlight the elaboration of pouches along the proximodistal axis and
phalloidin staining (D-F) has been used to visualise f-actin within the pouch
endoderm. (A) At stage 14, two pharyngeal pouches (1pp and 2pp) have formed
and begun to elongate along the proximodistal axis. (B) At stage 15, the third
pharyngeal pouch (3pp) is evident. (C) At stage 18, all three pouches (1pp,
2pp and 3pp) have further elongated and display typical narrow slit-like pouch
morphology. (D-F) Localised accumulation of actin is seen within the endoderm
of each pouch as they undergo proximodistal elongation. (G) High magnification
view of the third pharyngeal pouch (3pp) shows actin organised into a
supracellular actin cable (red arrowhead), assembled along the apical margin
of the endodermal cells (basal margin indicated by an asterisk). (H)
Longitudinal section through the pharyngeal region, (at level indicated in F),
showing pouches as distinct outpocketings of endoderm. The pharyngeal endoderm
is described as pouch endoderm (PE) or interpouch endoderm (IPE). A
two-dimensional web of supracellular cables, which follows the lumenal
contours of the pouch endoderm, appears to be localised to regions where
pouches are forming (red arrowhead) or have just formed, but at lower
abundance in the interpouch endoderm (IPE). OV, otic vesicle; aa, arch artery.
Anterior is towards the left.
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Although the pharyngeal pouches are important, we know very little about
how their morphogenesis is directed. It is clear that this must involve a
remodelling of the endoderm, such that the initial outpocketing that defines
the pouch elaborates along the proximodistal axis, adopting a narrow slit-like
morphology. Interestingly, studies into a variety of other morphogenetic
episodes in both vertebrates and invertebrates
(Jacinto et al., 2001
) have
described the involvement of actin cables that function at the level of the
tissue to bring about specific movements. We have therefore analysed whether
such actin structures are associated with the pharyngeal pouches and, if so,
what role they may play in directing their morphogenesis. We have found that
actin cables, linked via N-cadherin adherens junctions, are indeed a feature
of the pouch endoderm as it is undergoing proximodistal expansion.
Furthermore, we have also demonstrated that if these cables are disrupted, the
pouches fail to generate their normal morphology; thus we propose that these
cables are required to act as constraints directing the growth of the pouches
along the proximodistal axis.
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Materials and methods
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In situ hybridisation
Chick eggs were incubated at 38°C in a humid atmosphere, staged
according to Hamilton and Hamburger
(Hamburger and Hamilton, 1951
)
and fixed overnight at 4°C in MEMFA. In situ hybridisation was performed
as described by Henrique et al. (Henrique
et al., 1995
). The probes used have all been described previously:
Bmp7, Pax1, Dlx2 and N-cadherin. Following in situ hybridisation,
embryos were either bi-sected and mounted under 90% glycerol, or embedded in
gelatin-albumin with 2.5% glutaraldehyde and vibratome sectioned at 50
µm.
Wholemount phalloidin staining and immunohistochemistry
MEMFA-fixed embryos were washed in PBS + 1% Triton X-100 (PBSTx). For
phalloidin staining of f-actin, embryos were incubated overnight at 4°C
with 6.6 nM Alexa-Fluor 488 Phalloidin (Molecular Probes). For nuclear
staining, embryos were briefly washed in 2xSSC (pH 7.0) + 1% Triton
X-100, treated with RNase A (1 mg/ml) for 30 minutes at 37°C, then
incubated with propidium iodide (100 ng/ml) either 40 minutes on the bench or
overnight with phalloidin at 4°C. For analysis of N-cadherin protein,
embryos were first blocked with PBSTx + 10% heat-treated serum, incubated for
5-7 days with anti-chicken N-cadherin antibody (R&D Systems) diluted
1:500, then washed in PBSTx + 1% serum before adding Alexa-Fluor 568
goat-anti-rat IgG conjugate diluted 1:250, overnight at 4°C. Where actin
and N-cadherin double images were required, the above protocols were followed
with the phalloidin staining performed last.
Confocal analysis of fluorescently labeled embryos
Embryos were washed in PBSTx, then mounted whole under coverslip with
Prolong Anti-Fade (Molecular Probes). Some embryos were first embedded in 20%
gelatin:PBS, the gelatin block fixed in 4% PFA + 0.01% glutaraldehyde
overnight at 4°C, then vibratome-sectioned at 50-70 µm. Optical
sections were collected on a Leica DMRE and a Olympus FluoView FV500
laser-scanning microscope.
