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It's all in your head: new insights into craniofacial development and deformation

2005; Wiley; Volume: 207; Issue: 5 Linguagem: Inglês

10.1111/j.1469-7580.2005.00484.x

1469-7580

Minal Tapadia, Dwight R. Cordero, Jill A. Helms,

Craniofacial Disorders and Treatments

Journal of AnatomyVolume 207, Issue 5 p. 461-477 Free Access It's all in your head: new insights into craniofacial development and deformation Minal D. Tapadia, Minal D. Tapadia Department of Plastic and Reconstructive Surgery, Stanford University, Stanford, California, USASearch for more papers by this authorDwight R. Cordero, Dwight R. Cordero Department of Obstetrics and Gynecology, Brigham and Women‘s Hospital, Harvard Medical School, Boston, Massachusetts, USASearch for more papers by this authorJill A. Helms, Jill A. Helms Department of Plastic and Reconstructive Surgery, Stanford University, Stanford, California, USASearch for more papers by this author Minal D. Tapadia, Minal D. Tapadia Department of Plastic and Reconstructive Surgery, Stanford University, Stanford, California, USASearch for more papers by this authorDwight R. Cordero, Dwight R. Cordero Department of Obstetrics and Gynecology, Brigham and Women‘s Hospital, Harvard Medical School, Boston, Massachusetts, USASearch for more papers by this authorJill A. Helms, Jill A. Helms Department of Plastic and Reconstructive Surgery, Stanford University, Stanford, California, USASearch for more papers by this author First published: 02 November 2005 https://doi.org/10.1111/j.1469-7580.2005.00484.xCitations: 66 Jill A. Helms, Stanford University, 257 Campus Drive, Stanford, CA 94305, USA. T.: +1 650 736 0919; F: +1 650 736 4374; E: jhelms@stanford.edu AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Introduction ‘Our ignorance of the laws of variation is profound. Not in one case out of a hundred can we pretend to assign any reason why this or that part has varied. But whenever we have the means of instituting a comparison, the same laws appear to have acted in producing the lesser differences between varieties of the same species, and the greater differences between species of the same genus.’ (Darwin, 1859). Charles Darwin did not gild the truth when he candidly pointed out just how imperfect is our knowledge of how morphological diversity is generated. Over the intervening decades we have scrutinized the process of embryonic development and, in doing so, gained a deeper appreciation for the tissue interactions that lie at the basis of morphogenesis. More recently, some of the molecules involved in mediating these tissue interactions have been identified and, in a few rare cases, we even have gleaned insights into how species-specific development is controlled. These instances, however, are few and far between and much remains to be accomplished. In this review we will present some of the most recent advances that lie at the heart of understanding the mechanisms controlling normal craniofacial development, the consequences of when these normal pathways are disrupted and how this information sheds light on the basis for evolutionary diversity among the species. We choose to focus on morphogenesis of the craniofacial complex for two reasons. First, faces show tremendous phenotypic variation, first evident during the later stages of fetal development. Yet despite these differences in the facial appearance of embryos, they all look remarkably similar during earlier stages of embryogenesis. This suggests that whatever factors influence craniofacial diversity primarily act during a discrete period of embryonic development; and knowing when diversity first arises is an important first step towards understanding how diversity is generated. The second reason we use craniofacial development as a model is that variations in the craniofacial complex are tightly associated with adaptive radiations into ecological niches. If the relationship between craniofacial morphology and speciation is causal (and not merely correlative) then perhaps we can understand how modifying the spatial and temporal patterns of gene expression create diversity within a species. One final justification for using craniofacial morphology as a model system is the prodigious amount of documentation that has accrued on head and neck anomalies. As the noted embryologist William Harvey so eloquently wrote, ‘Nature is nowhere more accustomed openly to display her secret mysteries than in cases where she shows traces of her working apart from the beaten path; nor is there any better way to advance the proper practice of medicine than to give our minds to the discovery of the usual law of Nature by careful investigation of cases of rarer forms of disease’ (Harvey, 1657). Thus, thoughtful inspection of the malformed face may offer valuable clues into the mechanisms governing normal craniofacial morphogenesis. Components of the