Survey
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
* Your assessment is very important for improving the workof artificial intelligence, which forms the content of this project
This electronic thesis or dissertation has been downloaded from the King’s Research Portal at https://kclpure.kcl.ac.uk/portal/ Development, morphogenesis and evolution of pharyngeal segmentation in vertebrates Shone, Victoria Louise Awarding institution: King's College London The copyright of this thesis rests with the author and no quotation from it or information derived from it may be published without proper acknowledgement. END USER LICENCE AGREEMENT This work is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International licence. https://creativecommons.org/licenses/by-nc-nd/4.0/ You are free to: Share: to copy, distribute and transmit the work Under the following conditions: Attribution: You must attribute the work in the manner specified by the author (but not in any way that suggests that they endorse you or your use of the work). Non Commercial: You may not use this work for commercial purposes. No Derivative Works - You may not alter, transform, or build upon this work. Any of these conditions can be waived if you receive permission from the author. Your fair dealings and other rights are in no way affected by the above. Take down policy If you believe that this document breaches copyright please contact [email protected] providing details, and we will remove access to the work immediately and investigate your claim. Download date: 16. Jun. 2017 Title page Development, morphogenesis and evolution of pharyngeal segmentation in vertebrates Victoria L. Shone A thesis submitted to King’s College London in part fulfilment of the requirement for the degree of Doctor of Philosophy MRC Centre for Developmental Neurobiology King’s College London September 2013 1 In dedication to my dad 2 Abstract Pharyngeal arches are bulges found on the lateral surface of the head of vertebrate embryos. They are lined externally by ectoderm and internally by endoderm, with a mesenchymal core of neural crest cells and mesoderm. Lateral expansion of pockets of endoderm form pharyngeal pouches at specific locations along the pharynx. Each one of these aligns with invaginating portions of overlying ectoderm to form the anterior and posterior border of each pharyngeal arch. Current studies suggest endoderm plays a prominent role in patterning the arches, but little is known about how this tissue develops and is organised. Investigation of pharyngeal pouch morphogenesis revealed morphological differences between anterior and posterior pouches. These region-based differences are also evident during epithelial interaction at each ectoderm/endoderm interface, where the first interface does not sustain direct contact in contrast to those posteriorly. This results in the fusion and subsequent breakdown of their basement membranes and cell death of the overlying ectoderm. I have revealed that this morphogenetic program for posterior pouch development is conserved in vertebrates and outpocketing of the pouch endoderm represents an early conserved stage of ‘gill’ development. To molecularly characterise the differences between anterior and posterior pharyngeal regions I examined Hox gene expression revealing alignment with specific pouches, thereby separating the pharynx into anterior and posterior regions. Furthermore, the most posterior pouch is demarcated by Hoxb1 expression, so as new pouches form this expression ‘moves’ posteriorly. This dynamic expression pattern is conserved and therefore may underlie how pouch number is controlled within each species. Moreover, a general trend toward a reduction in the number of pharyngeal arches has occurred with vertebrate evolution and I have localised this reduction to the posterior pharyngeal region. By using morphogenetic, molecular and comparative anatomical data I have characterised pharyngeal development and highlighted key differences in anterior and posterior regions reflecting the two main functions of the pharynx: feeding and respiration. As vertebrates transitioned from water to land, their method of respiration was adapted and this is reflected by the reduction in posterior pharyngeal segments in tetrapod species. 3 Acknowledgments My huge thanks go to my supervisor, Anthony Graham, for giving me the opportunity to do this PhD and for the continuous support and encouragement he’s offered the entire time I’ve been doing it. His patience, perseverance, and subtle steering in the right direction is the primary reason I’ve been able to complete experiments, let alone the whole PhD! Anthony’s effortless style of emphasising the bigger picture while never compromising on the details has been inspirational and is a philosophy I will be trying to perfect throughout my career. I also need to thank all members of the Graham lab, past and present. A big thank you goes to Annabelle Scott for taking me under her wing when I was a clueless beginner and for teaching me almost everything I know about molecular biology. She not only helped me but treated me like a friend and equal, and our lively chats in the lab made those difficult early times so much more fun. Thanks also go to Bekah Carr, Aida Blentic and Suba Poopalasundaram for general support and their friendship. Tom Butts requires a big mention – firstly I need to thank him for proof-reading my thesis and for his helpful comments. He’s also not only given me sound experimental advice, but has always been willing to engage in lengthy scientific (and non-scientific) discussions about anything and everything at the drop of a hat. Tom has really helped me to develop as a scientist, but more than that, he’s become a good friend, and for that and all the reasons I listed above I thank him. I also want to thank everyone in the MRC Centre for generating a very special atmosphere to work in. It’s been an amazing place to spend the last 3 years and I’ve made some great friends who have offered lots of support during the ups and downs of PhD student life, often over a drink or two (or three) at the Miller. Last but no means least I want to thank my friends and family for their continued support and encouragement over the years. I want to thank my brother, Paul, my grandparents, Marilyn & Alan and Nanny June, my aunts, uncles and cousins for always being there for me. Thank you also to James for putting up with me for the last two years and being an absolute angel throughout my write-up period. You made it feel so easy and stress-free and without you I’m convinced I 4 would’ve fallen to pieces, so thank you for being my rock. And finally, a big thank you goes to my mum. Without her unwavering support and incredible strength I would not have gotten through the last 17 years or so, and for that I thank her dearly. 5 Table of contents Title page............................................................................................................. 1 Abstract ............................................................................................................... 3 Acknowledgments ............................................................................................... 4 Table of contents ................................................................................................. 6 List of figures .................................................................................................... 11 List of tables ...................................................................................................... 13 Abbreviations .................................................................................................... 14 Chapter 1. Introduction ..................................................................................... 16 1.1 Pharyngeal arch anatomy, development and derivatives ....................... 17 1.2 Patterning the pharyngeal arches ............................................................ 24 1.2.1 Mesoderm in the pharyngeal arches ................................................ 24 1.2.2 Ectoderm in the pharyngeal arches ................................................. 25 1.2.3 Neural crest cells in the pharyngeal arches ..................................... 25 1.2.4 Endoderm in the pharyngeal arches ................................................ 30 1.3 Evolutionary adaptations in pharyngeal arch development of vertebrates.. ....................................................................................................... 36 1.4 The aim of this study .............................................................................. 42 Chapter 2. 2.1 Materials and Methods .................................................................... 44 Solutions ................................................................................................. 44 2.1.1 Common solutions .......................................................................... 44 2.1.2 Bacterial culture media ................................................................... 45 2.1.3 In situ hybridisation solutions ......................................................... 46 2.1.4 Zebrafish immunostaining solutions ............................................... 47 6 2.1.5 Cell death inhibitor solutions .......................................................... 