Transmission electron microscopy
Embryos were collected and immediately fixed for approximately 4 hours at
4°C in 2.5% glutaraldehyde in 0.2 M phosphate buffer (pH 7.3), to which
6.6 nM phalloidin had been added to help stabilise the actin filaments.
Embryos were then washed in a phosphate buffer, post-fixed in osmium
tetroxide, dehydrated through an ethanol series and embedded in resin.
Ultra-thin sections were cut and mounted on 200 mesh grids, and these stained
with uranyl acetate and lead citrate before being viewed in a Hitachi H7600
transmission electron microscope.
Cytochalasin D treatment
A 100 mM stock solution of Cytochalasin D (Sigma) was prepared in
dimethylsulphoxide (DMSO). This was further diluted in DMSO and then sterile
Howard's Ringer to the required concentration. For delivery, chick eggs were
windowed and staged by injecting Pelikan 17 black ink under the blastoderm.
Two delivery methods were employed: beads soaked in 10 mg/ml cytochalasin D,
which were introduced into the pharyngeal cavity via the hindbrain, or; direct
injection of reagent into the pharyngeal cavity. For injection, final
cytochalasin D concentrations of 1 mM, 100 µM and 10 µM were assessed,
but phenotypes described here were generated at 100 µM cytochalasin D. All
injection solutions had Fast Green dye added (1:10) to visualise delivery of
the solution. Typically a 0.4 nl volume of solution was delivered by
microinjection, either through the hindbrain, for embryos up to approx. stage
12, or for older embryos, through individual pouches on the right-hand side.
For both delivery methods, control embryos were generated using undiluted
DMSO. Embryos were either collected immediately or the eggs resealed and
incubated for up to 20 hours. On collection, all embryos were fixed in MEMFA
and either processed for in situ hybridisation or phalloidin staining.
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Results
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The morphogenesis of the pharyngeal pouches can be readily followed through
the expression of Bmp7, which labels each of the pouches from their
earliest stages (Veitch et al.,
1999
). Pouches form sequentially over a protracted period; by
stage 12 the two most anterior pouches are just evident as localised
outpocketings that bring the endoderm into close contact with the overlying
ectoderm (data not shown). By stage 14, the first and second pouches have
begun to elongate along the proximodistal axis
(Fig. 1A). The presence of the
third pouch is evident by stage 15, and like the first two, it continues to
expand proximodistally as development proceeds
(Fig. 1B,C). Finally, the
fourth pouch forms by stage 19 (data not shown). Interestingly, we have found
that staining of whole embryos with fluorescently tagged phalloidin clearly
reveals a pronounced and localised accumulation of f-actin within each of the
pouches, coincident with their shape change. This abundance of actin is
evident within the endoderm of each pouch throughout the prolonged period of
their proximodistal expansion, which occurs over an
20-hour period. Thus
at stage 14, it is apparent at the apical surface of the first and second
pouches (Fig. 1D) and by stage
16, it is also apparent along the apical surface of the third pouch
(Fig. 1E,F). The nature of its
organisation is readily appreciated when viewed at high magnification, with
actin fibres seemingly forming a supra-cellular `cable' that assembles at the
apical margin of the cells of the pouch endoderm
(Fig. 1G). A longitudinal
section through the pharyngeal region of a stage 18 embryo further reveals
this actin structure following the lumenal contours of the pharyngeal endoderm
(Fig. 1H). It is clear from
Fig. 1G, which is a side view
of the pouch endoderm, and Fig.
1H, which views pouch endoderm in longitudinal section, that this
localised abundance of actin does not represent a simple cable within one
plane of the endoderm. Instead, the actin is organised into a two-dimensional
`web' of supracellular actin cables that runs just below the apical plasma
membrane of the pharyngeal endodermal cells. Importantly, this web of actin is
not found throughout the pharyngeal endoderm, but instead shows a marked
localised accumulation, being most abundant in regions where pouches are
forming and generally at much lower abundance in the interpouch endoderm
(Fig. 1H).
