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First published online 7 January 2004
doi: 10.1242/dev.00950
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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
 |
SUMMARY |
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 TOP SUMMARY IntroductionMaterials and methodsResultsDiscussionREFERENCES |
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Key words: Pharyngeal pouches, Pharyngeal endoderm, Actin cables, Morphogenesis, Chick
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Introduction |
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TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
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).
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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) 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. |
Materials and methods |
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TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
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)
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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TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
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). 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).
|
), 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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;
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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TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
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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.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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TOPSUMMARYIntroductionMaterials and methodsResultsDiscussionREFERENCES |
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Begbie, J., Brunet, J. F., Rubenstein, J. L. and Graham, A. (1999). Induction of the epibranchial placodes. Development 126,895 -902.[Abstract]
Bloor, J. W. and Kiehart, D. P. (2002).
Drosophila RhoA regulates the cytoskeleton and cell-cell adhesion in the
developing epidermis. Development
129,3173
-3183.
Graham, A. (2001). The development and evolution of the pharyngeal arches. J. Anat. 199,133 -141.[CrossRef][Medline]
Graham, A. and Smith, A. (2001). Patterning the pharyngeal arches. BioEssays 23, 54-61.[CrossRef][Medline]
Graham, A., Koentges, G. and Lumsden, A. (1996). Neural crest apoptosis and the establishment of craniofacial pattern: an honorable death. Mol. Cell Neurosci. 8,76 -83.[CrossRef][Medline]
Hamburger, V. and Hamilton, H. L. (1951). A series of normal stages in the development of the chick embryo. J. Morphol. 88,49 -92.[CrossRef]
Hatta, K., Takagi, S., Fujisawa, H. and Takeichi, M. (1987). Spatial and temporal expression pattern of N-cadherin cell adhesion molecules correlated with morphogenetic processes of chicken embryos. Dev. Biol. 120,215 -227.[CrossRef][Medline]
Henrique, D., Adam, J., Myat, A., Chitnis, A., Lewis, J. and Ish-Horowicz, D. (1995). Expression of a Delta homologue in prospective neurons in the chick. Nature 375,787 -790.[CrossRef][Medline]
Jacinto, A., Martinez-Arias, A. and Martin, P. (2001). Mechanisms of epithelial fusion and repair. Nat. Cell Biol. 3,E117 -E123.[CrossRef][Medline]
Jacinto, A., Wood, W., Woolner, S., Hiley, C., Turner, L., Wilson, C., Martinez-Arias, A. and Martin, P. (2002). Dynamic analysis of actin cable function during Drosophila dorsal closure. Curr. Biol. 12,1 -20.[CrossRef][Medline]
Karfunkel, P. (1972). The activity of microtubules and microfilaments in neurulation in the chick. J. Exp. Zool. 181,289 -301.[CrossRef][Medline]
Lee, H. Y. and Nagele, R. G. (1985). Studies on the mechanisms of neurulation in the chick: interrelationship of contractile proteins, microfilaments, and the shape of neuroepithelial cells. J. Exp. Zool. 235,205 -215.[CrossRef][Medline]
Morriss-Kay, G. and Tuckett, F. (1985). The role of microfilaments in cranial neurulation in rat embryos: effects of short-term exposure to cytochalasin D. J. Embryol. Exp. Morphol. 88,333 -348.[Medline]
Muller, T. S., Ebensperger, C., Neubuser, A., Koseki, H., Balling, R., Christ, B. and Wilting, J. (1996). Expression of avian Pax1 and Pax9 is intrinsically regulated in the pharyngeal endoderm, but depends on environmental influences in the paraxial mesoderm. Dev. Biol. 178,403 -417.[CrossRef][Medline]
Piotrowski, T. and Nusslein-Volhard, C. (2000). The endoderm plays an important role in patterning the segmented pharyngeal region in zebrafish (Danio rerio). Dev. Biol. 225,339 -356.[CrossRef][Medline]
Urbanik, E. and Ware, B. R. (1989). Actin filament capping and cleaving activity of cytochalasins B, D, E, and H. Arch. Biochem. Biophys. 269,181 -187.[CrossRef][Medline]
Veitch, E., Begbie, J., Schilling, T. F., Smith, M. M. and Graham, A. (1999). Pharyngeal arch patterning in the absence of neural crest. Curr. Biol. 9,1481 -1484.[CrossRef][Medline]
Ybot-Gonzalez, P. and Copp, A. J. (1999). Bending of the neural plate during mouse spinal neurulation is independent of actin microfilaments. Dev. Dyn. 215,273 -283.[CrossRef][Medline]
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