48 2.1.6 CCFSE ectoderm labelling solutions .............................................. 48 2.1.7 Wholemount LacZ staining solutions ............................................. 49 2.2 Embryo Collection ................................................................................. 50 2.2.1 Chick (Gallus gallus) ...................................................................... 50 2.2.2 Dogfish (Scyliorhinus canicula) ..................................................... 50 2.2.3 Mouse (Mus musculus) ................................................................... 50 2.2.4 Zebrafish (Danio rerio) ................................................................... 51 2.2.5 Lamprey (Lampetrus planeri) ......................................................... 51 2.3 Methods .................................................................................................. 52 2.3.1 Riboprobe generation for in situ hybridisation ............................... 52 2.3.2 In situ hybridisation (Chick and Dogfish)....................................... 53 2.3.3 Wholemount immunofluorescence (Chick, Dogfish, Mouse, Lamprey) ....................................................................................................... 56 2.3.4 Zebrafish immunostaining .............................................................. 56 2.3.5 Lysotracker staining for detection of cell death .............................. 61 2.3.6 Cell death inhibition ........................................................................ 61 2.3.7 CCFSE ectoderm labelling.............................................................. 62 2.3.8 Wholemount LacZ staining (mouse)............................................... 62 2.4 Analysing experimental results .............................................................. 63 2.4.1 Sectioning embryos. ........................................................................ 63 2.4.2 Bisecting embryos ........................................................................... 63 2.4.3 Wholemount embryos ..................................................................... 63 Chapter 3. Pharyngeal pouch/cleft interfaces during pharyngeal segmentation… ..................................................................................................... 65 3.1 Introduction ............................................................................................ 65 7 3.2 Results .................................................................................................... 70 3.2.1 Location of the chick ectoderm/endoderm interface during pharyngeal segmentation............................................................................... 70 3.2.2 CCFSE cell lineage tracing to track ectoderm and endoderm cells at their interface during intercalation ................................................................ 87 3.2.3 Lysotracker Red staining reveals bursts of cell death in the ectoderm ........................................................................................................ 92 3.2.4 3.3 Cell death inhibition ........................................................................ 96 Discussion .............................................................................................. 99 3.3.1 Each pharyngeal pouch has a unique morphology, reflecting their development into unique structures .............................................................. 99 3.3.2 Direct interaction of ectoderm and endoderm forms an opening following basement membrane degradation and apoptosis of ectodermal cells…… ..................................................................................................... 102 3.3.3 Apicobasal polarity is not maintained during growth and morphogenesis of the pharyngeal pouch/cleft interface ............................. 106 3.4 Summary .............................................................................................. 108 Chapter 4. Conservation of pharyngeal pouch/cleft interfaces during pharyngeal segmentation across vertebrates ....................................................... 109 4.1 Introduction .......................................................................................... 109 4.2 Results .................................................................................................. 113 4.2.1 Location of the shark ectoderm/endoderm interface during pharyngeal pouch formation ....................................................................... 113 4.2.2 Comparing pharyngeal pouch development at the endoderm/ectoderm interface between amniotes and anamniotes. ............. 122 4.2.3 Pharyngeal pouch development in the lamprey. ........................... 126 4.2.4 Endoderm cell lineage tracing reveals the location of the pharyngeal pouch/cleft interface in transgenic mouse and zebrafish lines. ................... 130 8 4.3 Discussion ............................................................................................ 135 4.3.1 Epithelial interactions at the pharyngeal pouch/cleft interface are conserved in fish and amniotes ................................................................... 135 4.3.2 Cell lineage tracing reveals conservation of pharyngeal pouch out- pocketing ..................................................................................................... 138 4.3.3 Outpocketing of the pharyngeal pouches and operculum development ................................................................................................ 141 4.4 Summary .............................................................................................. 143 Chapter 5. 5.1 Reduction in the number of pharyngeal segments ........................ 144 Introduction .......................................................................................... 144 5.1.1 Hox genes in vertebrate body patterning....................................... 144 5.1.2 Vertebrate evolution has resulted in a loss of pharyngeal arches . 146 5.2 Results .................................................................................................. 149 5.2.1 Cranial nerve innervation identifies where pharyngeal arch reduction has occurred ................................................................................ 149 5.2.2 Hox gene expression in the pharyngeal pouches of amniotes ....... 150 5.2.3 Hox gene expression with the pharyngeal pouches in gnathostomes… ........................................................................................... 155 5.2.4 Transient Hoxb1 expression marks the anatomical border for the posterior pharynx ........................................................................................ 158 5.3 Discussion ............................................................................................ 165 5.3.1 A conserved Hox code aligns with the pharyngeal pouches in the vertebrate pharynx....................................................................................... 165 5.3.2 Dynamic and transient Hoxb1 expression demarcates the posterior pharynx........................................................................................................ 170 5.3.3 Pharyngeal arch reduction has occurred from the posterior pharynx….................................................................................................... 172 9 5.4 Summary .............................................................................................. 176 Chapter 6. Discussion and Conclusions.......................................................... 177 6.1 Endodermal segmentation is conserved ............................................... 178 6.2 A pattern of Hox gene expression in the pharyngeal pouches governs regionalisation of the pharynx......................................................................... 180 6.3 A general trend toward a reduction in the number of pharyngeal arches in vertebrates: how and where does this occur? .............................................. 182 6.3.1 Why did arch reduction occur? ..................................................... 182 6.3.2 Where did arch reduction occur? .................................................. 184 6.4 Concluding remarks ............................................................................. 187 Chapter 7. Bibliography.................................................................................. 189 10 List of figures Figure 1.1. Pharyngeal arch anatomy................................................................... 18 Figure 1.2. Relationship between Hox gene expression in the hindbrain, neural crest streams, pharyngeal arches and pharyngeal pouches of an amniote and anamniote. ............................................................................................................. 28 Figure 1.3. Schematic of pharyngeal evolution in vertebrates ............................. 37 Figure 1.4. Vertebrate phylogeny......................................................................... 40 Figure 2.1. Wholemount immunofluorescence control........................................ 58 Figure 3.1. Morphology and maturation at the ectoderm/endoderm interface of the first pharyngeal pouch ..................................................................................... 