Supracellular actin cables have been described in other systems, most
notably in wounded epidermis and in dorsal closure of the epithelia of
Drosophila embryos (Jacinto et
al., 2001
). Importantly, in both these instances the actin cable
is able to function as a contractile purse-string, extending the full
circumference of the epithelial hole, and acting to close these holes over a
narrow time window of
2 hours. The actin structures observed within pouch
endoderm are not being used to close epithelia. Indeed, neither do they fully
circumscribe either a pouch (Fig.
1G) nor the pharyngeal endoderm per se
(Fig. 1H); instead, they
accumulate within regions of the endoderm that are undergoing morphological
change to form the pouches. The variable distribution of these structures
would be consistent with this web of actin cables being involved, over a
protracted period of around 20 hours in a progressive remodelling of the
endoderm along all three axes to generate the complex three-dimensional shape
of the final pouch structure.
To further detail the supracellular organisation of the actin, we analysed
the pouch endoderm using TEM. This clearly reveals the presence of apically
located filaments connecting via cellular junctions, morphologically similar
to adherens junctions (Fig.
2A). More specifically, we show that it is N-cadherin based
adherens junctions, which are associated with the actin structures.
N-cad expression has been previously described in a number of other
tissues, including the neural tube and somites
(Hatta et al., 1987
); however,
we detail its expression within pharyngeal endoderm and show, by its strong
expression within the pouch but not interpouch regions
(Fig. 2B,C), its association
with pouch morphogenesis. Additionally, transverse sections through pharyngeal
regions reveal that N-cad mRNA is also distinctly localised to apical
regions of the pouch endoderm (Fig.
2D). To demonstrate that the actin fibres are inserting into
N-cadherin adherens junctions, we analysed embryos using both phalloidin and
anti-N-cadherin antibody. A side view of a stage 18 embryo
(Fig. 2E) shows actin and
N-cadherin co-localised at the apical surface of each pouch. Higher
magnification further demonstrates that the actin fibres are joined via
N-cadherin adherens junctions (Fig.
2F). In keeping with our observations on actin deployment within
the pharyngeal endoderm, we found that the N-cadherin protein was most
strongly evident within regions of the endoderm where pouches were forming, or
had just formed. Thus, at stage 16, the highest levels of N-cadherin protein
is seen in the forming third pouch endoderm
(Fig. 2G). Interestingly, in
transverse section it is apparent that, like its mRNA, N-cadherin is only
found in pouch and not in the interpouch endoderm
(Fig. 2H,I).

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Fig. 2. The actin cables are networked within pouch endoderm via insertion into
N-cadherin adherens junctions. The cellular junctions connecting the actin
fibres into supracellular cables were characterised using (A) TEM, (B-D)
N-cadherin in situ hybridisation, and (E-I) confocal analysis of
phalloidin-stained f-actin (green) and anti-N-Cadherin antibody (red). (A) TEM
image of the apical margin of cells within pouch endoderm (stage 17), showing
a bundle of filaments (red arrowhead) running across the cells, just below the
plasma membrane, and connecting via adherens junctions (*). (B,C) Side views
showing N-cadherin expression within pouch endoderm of 1pp and 2pp at
stage 14 (B) and 1pp, 2pp and 3pp at stage 17 (C). (D) Transverse section
through second pouch (2pp) (stage 14); N-cadherin expression is
localised to the apical surface of the pouch endoderm (red arrow). (E) Side
view of a stage 18 embryo showing co-localisation (yellow) of actin (green)
and N-cadherin protein (red) at the apical margin of each pouch (white arrow).
(F) High magnification view of pouch endoderm showing N-cadherin protein (red)
localised to the cellular junctions (white arrow) that support the actin cable
(green). (G) Longitudinal section through a stage 16 embryo, showing an
abundance of N-cadherin protein (red) in the last-to-form pouch (3pp) endoderm
(white arrow). (H,I) Transverse confocal sections through a stage 18 embryo at
the level of (H) the third pouch, indicating the pouch endoderm (PE) and (I)
at lower magnification at the level of the second arch and therefore the
interpouch endoderm (IPE). N-cadherin protein is found in the lateral endoderm
of the pouch (PE in H), but is not apparent in the ventral pharyngeal endoderm
(asterisk in H), or the interpouch endoderm (IPE in I). OV, otic vesicle;
en, endoderm; nc, notochord; aa, arch artery; nt, neural tube; am,
arch mesenchyme; ec, ectoderm. Anterior is towards the left.