craniofacial complex In its postnatal form the vertebrate head has an intricate and highly varied morphology, but during early stages of embryonic development it exhibits a much more simple geometry (Fig. 1). There are seven prominences that comprise the vertebrate face: the midline frontonasal prominence, and three paired structures, the lateral nasal, maxillary and mandibular prominences (Fig. 1D–F). These maxillary and mandibular prominences are derived from the first pharyngeal (branchial) arch, whereas the frontonasal prominence is derived from a midline primordium that forms on top of the forebrain. The frontonasal prominence contributes to the forehead, middle of the nose, philtrum of the upper lip and primary palate (Fig. 1). The lateral nasal prominence forms the sides (ala) of the nose; the maxillary prominences contribute to the sides of the face and lips, and the secondary palate; and the mandibular prominences produce the lower jaw (Fig. 1). Disruptions in the rate, the timing or the extent of outgrowth of any of these prominences will adversely affect the fusion process. Consequently, one can appreciate the wide variety of facial clefts that occur (Tessier, 1976) and also why facial clefting is the most frequently occurring head and neck birth defect (Perrotin et al. 2001). Figure 1Open in figure viewerPowerPoint Development of the craniofacial primordia. (A–D) Representations of frontal views of mouse embryos showing the prominences that give rise to the main structures of the face. The frontonasal (or median nasal) prominence (pink) gives rise to the forehead (A), the middle of the nose (B), the philtrum of the upper lip (C) and the primary palate (D), whereas the lateral nasal prominence (blue) forms the sides of the nose (B,D). The maxillomandibular prominences (green) give rise to the lower jaw (specifically from the mandibular prominences), to the sides of the middle and lower face, to the lateral borders of the lips, and to the secondary palate (from the maxillary prominences). (E) Frontal view of a chick embryo, also showing which prominences give rise to different facial structures. (F) Frontal view of a human child, with different facial structures colour-coded to indicate the prominences from which each structure developed. Craniofacial patterning and the neural crest Each of the pharyngeal arches, and the frontonasal prominence, is composed of mesenchyme surrounded by epithelia. In the case of the frontonasal prominence, the mesenchyme is derived from the cranial neural crest, and the epithelia encasing the prominence include the neuroectoderm of the forebrain, and facial (surface) ectoderm (Fig. 2E,F). In the maxillary and mandibular prominences, the encasing epithelia are derived from facial ectoderm and pharyngeal endoderm, whereas the mesenchyme is derived from both cranial neural crest and mesoderm. Figure 2Open in figure viewerPowerPoint Neural crest induction and migration in the developing embryo. (A) The neural plate consists of a unified layer of ectoderm, beneath which lies the endoderm. The neural folds arise as the ectoderm begins to fold upwards. Interactions between signalling molecules cause the medial portion of ectoderm to begin to assume a neural character (green) while lateral portions of ectoderm begin to take on a non-neural character (blue). The prechordal plate mesendoderm (pcp) and the buccopharyngeal membrane (bpm) are indicated. (B) As the neural folds begin to fuse, the neural tube takes shape, giving rise to distinct tissue layers of neuroectoderm (green) and surface ectoderm (blue). Neural crest cells start to delaminate from the border region between the neuroectoderm and surface ectoderm. (C) Once the neural tube has closed, neural crest cells lie interposed between the facial (surface) ectoderm (fe) and the neuroectoderm (ne). (D–F) As the central nervous system begins to form from the neural tube, the neural crest starts to migrate anteriorly from rhombomeres (r1–r3) into different areas of the face, and into the pharyngeal arches. Abbreviations: C, caudal; is, isthmus; mes, mesencephalon; mn, mandible; PA, pharyngeal arch; pe, pharyngeal endoderm; rp, Rathke's pouch; R, rostral; tel ne, telencephalic neuroectoderm. The cranial neural crest cells populating each arch arise from distinct anterioposterior positions along the neural axis (Kontges & Lumsden, 1996), and once resident in the arches, proliferate in a highly regimented fashion. In some cases, the instructions for this proliferative activity are inherent within the neural crest cells themselves; in other cases, the surrounding epithelia provide the directives. For example, neural crest cells in which Homeobox transcription factor (Hox) genes such as Hoxa2 are expressed are constrained, to a certain degree, in their ability to respond to local cues from overlying or underlying epithelia (reviewed in Le Douarin et al. 2004). Cells devoid of Hox gene expression