74 Figure 3.2. Morphology and maturation at the ectoderm/endoderm interface of the second pharyngeal pouch ................................................................................ 76 Figure 3.3. Morphology and maturation at the ectoderm/endoderm interface of the third pharyngeal pouch .................................................................................... 78 Figure 3.4. Morphology and maturation at the ectoderm/endoderm interface of the fourth pharyngeal pouch ................................................................................. 81 Figure 3.5. A comparison of the morphology of all pharyngeal pouches ............ 84 Figure 3.6. Ectodermal cell lineage tracing using CCFSE................................... 90 Figure 3.7. Lysotracker Red staining reveals cell death in the ectoderm at the pouch interface ...................................................................................................... 94 Figure 3.8. Cell death inhibitors ........................................................................... 97 Figure 4.1. Location of the pharyngeal pouch/cleft interface in stage 19 shark embryos ............................................................................................................... 116 Figure 4.2. Location of the pharyngeal pouch/cleft interface in stage 21 shark embryos ............................................................................................................... 118 11 Figure 4.3. Location of the pharyngeal pouch/cleft interface in stage 22 shark embryos ............................................................................................................... 120 Figure 4.4. Comparison of the pharyngeal pouch/cleft interface during shark and chick pharyngeal development............................................................................ 124 Figure 4.5. Location of pharyngeal pouch/cleft interface in lamprey embryos . 127 Figure 4.6. The pharyngeal pouch/cleft interface of transgenic Sox17 mice and zebrafish .............................................................................................................. 133 Figure 5.1. Cranial nerve innervation of pharyngeal arches reveals region where pharyngeal arch reduction has occurred in vertebrates ....................................... 151 Figure 5.2. Hox gene expression in chick pharyngeal pouches ......................... 156 Figure 5.3. Hox gene expression in dogfish pharyngeal pouches ...................... 160 Figure 5.4. Conserved transient Hoxb1 expression in posterior pharynx of chick and shark ............................................................................................................. 162 Figure 5.5. A conserved pattern of Hox gene expression aligns with the pharyngeal pouches of gnathostomes ................................................................. 168 12 List of tables Table 1.1 Pharyngeal arch and pouch derivatives ................................................ 22 Table 1.2 Genes expressed in pharyngeal endoderm ........................................... 32 Table 2.1 In situ hybridisation probes .................................................................. 55 Table 2.2 Antibodies ............................................................................................ 60 13 Abbreviations A-P Antero-posterior cas casanova CNIX 9th cranial nerve: glossopharyngeal n. CNV 5th cranial nerve: trigeminal n. CNVII 7th cranial nerve: facial n. CNX 10th cranial nerve: vagus n. D-V Dorso-ventral ECM Extracellular matrix FA Focal adhesion NCC Neural crest cell nls neckless PG Paralogous group r rhombomere RA Retinoic acid RALDH Retinaldehyde dehydrogenase RAR Retinoic acid receptor RARE Retinoic acid response element 14 R26R R26 reporter Tg Transgenic TRs Thyroid hormone receptors vgo van gogh 15 Chapter 1. Introduction Embryology is arguably one of the most important ways to assess evolutionary changes. Early events during development will affect what structures form and the morphology that they have, therefore influencing how they function. One of the major alterations leading to the origin of vertebrates was a change in their feeding behaviour, evolving from passive feeders to active predators (Gans and Northcutt, 1983). This shift in behaviour was underpinned by a change in the chordate body structure, with modifications to the anterior part of the existing body plan (Gans and Northcutt, 1983). An important structure involved in this transition is the pharynx, having evolved from being an almost passive organ to comprising a much more complex suite of components, including cartilage and muscles, for a more active function (Gans and Northcutt, 1983). This thesis focuses on the development of the pharyngeal region in vertebrates and how this has been subsequently altered through evolution. Despite the variety of vertebrate species, each one with highly specialized structures, during a particular phase early in embryogenesis all vertebrates look similar. This stage, the ‘phylotypic stage’ (Haeckel, 1910), represents a time point in development when vertebrate features are highly conserved. One of the most striking features at this stage is the presence of several bulges on the lateral surface of the head, termed the pharyngeal arches. All vertebrates bear these structures, with a variation in their number differing between species. The pharyngeal arches are a key feature of vertebrates, eventually developing into the pharynx regardless of how morphologically different this region is across different species. The study of their development and evolution is important for elucidating mechanisms responsible for the emergence of vertebrates, including how they are patterned, and how modification of their developmental program allowed the formation of newly evolved structures, or indeed allowed old structures to develop a new form and/or function. 16 1.1 Pharyngeal arch anatomy, development and derivatives Pharyngeal arches develop around the third week of human development. They form a metameric series along the lateral surface of the embryonic head, within which the muscles, cartilages, nerves and glandular tissue are patterned and formed. Each arch will give rise to distinct structures, for example, the muscles of facial expression originate from the second arch mesoderm. This arch is also innervated by the facial nerve, and in the adult organism all muscles of facial expression are innervated by this nerve or its branches. Therefore, although the pharyngeal arches are not present in the adult, evidence of their metameric origins remain. The arches are formed following interaction between various embryonic cell populations. They are first evident following the segmentation and elongation of pharyngeal endoderm at spatially organized locations along the antero-posterior (A-P) axis. These ‘buds’ of pharyngeal endoderm are known as ‘pharyngeal pouches’, and they expand toward the lateral surface of the embryo to meet invaginating portions of the ectoderm, termed the ‘pharyngeal clefts’. The location where the ectodermal cleft and endodermal pouch meet forms the anterior and posterior border of each pharyngeal arch. Each arch is therefore lined laterally by ectoderm and medially by endoderm, and contains a mesenchymal core composed of both neural crest cells (NCCs) and mesoderm (see for Figure 1.1 schematic). The ectoderm will go on to form the epidermis and neurogenic placodes from which the sensory neurons that innervate the arches will develop. The trigeminal placode is located anterior to the pharyngeal arches and forms neurons of the trigeminal ganglion, which gives rise to projections contributing to the trigeminal nerve (CNV) which innervates the first pharyngeal arch. The epibranchial placodes are located just dorsally and posteriorly to each pharyngeal pouch/cleft interface and these give rise to sensory neurons that innervate the rest of the arches. The most anterior placode will form neurons of the geniculate ganglion and is associated with the first pouch interface. The geniculate ganglion will give rise to projections contributing to the facial nerve (CNVII) which innervates the second pharyngeal arch. The next posterior placode is located near the second pouch interface and will form neurons of the petrosal ganglion, giving 17 Figure 1.1. Pharyngeal arch anatomy (A) Schematic representation showing a lateral view of a chick embryo. The black dotted line marks the coronal plane through which the section in (B) has been taken. (B) This section represents a ventral view of the embryo (anterior at the top). The pharyngeal clefts and pharyngeal pouches have been labelled with orange and pink arrows respectively, and the white holes in each arch represent the aortic arch arteries. The black dotted box around the 3rd pharyngeal pouch demarcates the region which is magnified in (C). (C) The image has been rotated so lateral is at the top. Red – ectoderm; green – endoderm; beige – neural crest cells; blue – mesoderm; OV – otic vesicle; PA – pharyngeal arch; pc – pharyngeal cleft; pp – pharyngeal pouch. 