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To test the function of this web of supracellular actin cables in pouch
morphogenesis, we targeted the pharyngeal endoderm by introducing cytochalasin
D, which inhibits the new assembly of actin filaments
(Urbanik and Ware, 1989
), into
the pharyngeal lumen. To assess the effect of cytochalasin D treatment on the
actin assemblies, embryos (n=21) were treated for up to 6 hours
before being fixed, stained with phalloidin and then analysed using confocal
microscopy. It was apparent in all embryos treated in this manner that the
supracellular cables had been severely disrupted, and that these continued to
be affected for as long as 6 hours after treatment.
Fig. 3A shows an embryo that
was treated at stage 14 and then allowed to develop for a further 5
hours. The morphology of each of the pouches is clearly affected: the first
pouch appears contorted and the second and third pouches are rounded rather
than having a narrow and slit-like shape.
Fig. 3B shows a higher
magnification view of the second pouch of a different embryo treated as
before; in this specimen, actin within the dorsal tip endoderm is clearly
disorganised and the supracellular cable appears to be lost. These defects are
not seen in untreated embryos (Fig.
1, Fig. 3E) nor in
DMSO treated controls (n=9) (data not shown). Notably, at the
concentrations of cytochalasin D used here, cell proliferation has not been
blocked. At a gross level, the embryos have increased in size and, more
specifically, when these specimens were stained with a number of different
nuclear stains (such as propridium iodide and Topro-3) numerous mitotic cells
were seen within pouch endoderm (Fig.
3C,D), as is normally observed in untreated embryos
(Fig. 3E).

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Fig. 3. Cytochalasin D results in disorganized fibres that fail to form a coherent
actin cable. Confocal analysis of phalloidin-stained f-actin (green) and
propridium iodide (red) was used to assess the effects of cytochalasin D on
the organisation of the actin structures within the pouch endoderm (A-D)
compared with an untreated specimen (E), all at equivalent stages. (A) Side
view of an embryo that was treated by introducing cytochalasin D-soaked bead
into the pharyngeal cavity at stage 14 and incubated for a further 5
hours; already there is evidence of aberrant pouch morphology, where pouches
are contorted (1pp) or `relaxed' (2pp and 3pp). (B) High magnification of the
dorsal tip of a second pouch shows that actin fibres fail to form a
supracellular cable when treated with cytochalasin D; this embryo was treated
by introduction of a bead into the pharyngeal cavity and incubated for a
further 6 hours. (C,D) Mitotic cells (white arrows) are clearly evident within
the pouch endoderm of embryos treated with cytochalasin D, either via a bead
(C) or injection (D), as they are seen in the pouch endoderm of untreated
embryos (E). CD, cytochalasin D. Anterior is towards the left.
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To further investigate the effects of cytochalasin D treatment on pouch
morphogenesis, treated embryos (n=97) were incubated for longer
periods (up to 20 hours) and then in situ analysed using a number of markers
to highlight aspects of pouch morphology. Two different types of effects were
seen dependent on the stage of pouch development when cytochalasin D was
applied. Specifically, when cytochalasin D was delivered to the pharyngeal
cavity prior to stage 14, and the embryo allowed to develop further, dramatic
defects in pouch morphogenesis were observed. The specimens in
Fig. 4A,B typify such effects;
this embryo had cytochalasin D injected into the pharyngeal cavity at stage
11+ and the embryo then incubated for 20 hours to stage 15. This
approach uses the axial rotation of chick embryos at approximately stage 12 of
development, whereby they turn to lie on their left hand side; thus, the
highest concentrations of cytochalasin D within the cavity settle and lie over
the tissue lining the left hand side of the cavity. This method provides an
excellent experimental control as n=24/29 specimens treated in this
manner had severe effects to the left hand side pouches but with contralateral
pouches developing a normal morphology. By using Bmp7 to highlight
pouch morphology, it can be seen that the left hand side pouches of the embryo
shown in Fig. 4A display a
splayed, diamond-shaped morphology following exposure to cytochalasin D, but
by contrast, there is clear evidence of three narrow slit-like pouches on its
contralateral side (Fig. 4B).
These effects were not seen in control embryos injected at equivalent stages
with DMSO carrier alone; only minor defects were seen in these embryos with
n=19/21 displaying normal development
(Fig. 4C). Interestingly, of
all embryos treated with cytochalasin D, in no instance did we find that
cytochalasin D treatment resulted in a total failure to form any one of the
pouches. Instead, as evident in Fig.