appear much more responsive to signals emanating from the epithelial environment, even into very late stages of morphogenesis (reviewed in Helms et al. 2005). Some of these epithelial cues have been identified and in the next sections we will review these molecular pathways and what is known about their roles in craniofacial morphogenesis. Not all neural crest cells play by the same rules The presence or absence of a particular Hox code in a neural crest cell determines the extent to which neural crest cells are able to respond to cues from the surrounding milieu (reviewed in Trainor & Krumlauf, 2000; Le Douarin et al. 2004). Other neural crest cells, such as those destined for the frontonasal prominence, do not express Hox genes, and therefore our laboratory set out to determine the extent to which these cells were restricted in their developmental potential; or, phrased another way, the extent to which these cells exhibited plasticity and thus were able to respond to local cues from the facial ectoderm (Schneider & Helms, 2003). To test whether neural crest cells inherently possess directions for facial patterning, we began by exchanging neural crest cells between quail and duck embryos (Schneider & Helms, 2003) (Fig. 3). We specifically targeted neural crest cells destined for the upper beak for two reasons. First, the transplanted neural crest cells were Hox negative and therefore we would be assessing the plasticity vs. pre-patterned status of neural crest cells separate from the function of Hoxa2. Second, the short, narrow quail beak and the long, broad duck bill meant that we could use differences in the morphology of duck and quail faces (Fig. 3A) as a readout of whether neural crest cells contained patterning information. Figure 3Open in figure viewerPowerPoint Transplantation experiments provide evidence that the neural crest inherently contains species-specific patterning information. (A) Quail and duck embryos exhibit distinct anatomical features. For example, quails exhibit a shorter, narrower beak compared with the longer, broader duck bill. (B) When quail neural crest cells from forebrain (fb), midbrain (mb) and rhombomeres 1 and 2 (r1, r2) are transplanted into a duck host, a quail-like beak develops in lieu of a duck's bill. (C) When duck neural crest cells are transplanted into a quail host, the quail develops a duck-like bill. (Figure reproduced courtesy of Nature.) After neural crest cells were exchanged between duck and quail embryos the chimeras were allowed to develop to the stage where morphology of their beaks was evident (Schneider & Helms, 2003). Duck embryos that received grafts of quail frontonasal neural crest cells exhibited short quail-like beaks (‘qucks’) (Fig. 3B) whereas quail embryos that had received transplants of duck neural crest cells had duck-like bills (‘duails’) (Schneider & Helms, 2003) (Fig. 3C). These findings suggested that neural crest cells direct their own morphogenesis according to instructions inherent in the donor population (Schneider & Helms, 2003). To understand the molecular basis for these morphological transformations, we performed the same types of grafting experiments and then used molecular and cellular analyses to determine how the grafted cells acted in their new environment and, in turn, how the environment influenced the behaviour of the transplanted neural crest cells. We found that transplanted neural crest cells maintained the temporal gene expression patterns of their original environment despite being transplanted into a new site. In addition, the transplanted neural crest cells altered the temporal pattern of gene expression to reflect the donor, and not the host, environment (Schneider & Helms, 2003). The same conclusion was reached by Abigail Tucker and Andrew Lumsden, who independently performed similar types of interspecies transplants (Tucker & Lumsden, 2004). They, too, found that the capacity to form species-specific skeletal elements in the head was an inherent property of the neural crest, and concluded that this characteristic is articulated is response to signals from epithelia (Tucker & Lumsden, 2004). In fact, other new experiments directly show that most neural crest cells acquire at least some patterning information from nearby epithelia (reviewed in Helms et al. 2005). The extent to which facial features were transformed was directly proportional to the number of transplanted neural crest cells that made their way into the chimeric tissue. In other words, the transformation was a ‘population-dependent’ effect, quite analogous to previous transplantation studies (reviewed in Helms et al. 2005). So it seems that when the contingency is large enough, neural crest cells follow molecular cues that are generated and maintained by the assemblage itself, and disregard signals emanating from the local environment. Just what these population-dependent cues are, and how many cells are