18 ` 19 rise to projections that contribute to the glossopharyngeal nerve (CNIX) that innervates the third arch. The most posterior placode is near the caudal pouches and forms neurons of the nodose ganglion, which gives rise to projections that contribute to the vagus nerve (CNX) innervating the posterior arches (Graham and Begbie, 2000). In amniotes, the endoderm will develop into the epithelial lining of the pharynx as well as the taste buds, lymphatic structures and certain glands, and the first pharyngeal pouch endoderm contributes to the internal ear canal. In humans, the second pharyngeal pouch will form palatine tonsils around the third month of development following proliferation of the endodermal epithelium to form solid buds of tonsillar tissue (Larsen, 1997). The third pharyngeal pouch will form the thymus and inferior parathyroid glands. The thymus gland emerges around week four of human development following proliferation of the ventrally expanded endoderm, and inferior parathyroid glands arise from the dorsal portion of the third pouch endoderm in the fifth week of development (Larsen, 1997). The fourth pouch will form the superior parathyroid glands and ultimobranchial body. The superior parathyroid glands develop in the fifth week as do the ultimobranchial bodies, which arise from a posterior bud coming off the fourth pouch. The ultimobranchial bodies will eventually migrate to the thyroid gland and implant in its dorsal wall before differentiating into parafollicular, or C, cells that produce calcitonin and play a role in calcium regulation (Larsen, 1997). The ventral surface, or floor, of the pharyngeal endoderm will contribute to the thyroid gland around the fourth week of development, following proliferation of a small region of the endoderm located at the foramen caecum (Larsen, 1997). See Table 1.1 for overview of pharyngeal pouch and arch derivatives. The mesenchymal core of each pharyngeal arch is composed of a combination of mesoderm and NCCs. The mesoderm will give rise to the musculature and blood vessels associated with each arch. The musculature associated with the first arch are the muscles of mastication, including masseter, temporalis, lateral and medial pterygoids, as well as some suprahyoid muscles including mylohyoid and the anterior portion of digastric, tensor tympani, associated with constriction of the tympanic membrane in the middle ear, and tensor veli palatini, which tenses the 20 soft palate during feeding. The first arch is innervated by CNV, so these muscles are also innervated by this nerve, and the maxillary artery supplies the first arch. The second arch mesoderm gives rise to the muscles of facial expression including platysma, orbicularis oculi, orbicularis oris, auricularis, risorius, buccinator and fronto-occipitalis, the rest of the suprahyoid muscles (except geniohyoid) including the posterior portion of digastric and stylohyoid, as well as stapedius, the only other muscle found in the middle ear. The second arch is innervated by CNVII, and so all these muscles are innervated by CNVII too, and it is supplied by the stapedial artery. The third arch mesoderm forms only one muscle, stylopharyngeus, a pharyngeal elevator and dilator. This arch, and subsequently muscle, is innervated by CNIX and supplied by the common carotid artery. The fourth arch mesoderm gives rise to the cricothyroid, the only laryngeal tensor, and levator veli palatini which elevates the soft palate during feeding. These muscles are innervated by the vagus nerve (CNX) and supplied by the right subclavian artery. The sixth pharyngeal arch mesoderm gives rise to intrinsic muscles of the larynx which are also innervated by CNX and supplied by the pulmonary arteries (Larsen, 1997). The neural crest will give rise to all cartilaginous and skeletal elements associated with each arch. The neural crest that populates the first arch will differentiate into maxillary cartilage, which will eventually ossify to form both the greater wing of the sphenoid and the incus, and mandibular, or Meckel’s, cartilage that forms the malleus. Also derived from this arch is the maxilla, mandible, zygomatic bone, and the squamous part of the temporal bone, via direct ossification of the arch mesenchyme (Larsen, 1997). Second arch neural crest gives rise to Reichert’s cartilage which is the cartilaginous precursor of the stapes, styloid process, stlyohyoid ligament and the lesser horns and upper rim of the hyoid bone, while the lower rim and greater horns of the hyoid bone are derived from the third arch neural crest. Laryngeal cartilages arise from the fourth and sixth arch neural crest, while some cells migrate through these arches to reach the developing heart. Different species within the vertebrates have varying numbers of pharyngeal arches. For example, in this study I have used the most basal extant vertebrate species, the agnathan (or jawless) lamprey which has 9 pharyngeal arches, basal 21 Table 1.1 Pharyngeal arch and pouch derivatives Main derivatives PA1 Maxilla & mandible, greater wing of sphenoid, zygomatic bone, squamous portion of temporal bone, malleus & incus; muscles of mastication, anterior portion of digastric, mylohyoid, tensor tympani and tensor veli palatini; innervated by CNV Pharyngeal arches PA2 Hyoid bone, styloid process, lesser horns and upper rim of hyoid bone, stapes; muscles of facial expression, posterior portion of digastric, stylohyoid and stapedius; innervated by CNVII PA3 Lower rim and greater horns of hyoid bone; stylopharyngeus; innervated by CNIX PA4 Laryngeal cartilages; cricothyroid, levator veli palatini; innervated by CNX PA6 Laryngeal cartilages; intrinsic muscles of larynx; innervated Pharyngeal pouches by CNX pp1 Internal auditory meatus pp2 Palatine tonsils pp3 Thymus gland and inferior parathyroid glands pp4 Superior parathyroid glands and ultimobranchial bodies 22 gnathostomes the shark and zebrafish which both have 7 arches, and amniotes the chick and mouse, both of which have 5 arches. In the agnathan and basal gnathostome species the arches are numbered 1-9 or 1-7 respectively, but the arch numbering is different in the amniotes. The arches are numbered 1, 2, 3, 4 and 6, with the missing ‘5th arch’ historically representing an arch which either begins to form a vestigial remnant before regressing, or represents the region within the pharynx where the arches have been lost from. The fifth arch does not actually form, and part of the aim of this thesis is to determine where within the pharyngeal region arches have been lost with evolution, the findings of which could challenge this nomenclature (see Chapter 5). There are also differences in the continued development of the pharyngeal region between different species. The pharyngeal arches and pouches will break through to form the gills in agnathan and chondrichthyan species, which are clearly visible on the external surface. Osteichthyan species still form gills from their arches and pouches, but the second arch will expand caudally to cover over, but not obstruct, the posterior arches, appearing as a ‘flap’ on the lateral surface of the head in adult organisms. This flap is called the operculum, and functions both to protect the gills and to help draw water inward. In tetrapod species, the pharyngeal arches are completely obliterated as development progresses. During embryonic development a similar caudal expansion of the second arch is seen as described in osteichthyans, but in amniotes it will continue expanding until it reaches and fuses with the epithelium at the level of the cardiac eminence. The arches then become enclosed within a cavity, the cervical sinus of His, which eventually fuses completely under the control of thyroid signalling (Richardson et al., 2012), internalising the pharynx and forming the smooth outline of the neck. 23 1.2 Patterning the pharyngeal arches Given that pharyngeal arch development involves numerous different embryonic populations derived from all three embryonic germ layers: ectoderm, mesoderm and endoderm, as well as the neural crest, there has been much debate over which of these cell types is the principal organising tissue for regionalisation of the pharyngeal apparatus (Graham et al., 2005). NCCs have historically been implicated as the major component patterning the arches, but some key experiments have revealed the endoderm is probably the principle tissue imparting regionalising cues to the pharyngeal apparatus and I will discuss these two tissues, as well as molecular cues involved in patterning the pharyngeal region in more detail below. However, although no strong evidence suggests mesoderm or ectoderm have patterning capabilities, they are still worth considering when discussing tissues that induce and maintain appropriate patterning cues. 1.2.1 Mesoderm in the pharyngeal arches To test whether mesoderm plays a role in patterning the arches, Trainor and Krumlauf (2000) investigated if it was involved in the activation and maintenance of Hoxb1 expression in second arch NCCs, as Hoxb1 imparts an identity onto the second arch crest. Transplantation of second arch neural crest into a first arch environment resulted in a loss of Hoxb1 expression, but when the crest was transplanted with second arch mesoderm, Hoxb1 expression was maintained. In contrast, when mesoderm alone was transplanted into the first arch, it failed to activate Hoxb1 expression in first arch NCCs. It therefore functions in the maintenance, but not the activation, of Hoxb1 expression in NCCs (Trainor and Krumlauf, 2000). Therefore, although mesoderm is an important tissue to ensure correct patterning signals are maintained in the appropriate region, this tissue has no instructive capability and does not impart any patterning information. 