4A, pouches form but their growth is not directed proximodistally,
and as such they fail to generate the typical narrow, slit-like morphology. It
seems clear that the actin web is responsible, not for induction, but for
shaping of the pouches and for the subsequent maintenance of their shape.

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Fig. 4. Disruption of actin cable assembly results in pouches with aberrant
morphology, owing to a failure in proper proximodistal elongation. Effects of
cytochalasin D treatment on pouch morphogenesis analysed through (A-C)
Bmp7; (D,F) Pax1; or (G,H) Dlx2 in situ
hybridization. (A,B) Left- and right-hand side views of the same embryo that
had cytochalasin D injected into the pharyngeal cavity at stage 11+, prior to
pouch formation, and the embryo allowed to develop to stage 15. As the
chick embryo turns, gravity causes cytochalasin D to accumulate over the
left-hand side (LHS) tissue, which results in aberrant pouch morphology on
that side (A) but not on the right-hand side (RHS) (B). LHS pouches have an
open diamond shape compared with the narrow, slit-like pouches on the RHS or
in the DMSO control (C) embryo treated in the same manner. (D) Side view of an
embryo where cytochalasin D was injected into the vicinity of the first pouch
(1pp) at stage 15, and allowed to develop to stage 19. (E) High
magnification of the contralateral first pouch, which did not receive an
injection of cytochalasin D, displaying normal slit-like morphology. The dots
outline the contours of the pharyngeal endoderm. (F) High magnification of a
first pouch that did receive a cytochalasin D injection showing a contorted
morphology of the pharyngeal endoderm along the proximodistal axis. Again, the
dots outline the contours of the pharyngeal endoderm. (G) Side view of an
embryo, treated with cytochalasin D by injection into the pharyngeal cavity at
stage 12 and allowed to develop to stage 18 and (H) an embryo similarly
treated with DMSO at stage 13 and allowed to develop to stage 19. In both G
and H, Dlx2 expression shows the arches have been properly populated
with neural crest cells. OV, otic vesicle. Anterior is towards the left.
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When cytochalasin D was injected into the pharyngeal cavity of embryos at
later stages from stage 14 onwards, pouch morphology was also affected, but in
a different manner in n=4/9 specimens.
Fig. 4D shows a side view of an
embryo, with pouches highlighted through Pax1 expression
(Muller et al., 1996
;
Veitch et al., 1999
), that had
received a cytochalasin D injection at stage 15 into the vicinity of
the first pouch. Clearly, further pouch development has not been blocked,
rather the pouch endoderm has assumed a contorted morphology, owing to growth
not being directed appropriately (Fig.
4F). By contrast, the first pouch on the contralateral side, which
did not receive an injection of cytochalsin D, displays the normal slit-like
morphology (Fig. 4E). The
effects on pouch morphology we describe here, are not simply due to an
indirect effect on the pharyngeal arch mesenchyme, because in situ analysis
using Dlx2, which labels the neural crest cells of the pharyngeal
arches (Veitch et al., 1999
),
shows that in both experimental (Fig.
4G) and DMSO control specimens
(Fig. 4H), the neural crest
cells have normally populated the arches. This finding is also consistent with
the fact that the neural crest are not required for normal pouch development;
in the absence of crest cells pharyngeal pouches form, elongate and are
normally patterned (Veitch et al.,
1999
). Thus, the consequences for pharyngeal morphogenesis,
resulting from the direct introduction of cytochalasin D into the pharyngeal
lumen, are specifically due to its disruption of the actin cable network
assembled in the cells of the pharyngeal endoderm.
 |
Discussion
|
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This study has analysed the role of actin in the complex morphogenesis of
the pharyngeal pouches, by revealing how the actin network is organised within
pharyngeal endoderm and by observing the consequences of its disruption with
cytochalasin D. We show that an assembly of actin, networked by N-cadherin
adherens junctions, forms supracellular cables that follow the lumenal
contours of the pouch in such a way as to form a two-dimensional web-like
structure. These actin cables are required for the establishment and
subsequent maintenance of normal pouch morphology and we propose that one
important function of these cables is in ensuring the proper proximodistal
elaboration of the pouches to generate their long, slit-like shape.
Insights into the morphogenesis of the pharyngeal pouches are of crucial
and timely importance. With emphasis now being placed on the role of the
pouches in organising the pharyngeal apparatus as a whole
(Graham, 2001
), it is even more
pertinent to investigate how the morphogenesis of this tissue is controlled.