required to maintain them, is completely unknown. The nature of prespecification Hox genes are probable mediators of the population-based behaviour exhibited by neural crest cells (Capecchi, 1997; Barrow & Capecchi, 1999). In the craniofacial region, Hox genes are expressed by neural crest before and after migration to the arches (Hunt et al. 1991b,c; Wilkinson, 1993; Favier & Dolle, 1997; Couly et al. 1998; Rijli et al. 1998; Trainor & Krumlauf, 2000, 2001; Trainor, 2003). These domains are preserved even after neural crest cells take up residence in the branchial arches (Hunt et al. 1991a,b). After migration, Hoxa2-positive neural crest cells occupy the second and more posterior arch mesenchyme, which gives rise to the hyoid bone and other structures (Creuzet et al. 2002) (Fig. 4A). Hoxa2 is not expressed in crest cells of the first arch in most vertebrates (Creuzet et al. 2002) (Fig. 4A). This distinct expression boundary of Hoxa2 has led to speculation that neural crest cells may be prespecified by virtue of the expression of this Hox gene. Figure 4Open in figure viewerPowerPoint Manipulation of Hox expression. (A) In jawed animals, Hoxa2 is expressed up to pharyngeal arch 2 (PA2). (B) Loss of Hoxa2 expression in pharyngeal arch 1 (PA1) gives neural crest cells in PA1 increased plasticity, and they undergo transformation to a first arch fate. (C) When Hoxa2 is ectopically expressed in PA1, neural crest cells in this arch take on second arch fates, giving rise to a duplication of the hyoid arch. Gain- and loss-of-function studies bolster this hypothesis. For example, loss of Hoxa2 from the second arch allowed the second arch to take on first arch character, eventually resulting in duplication of maxillary and mandibular structures (Gendron-Maguire et al. 1993; Rijli et al. 1993) (Fig. 4B). By contrast, over-expression of Hoxa2 in all tissues of the first arch caused the first arch to take on a second arch character (Grammatopoulos et al. 2000; Pasqualetti et al. 2000) (Fig. 4C). Additionally, if Hoxa2 is transfected into the rostral domain of the cephalic neural crest, these cells lose their ability to differentiate into skeletal structures (Creuzet et al. 2002). Transplantation and ablation experiments lend further confirmation that Hoxa2 expression confers upon neural crest cells an anterioposterior identity (Couly et al. 1998; Ruhin et al. 2003). Craniofacial epithelia as sources of instructive signals Given that the Hox transplantation and ablation experiments demonstrated that some neural crest cells are capable of responding to local cues from the surrounding environment, the question then becomes which tissues in the environment are ‘talking’ to the neural crest. Once the neural crest delaminates from the surface ectoderm during neurulation (Fig. 2B), it lies sandwiched between several epithelia: the surface ectoderm, the neuroectoderm and the pharyngeal endoderm (Fig. 2C–F). Its close contact with these epithelia during development (Fig. 2D–F) allows these tissues to provide instructive signals that help to pattern the neural crest. Recent studies have shed light on the role of the surface ectoderm in initiating outgrowth of the frontonasal prominence, and studies on pharyngeal ectoderm have revealed the importance of endoderm in patterning of the pharyngeal arches. The neuroectoderm also has a significant influence on patterning the neural crest into the craniofacial skeleton, as blocking molecular signals from the neuroectoderm leads to craniofacial syndromes such as holoprosencephaly (HPE). Surface ectoderm as a source of craniofacial patterning information Are neural crest cell fates dictated by patterning information inherent in this population of cells, or do neural crest cells respond to signals from their local environment? Studies from our laboratory also suggest that in the frontonasal prominence, the surface ectoderm provides instructive signals that influence neural crest cell fate, even late in development. We identified a region of facial ectoderm that was delineated by the gene expression boundaries of Fibroblast Growth Factor 8 (Fgf8) and Sonic Hedgehog (Shh) (Fig. 5). The junction of the dorsal, Fgf8-positive domain and the ventral, Shh-positive domain coincided with the tip of the upper beak, such that the dorsal (top) surface was derived from Fgf8-expressing epithelium and the ventral (inside) surface arose from Shh-expressing epithelium (Hu et al. 2003). We termed this junction between Fgf8 and Shh the frontonasal ectodermal zone (FEZ). Figure 5Open in figure viewerPowerPoint Transplantation experiments reveal that the facial ectodermal zone (FEZ) is crucial for patterning of the frontonasal primordia. (A) In situ hybridization on a sagittal section of a stage-20 chick embryo; red corresponds to an Shh-expressing domain and green corresponds to an Fgf8-expressing domain. (B,D) Representations of similar lateral sections of donor stage-20 (B) and host stage-25 (D) chick embryos, respectively. (C,E) Trichrome-stained sections of stage-36 control and transplanted embryos, respectively. When the FEZ is transplanted (yellow arrows) from a stage-20 chick donor (B) to a stage-25 chick host (D), an ectopic beak forms by stage 36 (E, black arrowhead). Donor FEZ tissue is indicated by darker red and green colouring, while host FEZ is indicated by lighter red and green colouring. Not only does the FEZ demarcate the initial site of frontonasal outgrowth, it also is responsible for setting up the dorsoventral axis of the upper beak. We showed this by ectopically transplanting the FEZ to a more dorsal position of the frontonasal prominence (Fig. 5B–E). In this new site, the FEZ induced a molecular cascade that re-programmed the fate of neural crest cells at the transplant site (Hu et al. 2003). The net result was duplications of upper beak structures (Hu et al. 2003) (Fig. 5D–E). We found that transplantation of the FEZ to the mandible also induced the duplication of lower beak structures, indicating that the neural crest cells in both locations remained highly responsive to signals emanating from the facial ectoderm (Hu et al. 2003). The dorsoventral polarity of the upper beak is also controlled by the FEZ. When the FEZ graft was inverted, the polarity of the ectopic beak structures was also inverted (Hu et al. 2003). Clearly, some neural crest cell fates can be modulated by external cues from surrounding tissues. When the FEZ graft was transplanted to the second (Hoxa2-expressing) arch, however, the transplant failed to induce a molecular cascade as we had observed earlier, nor was it able to re-pattern the neural crest cells to duplicate any skeletal structures (Hu et al. 2003). Seemingly, these Hoxa2-positive cells disregarded the patterning signals being sent by the transplanted FEZ (Hu et al. 2003). This location-dependent response suggested that the plasticity of a neural crest cell, and its ability to respond to environmental signals, is context dependent (Hu et al. 2003). By extension, these data suggest that removing the influence of Hox genes conferred greater plasticity upon neural crest cells. Pharyngeal endoderm as a source of craniofacial patterning information Some of the first experimental evidence demonstrating that the pharyngeal endoderm is a source of patterning information came from a study that was originally designed to test the role of neural crest cells on patterning of the pharyngeal skeleton. In this study, the neural tube was ablated prior to neural crest migration (Veitch et al. 1999). Despite the absence of neural crest cells post-ablation, the pharyngeal arches were properly organized (Veitch et al. 1999). These data indicated that neither the formation nor the patterning of the pharyngeal arches was absolutely dependent upon neural crest cells. The most likely candidate tissue that controlled patterning in this region of the craniofacial complex was the pharyngeal endoderm. From an evolutionary point of view, pharyngeal ‘perforations’, the predecessors of the pharyngeal clefts, preceded the development of the neural crest as a cell population (Gans & Northcutt, 1983; Northcutt & Gans, 1983). This type of correlation has been interpreted as indicating that the pharyngeal endoderm was probably the initial source of patterning information. Studies from zebrafish and quail/chick chimeras now offer direct experimental evidence that the pharyngeal endoderm can influence that pattern of the lower face. In zebrafish, the van gogh (vgo) mutant shows an absence of pharyngeal segmentation and a failure of the surrounding mesoderm to pattern correctly (Piotrowski & Nusslein-Volhard, 2000). Although hindbrain segmentation proceeds normally, the vgo pharyngeal clefts do not form. Consequently, neural crest cells exiting from the rhombencephalon fuse in the ventral surface because of the lack of pharyngeal pouches; the end result is a lack of skeletal elements in the pharyngeal region (Piotrowski & Nusslein-Volhard, 2000). This phenotype suggests that the segmentation and differentiation of crest cells to form the pharyneal skeleton is primarily determined by endodermal signalling. Using the quail/chick chimeric system, Couly, Le Douarin and their colleagues showed that if regions of pharyngeal endoderm were removed then corresponding regions of the facial skeleton were affrected (Couly et al. 2002). For example, removing one strip of pharyngeal endoderm resulted in the reduction or absence of the nasal capsule and upper beak; when another strip of tissue was removed then Meckel's cartilage was affected, and when a third strip of tissue was removed then the articular, quadrate and proximal portions of Meckel's cartilage were perturbed (Couly et al. 2002). Transplantation of quail endodermal strips of tissue to an ectopic location resulted in supernumerary lower jaws that were malpositioned