24 1.2.2 Ectoderm in the pharyngeal arches Ectoderm has also been considered as a potential instructive tissue. It has been shown to be involved in pharyngeal development by inducing tooth development. Studies have shown that the ectoderm overlying the first arch instructs underlying crest-derived mesenchyme to develop the capacity for forming teeth (Lumsden, 1988, Tucker and Sharpe, 1999). Furthermore, crest from other arches have also shown the capacity to respond to ectoderm to form teeth, although whether this is restricted to first arch ectoderm or whether it is a general mechanism of ectoderm from any of the arches is not clear. It has also been suggested that the ectoderm undergoes early regionalization to form segments termed ‘ectomeres’, and that each of these will correspond to the later development of the pharyngeal arches (Couly and Le Douarin, 1990, Haworth et al., 2004). However, there is no evidence to show that these are units imparting any developmental organization or that this tissue has any instructive potential. 1.2.3 Neural crest cells in the pharyngeal arches Initially it was believed that NCCs were primarily responsible for patterning the pharyngeal arches and early experiments elucidated whether positional information is indeed imparted by NCCs. Neural crest from the posteriormidbrain and anterior-hindbrain populates the first arch, crest from the midhindbrain populates the second arch and crest from the caudal hindbrain populates the posterior arches. Following transplantation of anterior hindbrain into the place of mid hindbrain, the NCCs migrating out of the transplanted hindbrain streamed into the second arch, as is typical of crest cells migrating from this region (Noden, 1983). However, once the crest reached this arch it induced the generation of skeletal components and associated musculature typical of the first arch (Noden, 1983). This suggested that not only is the neural crest responsible for patterning the arches but that it also obtains its identity within the neural primordium before leaving the neural tube. Lumsden and Keynes (1989) showed that the hindbrain is divided into eight segments, termed rhombomeres (r), from which NCCs will migrate. Fate 25 mapping studies have shown that during development, NCCs migrate from the posterior-midbrain and anterior-hindbrain, or r1 and r2, into the first pharyngeal arch, from the mid-hindbrain, or r4, into the second pharyngeal arch, and from the posterior-hindbrain, or r6 and r7, into the posterior arches (Lumsden et al., 1991). NCCs therefore migrate into the arches in three separate streams. Due to the segregated nature of these streams, it was thought the neural crest carried positional information to pattern the arches based on its organisation within the hindbrain, particularly as this segregation is a conserved feature in vertebrates (Horigome et al., 1999, Lumsden et al., 1991, Schilling and Kimmel, 1994). Kontges and Lumsden (1996) used long-term fate mapping of neural crest cells in quail-chick chimeras to show that the migrating cells within these streams never mix, even when cells from different streams will ultimately contribute to the same structure. These streams are able to retain their separation by expressing the apoptosis-inducing gene Bmp4 in r3 and r5, areas distinctly void of NCC production, therefore playing a key role in the segmentation of the neural crest streams (Graham et al., 1994, Graham et al., 1993). Furthermore, the exit-point of branchiomotor neurons from the hindbrain have been shown to be restricted to even-numbered rhombomeres (Guthrie and Lumsden, 1991), and these correlate with the segmentation of neural crest streams migrating into the pharyngeal arches. For example, the trigeminal nerve root originates from r2 and 3, and exits via r2 to innervate the first pharyngeal arch (Guthrie and Lumsden, 1991), a pattern the most anterior neural crest stream also follows. Similarly, the facial nerve root originates in r4 and 5, exiting r4 to innervate the second arch, the same location the second stream of NCCs migrating out of r4 head toward. Post-otically however CNIX and CNX are not associated with specific rhombomeres, but rather they share a common root which is more widespread along the hindbrain and spinal cord, much like the neural crest migrating out of this region (Kuratani, 1997). Due to the antero-posteriorly patterned origin, route and destination of the neural crest migratory streams, and the fact that Hox genes are well known to be involved in A-P patterning, it has been suggested that a transfer of Hox expression from the hindbrain to migrating crest cells may provide a code for 26 patterning of the pharyngeal arches along their A-P axis (Hunt et al., 1991a, Hunt et al., 1991b). Hoxb1 is expressed in r4 of the hindbrain, and is also expressed in the second pharyngeal arch, suggestive of some transference of positional information (Hunt et al., 1991a). However, Prince and Lumsden (1994) showed that although Hoxa2 is expressed in the hindbrain up to the r1/2 boundary, this expression is not transferred to the NCCs migrating out of r2, suggesting Hox gene expression is established separately in the hindbrain and neural crest. Hoxa2 is a clear indicator of this hypothesis, and Maconochie et al. (1999) identified the cis-components that enable regulation of Hoxa2 independently within NCCs and hindbrain. (See Figure 1.2 for the relationship between Hox gene expression in the hindbrain, neural crest streams, pharyngeal arches and pharyngeal pouches and Chapter 5 for data and discussion on Hox gene expression in the pharyngeal pouches.) Despite no positional information being transferred from hindbrain to neural crest, the Hox genes do play an integral role in the patterning of the crest and its derivatives. When Hoxa2 expression is down-regulated in the second arch, skeletogenic structures of first arch morphology develop (Rijli et al., 1993, Gendron-Maguire et al., 1993, Hunter and Prince, 2002). Similarly, when Hoxa2 is overexpressed in the first pharyngeal arch, skeletal structures typical of the second arch form (Grammatopoulos et al., 2000, Hunter and Prince, 2002, Pasqualetti et al., 2000). However, these transformations can only occur when other tissues express Hoxa2, not just the neural crest cells alone (Grammatopoulos et al., 2000). These experiments therefore show that pharyngeal arches are patterned through the cross-talk between different tissues. In order to test the hypothesis that crest is required for the formation and patterning of the arches, Veitch et al. (1999) ablated the neural tube prior to neural crest cell formation, therefore preventing their migration out of the hindbrain and into the arches. The epithelia of the arches normally express a specific subset of regional markers: Fgf8 is expressed in the anterior endodermal domain of each arch and the overlying ectoderm, while Bmp7 is expressed in the posterior endodermal domain of each arch (Wall and Hogan, 1995, Veitch et al., 1999). Pax1 is expressed in dorsal pouch endoderm, and is therefore a marker 27 Figure 1.2 Relationship between Hox gene expression in the hindbrain, neural crest streams, pharyngeal arches and pharyngeal pouches of an amniote and anamniote. (A) Schematic representation of a chick and (B) shark embryo. In both species, the most anterior limit of Hox gene expression in the hindbrain is Hoxa2 at the r1/r2 border (pink). However, no Hox gene expression is transferred into the arches via the first neural crest stream, and no Hox gene expression is seen in the first pharyngeal pouch (blue). Hoxa2 expression is however transferred into the second pharyngeal arch via the second neural crest stream and is also expressed in the second pharyngeal pouch. Hoxb1 has the next most anterior limit of expression in the hindbrain but is restricted to r4 (green). Hoxb1 expression is also evident in the most posterior pharyngeal pouch in both species, being the fourth pouch in the chick and sixth pouch in the shark. Hoxa3 expression has its anterior limit of expression at the r4/r5 boundary in the hindbrain (purple), and the anterior limit of Hoxb4 is at the r6/r7 boundary (orange). Both of the genes are transferred via the post-otic neural crest stream into the posterior arches, with Hoxa3 having its anterior boundary of expression in the third arch and Hoxb4 in the fourth. Hoxa3 is transiently expressed in the third pharyngeal pouch in the chick but not in the shark, while Hoxb4 is not expressed in any pharyngeal pouches. Adapted from (Couly et al., 1998, Couly et al., 2002). ov: otic vesicle; r: rhombomere; I-VII: pharyngeal arch number; i-vi: pharyngeal pouch number; blue: no Hox gene expression; pink: Hoxa2; green: Hoxb1; purple: Hoxa3; orange: Hoxb4 28 29 for dorso-ventral (D-V) patterning in the arches, and Shh is expressed in posterior second arch endoderm (Muller et al., 1996, Wall and Hogan, 1995, Veitch et al., 1999). The neural crest ablation experiments showed no change to this array of markers, and the pharyngeal arches still formed normally with their sense of