The results presented here provide us with the first insights into the
cellular mechanisms that guide the complex morphological movements that give
rise to the pharyngeal pouches. During the period of pouch morphogenesis, the
pharyngeal endoderm as a whole is expanding in all directions, but the pouches
themselves are primarily growing along the proximodistal axis. Importantly, it
is during this period of directed proximodistal expansion that the actin
cables form at the lumenal surface of the pouches. Furthermore, interfering
with the actin cables during this time frame results in a failure in directed
proximodistal expansion, and instead the pouches splay open or, if inhibited
at later stages, have a randomly contorted shape. More specifically, these
results suggest these actin cables are functioning as a constraining force
upon the endodermal sheet directing the pouches to primarily elongate along
the proximodistal axis and thus generate their typical narrow, slit-like
shape. Besides directing the proximodistal expansion of the pouches, the actin
cables could also act to give rigidity to the pouch endoderm. For instance, in
a growing sheet, the rigidity afforded by the actin cables would prevent
random buckling caused by the tissue taking up a shape that was simply
preferred by its physical properties and those of its surrounds, thus allowing
the maintenance of specific pouch shapes.
We propose that the actin network operates as a constraint within the pouch
epithelium, thus forcing directed expansion and eventual slit-like morphology.
A useful analogy might be to consider the effect of applying a piece of tape
to a balloon as it is being inflated. Filling the balloon with air equates to
increasing the number of cells within the endodermal sheet, the result being a
uniform expansion of the balloon. Applying a piece of non-expanding tape to
the balloon, however, constrains that region preventing a change in its
surface area at this position; thus, directing expansion and generating a
particular shape. Additionally, the actin cables within the pharyngeal
endoderm, like the tape, act as a constraint that holds a particular shape
over time, while other unconstrained areas continue to expand.
The function of actin as a constraining force within the pharyngeal
endodermal sheet is a departure from the role normally ascribed to actin
networks during morphogenesis, as contractile tools for drawing tissues
together. However, recent studies of dorsal closure in Drosophila
show that here too cables operate not only as purse strings, closing this hole
within 2 hours, but also supply constraining forces, restricting forward
movement of the leading edge, by keeping it taut and thus facilitating an
orderly movement of the epithelia towards the dorsal midline
(Bloor and Kiehart, 2002
;
Jacinto et al., 2002
). It
seems more accurate to view the cables as a contractile apparatus that, rather
than contracting the sheet of cells, is functioning to maintain local tensions
and constrain various regions in order to shape and maintain form and
rigidity. Hence, it would seem that there is accumulating evidence that the
constraining role for actin cables proposed here in pouch morphogenesis is a
fundamental feature of actin cables during the shaping of epithelial tissues.
Indeed, it is likely that actin cables functioning as constraints,
particularly during vertebrate morphogenetic episodes which often involve
complex three-dimensional modelling of tissues over considerably longer time
periods; for example, the elongation of the pharyngeal pouches taking some 20
hours. Neural tube morphogenesis is likely to be another such example of this.
For some time it has been thought that contraction of actin filaments aligned
along the lumenal surface of the neuroepithelium was facilitating aspects of
the elevation and fusion of the tube
(Karfunkel, 1972
;
Lee and Nagele, 1985
;
Morriss-Kay and Tuckett,
1985
). However, more recently it has been suggested that one
function of the actin cables may be to generally maintain rigidity throughout
the neuroepithelium (Ybot-Gonzalez and
Copp, 1999
), which would suggest that in this situation actin
cables are also acting to constrain and hold shape. Furthermore, the role that
we describe here for actin in constraining the morphogenesis of an epithelial
sheet is also important as it is likely to be relevant to the formation of
many vertebrate organs, as these also invariably involve epithelial
remodelling rather than closure of a hole.
 |
ACKNOWLEDGMENTS
|
---|
We thank Ken Brady of the EM unit at Guys Campus (KCL), for his expert
assistance with the TEM study. We also thank Richard Wingate and Imelda
McGonnell for constructive comments on the manuscript. This work was supported
by the Medical Research Council (UK).
 |
Footnotes
|
---|
* Present address: Departments of Physiology and Biochemistry, University of
Bristol, School of Medical Sciences, University Walk, Bristol BS8 1TD, UK 
 |
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