immediately above the host jaw, and as we had found for facial ectoderm, the rostrocaudal inversion of these grafts resulted in the ectopic lower jaw developing in an inverted position (Couly et al. 2002). Taken together, these data demonstrate that patterning and orientation of the pharyngeal arch skeleton is dependent upon the endoderm. In turn, the pharyngeal endoderm is able to instruct the Hox-expressing neural crest as to the size, morphology and orientation of the pharyngeal skeletal elements. At least some of the patterning information in the pharyngeal endoderm is mediated via Fgf signaling. In the zebrafish acerebellar (ace) mutant the loss of Fgf8 results in deformed pharyngeal pouches and the reduction of the hyoid cartilage (Reifers et al. 1998; Draper et al. 2001; Roehl & Nusslein-Volhard, 2001). More recently, Chuck Kimmel and colleagues showed that disrupting Fg8 signalling results in a complete failure of pouch formation (Crump et al. 2004). Although the pharyngeal endoderm was present in their Fgf mutants, the lateral migration of the endoderm was disorganized and consequently the hyoid and branchial cartilages were truncated, similar to the ace phenotype (Crump et al. 2004). In mice, Fgf8 compound heterozygous (Fgf8neo/–) mutant embryos exhibit hypoplasia of the first and second pharyngeal arches and their associated clefts (Abu-Issa et al. 2002), and although neural crest cells migrate appropriately into the arches, once they arrive they undergo premature programmed cell death (Abu-Issa et al. 2002). Fgfs are not the only molecular signal in the pharyngeal endoderm; endothelin-1 (Edn1) is an intercellular signalling molecule that is expressed in the mesoderm of the pharyngeal arches, as well as in the epithelia of the arches. Both mammalian and teleost data indicate that Edn1 is involved in dorsovental patterning of the arches (Schilling et al. 1996; Miller et al. 2000; Remuzzi et al. 2002; Ozeki et al. 2004), perhaps by establishing a morphogen gradient. Downstream targets of Edn1, such as the bHLH transcription factor Hand2 and the homeobox transcription factor Bapx1, are also involved in dorsoventral patterning in the anterior pharyngeal arches (Miller et al. 2003). Specifically, Hand2 plays a role in specifying the ventral pharyngeal cartilages of the lower jaw and Bapx1 in specifying the jaw joint (Tucker et al. 2004b). Sonic Hedgehog and craniofacial patterning Over the past decade, studies have begun to delve into the precise nature of the environmental cues that influence the neural crest. Many of these studies have implicated Shh as a critical factor in regulating craniofacial morphogenesis. Indeed, Shh seems almost omnipresent, with expression domains in several epithelia presenting themselves at various stages of development. The dynamic expression of Shh in the craniofacial tissues is one indicator of the multiple roles this growth factor plays in modulating normal, as well as abnormal, craniofacial development. Its role in all of these tissues has been examined in mammals (humans and mice), birds, and most recently, zebrafish. Despite the species differences in facial forms, the results have been remarkably consistent. Shh expression is dynamic during embryonic development Shh is expressed in the facial ectoderm, the neuroectoderm and the pharyngeal endoderm at various stages during development. In birds, Shh is sequentially induced in the brain and subsequently in the face (Fig. 6). At stage 15 (Hamburger & Hamilton, 1951), Shh expression in the brain is initially restricted to the ventral diencephalon (Cordero et al. 2004) (Fig. 6B). At stage 17, a new domain of Shh is induced in the ventral telencephalon, which is separated from the diencephalic domain by an Shh-negative optic recess (Cordero et al. 2004) (Fig. 6C). Subsequently, around stage 20, Shh expression is induced in the facial ectoderm (Cordero et al. 2004) (Fig. 6D). These same expression patterns for Shh are appreciated in zebrafish as well (Wada et al. 2005). Figure 6Open in figure viewerPowerPoint Sequence of Shh expression in a developing chick embryo. (A–F) In situ hybridization performed on sagittal sections of chick embryos, where red (pseudocoloured using Photoshop) represents Shh expression as obtained by in-situ hybridization with S35. (A) At stage 10 Shh is expressed in the forebrain (fb) in tissues such as the ventral prosencephalon (vp) and pharyngeal endoderm (pe). (B) At stage 15, the prosencephalon has divided into the telencephalon (te) and the diencephalon (di); at this stage Shh transcripts are localized to the diencephalic neuroectoderm (ne). (C) At stage 17, Shh is now expressed in the telencephalon. (D) Around stage 20, Shh is expressed in a new domain, the facial ectoderm (fe) in addition to being expressed in the diencephalic and telencephalic neuroctoderm. (E) Shh expression remains constant in the neuroectoderm and facial ectoderm from stage 22. (Panel