identity intact (Veitch et al., 1999). Veitch et al. (1999) therefore demonstrated that in the absence of neural crest pharyngeal arch and pouch formation occurs normally, their dorso-ventral (D-V) and A-P polarity is retained, and so arch patterning is not reliant on NCCs. 1.2.4 Endoderm in the pharyngeal arches Endoderm has been shown to be responsible for the expression of many signals that will induce the generation of the majority, if not all, of the arch derivatives, including epibranchial placodes (Begbie et al., 1999) and all associated glands (Cordier and Haumont, 1980). As well as lining the medial part of each pharyngeal arch, the endoderm forms pharyngeal pouches that make contact with overlying ectoderm and elongate in a D-V direction, separating each arch. The pouches arise along the A-P axis and define the anterior and posterior borders of each arch, and it has been shown that if the pharyngeal pouches do not form, neither do the arches. Of 109 mutations affecting development of the jaw in zebrafish, the van gogh (vgo) mutant is the only one that results in an interruption of pharyngeal arch segmentation and this is as a result of the failure of the endoderm to segment into pharyngeal pouches (Schilling et al., 1996, Piotrowski et al., 1996, Piotrowski and Nusslein-Volhard, 2000). These embryos retained normal hindbrain segmentation and normal migration of NCCs, but when the NCCs reached the region where the arches should be, the crest-derived cartilages fused abnormally (Piotrowski and Nusslein-Volhard, 2000). Similarly, David et al. (2002) used the zebrafish mutant casanova (cas), to show that the depleted endoderm in this mutant resulted in the cartilage within the arches being unable to form. In the vgo mutant tbx1 is disrupted, a gene that is part of the T-box transcription factor family and acts autonomously in the endoderm to pattern 30 neural-crest derived cartilages properly (Piotrowski et al., 2003). cas is a Soxrelated transcription factor which acts downstream of nodal signalling and is important for endoderm formation. It is clear from these experiments that the endoderm is integral for organisation of the pharyngeal region. The endoderm is also the site of expression of many important signals involved in patterning the pharynx (see Table 1.2). Among these signals is the expression of Fgfs, which have been shown to be important for the correct formation of the pharyngeal pouches. Fgf8 mutant mice have been shown to have reduced or absent caudal pharyngeal pouches resulting in abnormal formation of the thymus gland (Abu-Issa et al., 2002, Roehl and Nusslein-Volhard, 2001). Interestingly, although the first arch is affected in Fgf8 mutants, it is the crest-derived cartilaginous structures that do not develop correctly, while the first and second pharyngeal pouches appear to develop normally (Abu-Issa et al., 2002). This difference between anterior and posterior arches and pouches will be a recurring theme throughout this thesis and will be examined in more detail in the following chapters. The relatively mild phenotype of the Fgf8 mutant indicated the functional redundancy of other Fgfs, so Crump et al. (2004) showed in zebrafish that when both fgf8 and fgf3 signalling was repressed by genetic knockdown (fgf8) and morpholino injection (fgf3), pharyngeal pouches completely failed to form, and as a result mandibular cartilages were reduced and hyoid and posterior arch cartilages were mostly absent. Fgf8 mouse mutants have also been shown to phenocopy the human 22q11 deletion syndrome, with abnormalities reminiscent of the vgo zebrafish described earlier, a mutant of tbx1, one of the genes implicated in human 22q11 deletion syndrome (Frank et al., 2002, Piotrowski and Nusslein-Volhard, 2000). Moreover, Fgf8 and Tbx1 have been shown to interact at a genetic level (Vitelli et al., 2002). Bmps are also expressed by endoderm, and have been shown to be crucial for the induction of surrounding ectoderm to form epibranchial placodes (Begbie et al., 1999, Holzschuh et al., 2005). Begbie et al. (1999) showed in the chick that when ectoderm is cultured in isolation no neuronal cells will form, but when endoderm is cultured alongside it they will. Furthermore, when BMP7 was added to isolated ectodermal cultures placode formation occurred, and similarly when ectoderm 31 Table 1.2 Genes expressed in pharyngeal endoderm Gene Function Species found in Tbx1 Endodermal Human (Yagi et al., segmentation 2003); chick (Garg et al., 2001); mouse (Jerome and Papaioannou, 2001); zebrafish (Piotrowski et al., 2003) casanova Endoderm specification Zebrafish (David et al., 2002) Fgf3 and Fgf8 Pharyngeal pouch Zebrafish (Crump et al., formation 2004); mouse (Abu-Issa et al., 2002); chick (Veitch et al., 1999) Bmp7 (chick); bmp2b Induction of Chick (Begbie et al., and bmp5 (zebrafish) epibranchial placode 1999); zebrafish formation (Holzschuh et al., 2005) Development of the Mouse (Manley and ultimobranchial body, Capecchi, 1995, Peters thymus and et al., 1998); chick parathyroid glands (Veitch et al., 1999) Development of Mouse (Xu et al., 2002, thymus, parathyroid Zou et al., 2006); chick and thyroid glands (Ishihara et al., 2008); Pax1 and Pax9 Eya1 zebrafish (Nica et al., 2006) 32 Six1 Thymus development Mouse (Laclef et al., 2003, Zou et al., 2006) Gcm2 Parathyroid gland Rat (Kim et al., 1998); development mouse (Gordon et al., 2001); zebrafish and chick (Okabe and Graham, 2004) Shh Posterior growth of 2nd Chick and zebrafish pharyngeal arch (Richardson et al., 2012) 33 was co-cultured with endoderm with the addition of a BMP7 antagonist, no neuronal cells developed (Begbie et al., 1999). In zebrafish, bmp2b and bmp5 are responsible for inducing epibranchial neurogenesis (Holzschuh et al., 2005). Furthermore, Pax1, Pax9, Eya1, Six1 and Gcm2 are all expressed in the third pouch epithelium and are required for proper thymus gland development in the mouse (Manley and Capecchi, 1995, Wallin et al., 1996, Peters et al., 1998, Su et al., 2001, Laclef et al., 2003, Zou et al., 2006, Xu et al., 2002, Kim et al., 1998, Gordon et al., 2001). The endoderm has also been shown to be directly responsible for imparting patterning information on neural crest-derived structures. Ablation of posterior foregut endoderm results in a reduction of hyoid skeletal elements, and ectopic endoderm grafts result in ectopic formation of hyoid elements according to the positional origin of the graft (Ruhin et al., 2003). Anterior Hox-negative NCCs from the mid-diencephalon to r3 normally contribute to the facial skeleton, and have been shown to be an ‘equivalence group’ (Couly et al., 2002). Following ablation of this region, it was found that a quarter of this entire domain was sufficient to form the whole facial skeleton along with all connective crestderived tissues, indicating the neural crest does not contain information specifying each unit of the facial skeleton, but rather is specified by other tissues (Couly et al., 2002). This tissue was shown to be endoderm following a series of endodermal stripe ablations which resulted in the reduction or absence of crestderived cartilages that normally develop in the endoderm-ablated regions (Couly et al., 2002). Conversely, when ectopic stripes of endoderm were grafted alongside NCCs, the NCCs were induced to form ectopic cartilaginous structures depending on the origin of the endoderm. For example, when anterior quail endoderm was grafted into a chick host around the same region, a supernumerary jaw was formed (Couly et al., 2002). Furthermore, if these stripes of grafted endoderm were rotated, the orientation of the jaw was subsequently affected (Couly et al., 2002). This is reminiscent of observations made in homeotic transformation experiments discussed earlier, where the duplicated jaw structure formed from down-regulation of Hoxa2 in the second pharyngeal arch is a mirrorimage of the normal one (Rijli et al., 1993, Gendron-Maguire et al., 1993), with 34 the pharyngeal pouch acting as the ‘mirror’ and therefore imparting information both for position and orientation of surrounding structures. Similarly, the endoderm has been shown to pattern the crest-derived skeletal structures not only according to their A-P position, but also medio-laterally (Ruhin et al., 2003). 35 1.3 Evolutionary adaptations in pharyngeal arch development of vertebrates Vertebrates arose from within the chordates. Primitive vertebrates are jawless, or known as agnathans, and have varying numbers of pharyngeal arches. For example, extant species of agnathans include the hagfish, with between 6 and 14 pairs of gill slits or pores, and the lamprey with 9 pairs. These species will develop a ‘velum’ from the first pharyngeal arch, which is a series of muscular folds that function in moving water through the mouth and into the pharynx. The jawed vertebrates, or gnathostomes, evolved from agnathan species when the first gill arch developed a hinge to form a pair of upper and lower jaws (see Figure 1.3 for schematic). The general blueprint of pharyngeal arches in gnathostome species, despite the varying number of pharyngeal arches present, is for the first arch to develop into the mandible, the second arch into hyoid structures that support the mandible, and the posterior arches to develop into gill arches while all associated pouches develop into gill slits. The first pharyngeal pouch in agnathans develops into a gill pore. Coincident with the evolution of the first two arches, the first pouch was displaced, moving dorsally and posteriorly to the eye, while its function was also altered to become the spiracle which assists in respiration when the organism has its mouth closed (see Figure 1.3. B). As gnathostomes continued to evolve, the spiracle adapted toward an auditory function with the first pharyngeal cleft and pouch in amniote species contributing to the external and internal auditory canal respectively. Following the transition from water to land, the function of the posterior arches changed dramatically. In water-based species they served as gills for respiration, but tetrapods no longer required gills and so a refinement in the function of each arch ensued resulting in a general trend toward a reduction in the number of pharyngeal arches as well as their internalisation in the fully developed organism (see Figure 1.3. C). To explore evolutionary changes that have arisen in vertebrates, I wanted to look at pharyngeal arch development while focusing on endodermal segmentation in a variety of species representing various phylogenetic groups within the 36 Figure 1.3. Schematic of pharyngeal evolution in vertebrates Schematisation of an agnathan (A), basal gnathostome (B) and amniote (C). Each drawing shows location of pharyngeal arches and pouches and nerves that innervate each with a general trend toward a reduction in the number of arches clearly evident. Note the displacement of the first pouch between agnathan and gnathostomes with the emergence of the jaw. Key: orange – pharyngeal pouches; blue – pharyngeal cartilages in arches; CNV – trigeminal nerve; CNVII – facial nerve; CNIX – glossopharyngeal nerve; CNX – vagus nerve; CNXII – hypoglossal nerve; T – trigeminal ganglion; G- geniculate ganglion; P – petrosal ganglion; N – nodose ganglion; hyoid – hyoid structures that support the jaw; mand – mandible; max – maxilla; pp – pharyngeal pouch; s – spiracle. 37 38 vertebrates. I have therefore decided to study the agnathan lamprey, Lampetrus planeri, basal gnathostomes, the chondrichthyan shark, Scyliorhinus canicula, and the actinopterygian zebrafish, Danio rerio, and amniotes including an avian, the chick, Gallus gallus, and a mammal, the mouse, Mus musculus (see Figure 1.4 for a phylogenetic tree). By comparing and contrasting my results from across these different species I aim to identify how pharyngeal endoderm segmentation occurs, whether the pharynx is regionalised and how, and how modifications have occurred within the pharynx with evolution by defining where arches have been lost. By studying these processes across these different vertebrate species, I can also conclude whether early pharyngeal development is conserved. 39 Figure 1.4. Vertebrate phylogeny This cladogram gives a general overview of the relationships between the vertebrate clades. Sister taxa agnatha and gnathostomes show the emergence of jawless and jawed vertebrates respectively. Jawless vertebrates have between 614 gill pores. All other vertebrates are grouped within the gnathostome clade. Chondrichthyans are vertebrates with a cartilaginous skeleton and between 4-8 exposed gill slits. Osteichthyans have bony skeletons and can be split into actinopterygii and sarcopterygii. Actinopterygii are ray-finned fish with 7 gills that are covered by the operculum. Sarcopterygii are lobe-finned fish, such as the coelacanth, from where the tetrapods arose. Within the tetrapod clade, Amphibia have 5 gills as tadpoles but the operculum covers over and internalises them during metamorphosis into an adult, while birds and mammals have 4 transient pharyngeal pouches which are covered by the opercular flap during embryonic development and therefore never function as gills. Adapted from (Graham and Richardson, 2012). pps – pharyngeal pouches 40 41 1.4 The aim of this study Throughout this thesis I aim to analyse pharyngeal development by focusing on its morphogenesis, regionalisation and evolution within the vertebrates. I have divided my results into three chapters which can be found in Chapters 3, 4 and 5. The aim of Chapter 3 was to catalogue for the first time the morphogenesis of the pharyngeal pouches. Amniotes have five pharyngeal arches and four pouches, each of which will develop into different structures. Using the chick model I investigated the morphogenesis of each pouch to determine whether their early morphology reflected the different structures they will later form. I also wanted to analyse the relationship between the endoderm and ectoderm at the pouch/cleft interface and to uncover whether this interaction is uniform across all the pouches. This revealed distinct differences in the morphogenetic program of anterior and posterior portions of the pharynx, with all posterior pouches following a similar program. This too is reflective of the later structures that are derived from these pouches. In Chapter 4 I wanted to determine whether this morphogenetic program for posterior pouch formation is conserved in other vertebrate species, particularly those that develop gills. Basal gnathostomes the shark and zebrafish represented gill-bearing species in my study, and comparative studies revealed a similar morphogenetic process of the posterior pharyngeal pouches when compared with amniotes, the chick and mouse. This revealed that an early program for ‘gill’ development is conserved in the posterior pouches of vertebrates, and the later remodelling and internalisation of this region in amniotes generates the smooth external surface of the neck in their adult form. The final set of results can be found in Chapter 5. Here I have investigated the instructive potential of the endoderm, particularly the pharyngeal pouches. I have aimed to determine whether they are regionalised and whether they have the potential to impart regionalisation onto surrounding tissues. As Hox genes are responsible for axial regionalisation along the A-P axis of the developing embryo, I analysed Hox gene expression in the pharyngeal pouches. I hypothesised that a particular pattern of Hox gene expression aligns with certain pouches dividing the 42 pharynx into anterior and posterior portions. I also wanted to determine whether this regionalisation was conserved across the vertebrates. A general trend toward a reduction in arch number is evident with more derived vertebrate species, so I aimed to determine where within the pharynx where this variation in arch number is localised among the vertebrates. By using a comparative analysis of species from different phylogenetic groups, I aim to produce a comprehensive study that will contribute to further our understanding not only of the mechanisms behind how the pharyngeal arches and pouches develop, but also how they have evolved as new vertebrate taxa have appeared. By using a combination of morphogenetic, molecular and anatomical data, I aim to refine the details of how the pharynx is regionalised and where modifications have occurred and to speculate on why and how this has happened. 43 Chapter 2. Materials and Methods 2.1 Solutions 2.1.1 Common solutions All reagents are from Sigma, Fisher Scientific or VWR unless otherwise stated. PBS Phosphate Buffered Saline: 1 tablet (Oxoid) in 100ml dH2O, autoclaved. PBST PBS + 0.1% Tween-20 PBSTx PBS + 1% Triton-X 4% PFA (SLS) Paraformaldehyde: 4g PFA dissolved in 100ml PBS at 65˚C and pH adjusted to 7.4, then aliquoted and stored at -20˚C. 50x TAE Stock Tris-Acetate-EDTA Buffer: 242g Tris base (0.8M); 57.1ml glacial acetic acid ; 37.2g EDTA (40mM); topped up to 1L with dH2O. DEPC-H2O Diethylpyrocyanate treated H2O: 0.1% 44 DEPC in dH2O, leave overnight then autoclave. MAB Maleic Acid Buffer: 58.035g Maleic acid (100mM); 43.8g NaCl (150mM); 37.5g NaOH; made up to 5L with dH2O; adjusted pH to 7.5; autoclaved. 20% Gelatin Dissolve 200g gelatin in 1L PBS. Aliquoted and stored at -20˚C. Warmed to 55˚C before embedding. 10% BBR Blocking Reagent (Roche); 10g blocking reagent in 100ml MAB; autoclave; aliquoted and stored at -20˚C. 2.1.2 Bacterial culture media LB Agar Luria-Bertani Agar: 35g in 1L dH2O, autoclaved temperature. and stored at room Working solution was melted, cooled, ampicillin added at 50µl/ml and poured into plates. LB Broth Luria-Bertani Broth: 20.6g LB Broth in 1L dH2O and autoclave. 45 2.1.3 In situ hybridisation solutions 6% H2O2/MeOH 30% H2O2: Made fresh with each use at a 1:5 dilution with 100% MeOH. MABT Add 1% Tween-20 to MAB (after it’s been autoclaved). 10% Deoxycholate Add 50g deoxycholate to 400ml dH2O and stir. Top up to 500ml with dH2O. 10% SDS Sodium Dodecyl Sulphate: 100g SDS; up to 1L with dH2O. 0.5M EDTA pH8 Ethylenediaminetetraacetic acid: 186.1g EDTA; ~50ml 10M NaOH to pH8; up to 1L with dH2O. 2M tris-HCl pH8 242.2g Tris base; up to 1L with dH2O; adjust to pH8 with concentrated HCl. 5M NaCl 292.2g NaCl; up to 1L with dH2O. Detergent Mix 1% IGEPAL; deoxycholate; 1% 50mM SDS; 0.5% tris-HCl pH8; 1mM EDTA; 150mM NaCl; up to 500ml with DEPC-H2O. Sterile filtered before use. Pre-hybridisation Buffer 50% Formamide; 5x SSC pH4.5; 2% SDS; 2% BBR; DEPC-H2O 20x SSC pH4.5 175.3g NaCl; 88.2g sodium citrate; up to 1L with dH2O; pH adjusted to 4.5 with 1M citric acid. Solution X 50% Formamide; 2x SSC pH4.5; 1% 46 SDS; up to 500ml with DEPC-H2O. Blocking solution Make fresh with each use: 2% Blocking reagent (2ml 10% BBR); 20% goat serum (2ml); up to 10ml with MABT. 2M tris-HCl pH9.5 242.2g Tris base; up to 1L with dH2O; adjust to pH9.5 with concentrated HCl. NTMT 100mM NaCl; 100mM Tris-HCl ph9.5; 50mM MgCl2; 1% Tween-20; topped up with DEPC-H2O. Make fresh with each use. NBT/BCIP 4-nitroblue-tetrazolium chloride and 5bromo-4-chloro-3-indolyl-phosphate (Roche): 18.75mg/ml NBT and 9.4mg/ml BCIP in 67% DMSO v/v Colour reaction 1:200 dilution NBT/BCIP in NTMT. 2.1.4 Zebrafish immunostaining solutions PBS-SSDT 10ml: 9.59ml PBS; 100μl 100xBSA; 200μl goat serum; 100μl DMSO; 10μl Triton-X PBS-DT 100ml: 98.9ml PBS; 1ml DMSO; 100μl Triton-X 47 2.1.5 Cell death inhibitor solutions PBS-PenStrep 1:1000 dilution of penicillin/streptomycin in PBS Ink 1:5 dilution of Pelikan Fount India ink (221143) in PBS-PenStrep 20mM z-VAD-fmk stock solution Provided as a stock solution in DMSO. (Promega) 200µM z-VAD-fmk working solution 5µl z-VAD-fmk, 15µl fast green, up to 500µl with PBS-PenStrep 50mM pifithrin-α stock solution Provided as a solid that was dissolved (Calbiochem) in DMSO at 20mg/ml to make stock solution. 500µM pifithrin-α working solution 5µl pifithrin-α, 15µl fast green, up to 500µl with PBS-PenStrep 2.1.6 CCFSE ectoderm labelling solutions PBS-PenStrep As described above Ink As described above 50mM CCFSE stock solution 25mg CCFSE diluted in 798µl anhydrous DMSO, aliquoted and stored at -20°C. 250µM CCFSE working solution 2.5µl CCFSE , 15µl fast green, up to 500µl with PBS-PenStrep 48 2.1.7 Wholemount LacZ staining solutions X-gal 200mg X-gal dimethylformamide, diluted in 4ml aliquoted and stored at -20°C in the dark. Base solution 10mM Tris-HCl pH7.3; 0.005% Nadeoxycholate; 0.01% IGEPAL; 0.25mM K4; 0.25mM K3; 2mM MgCl2; PBS up to 50mls 49 2.2 Embryo Collection 2.2.1 Chick (Gallus gallus) Fertilized brown hen eggs (Henry Stewart & Co Ltd, Lincolnshire) were laid flat on one side and marked to distinguish the superior side where the embryo will develop. They were placed in an incubator at 37˚C and left until the required stage had been reached. Embryos were staged according to Hamburger and Hamilton (HH) stages (Hamburger and Hamilton, 1951), dissected out of their extraembryonic membranes in PBS and fixed in 4% PFA. 2.2.2 Dogfish (Scyliorhinus canicula) Fertilized dogfish egg cases, also known as ‘mermaid’s purses’ were obtained from Bangor University. A tank filled with 20L cold water and 670g Tropic Marin sea salts (dilution according to manufacturer instructions: 2kg sea salts in 60L water) was placed in a 4˚C cold room. The embryos normally grow at sea temperature of around 14˚C, so the water was warmed to this temperature with a heater. When held up to the light the embryo inside is visible and can be staged accordingly (Ballard et al., 1993). As embryos are slow to develop, taking 25 weeks to reach hatching stage, the eggs were checked weekly for embryos at the desired stage. Once they had reached this stage, the egg cases were cut at one end using scissors, and the yolk poured into a glass petri dish. No extraembryonic membranes surround the embryo, so once the embryo had been located the yolk stalk connecting the embryo to the yolk was cut and the embryo fixed in 4% PFA. 2.2.3 Mouse (Mus musculus) Transgenic Sox17-2A-iCre;R26R mice were supplied by Albert Basson, King’s College London. The day the vaginal plug was found was demarcated as E0.5. Dams were sacrificed by CO2 asphyxiation before the uterus was dissected out and placed in cold PBS. Each uterus was separated from the next and the embryo 50 inside dissected free of its extraembryonic membranes. Embryos were staged according to the day collected, i.e. if collected 9 days after discovery of the plug, the stage was E9.5, and immediately used for x-gal staining. 2.2.4 Zebrafish (Danio rerio) Transgenic sox17:GFP zebrafish embryos were kindly donated by Fiona Wardle (King’s College London). Embryos were kept in a Petri dish containing aquarium water at 28.5˚C until they had reached the desired stage as described by Kimmel et al. (1995). Embryos were then freed from their chorions and fixed in 4% PFA for 2 hours at room temperature, then stored in PBST at 4˚C. 2.2.5 Lamprey (Lampetrus planeri) Embryos were kindly donated by Dr Sebastian Shimeld, Oxford University. Embryos had been fixed in 4% PFA and were stored in PBS at 4˚C until ready for use. 51 2.3 Methods 2.3.1 Riboprobe generation for in situ hybridisation 2.3.1.1 Hoxa3 plasmid from an EST clone An EST clone of Gallus gallus Hoxa3 was obtained from Ark Genomics. Colonies were spread over an agar plate and incubated overnight at 37°C. A single colony was picked with a pipette tip and dropped into 5ml LB broth/amp, then left to grow over night while shaking at 37°C. 200µl of this starter culture was then added to 200ml LB broth/amp and left shaking all day at 37°C. Cells were then harvested by centrifuging at 4,200rpm at 4°C for 20 minutes, before extracting the DNA using the QIAGEN HiSpeed Plasmid Maxi Kit according to manufacturer’s instructions. The plasmid was then sequenced by Source Bioscience’s Sanger overnight sequencing service. 2.3.1.2 Plasmid linearisation Each linearisation reaction consisted of the following parameters: 10µg plasmid DNA, 10µl 10X NEB digestion buffer, 5µl restriction enzyme, 1µl BSA (if needed), made up to 100µl with dH2O. This reaction was incubated at 37°C for 2 hours, and linearisation was verified by running the digest on a 1% agarose/TAE gel before purifying the linearised plasmid using the QIAGEN PCR Purification Kit according to manufacturer’s instructions. All probes used and relevant restriction and transcription enzymes are listed in Table 2.1, and all enzymes and buffers are manufactured by NEB. 2.3.1.3 Synthesis of DIG-labelled riboprobes The linearised plasmid was transcribed in the following reaction: 1µg linear plasmid, 2µl 10x transcription buffer, 2µl 10x Dig labelled nucleotide mix, 2µl transcription enzyme, 1.5µl RNase inhibitor (Roche), up to 20µl with dH2O. Both anti-sense and sense strands were transcribed using the appropriate transcription enzyme (see Table 2.1). Anti-sense probes will hybridise to mRNAs present in the embryo that I want to detect, while the sense probes were used as a negative 52 control to detect any non-specific binding. This was incubated at 37°C for 2 hours, then 2µl DNase I and 28µl dH2O was added to the mix and incubated for a further 45 minutes at 37°C. The product was run on a 1% agarose/TAE gel to verify transcription and cleaned using Microspin G-50 columns (GE Healthcare) according to manufacturer’s instructions. The RNA was then added to prehyb buffer at a final concentration of 500ng-1µg/ml and stored at -20°C. 2.3.2 In situ hybridisation (Chick and Dogfish) Day 1 – Embryos fixed in 4% PFA were washed twice for 5 minutes in PBST to remove fixative. At this point they also had their hearts removed, and their forebrains and midbrains cut with Lumsden scissors to prevent probe trapping. They were then dehydrated through graduated steps by washing for 5 minutes (or until embryos settled) in 25% MeOH/PBST, followed by 75% MeOH/PBST, and then 100% MeOH. Embryos were then bleached in 6%H2O2/MeOH for 15 minutes and rinsed twice in MeOH to remove traces of bleach prior to rehydrating in graduated steps of 75% MeOH/PBST for 5 minutes (or until embryos settled), 25% MeOH/PBST and PBST twice for 5 minutes. The embryos were permeabilised with detergent mix twice for 20 minutes before fixing in 4% PFA at room temperature for 20 minutes. They were rinsed then washed for 5 minutes in PBST to remove traces of fixative, then left in pre-hybridisation buffer preheated to 70˚C for 1 hour at 70˚C before replacing with fresh pre-hybridisation buffer containing the 1μg/ml DIG-labelled anti-sense RNA probe, or the sense RNA probe for controls, and left overnight at 70˚C. Day 2 – Solution X was pre-heated to 70˚C to be used for a series of high stringency washes in order to remove any excess probe, non-specific interactions and pre-hybridisation buffer. Embryos were rinsed twice in Solution X, then left for 30 minutes twice in Solution X before being rinsed with MABT 3 times and left in MABT for 30 minutes. Blocking solution was then placed on the embryos for 1-2 hours while rocking at room temperature, before fresh blocking solution was added with anti-DIG antibody at a 1:2000 concentration and left rocking overnight at 4˚C. 53 Day 3 – Embryos were thoroughly washed in MABT by rinsing three times before four 10 minute washes and finally two 30 minute washes. NTMT was then added to the embryos twice for 10 minutes, before the colour reaction was added. This was left to develop in the dark while checking regularly every hour or so until the colour had developed. Embryos were then left for 1-2 days in NTMT to reduce background, washed in PBST for 10 minutes before fixing again in 4% PFA overnight. 54 Table 2.1 In situ hybridisation probes Probe Hoxb1 Species S. Canicula Source Patrick Restriction Transcription enzyme enzyme SpeI Antisense - T7 Sense – SP6 Laurenti Hoxb1 G. gallus Anthony XbaI Sense – T3 Graham Hoxa2 G. gallus Anthony HindIII G. gallus EST clone Antisense - T3 Sense – T7 Graham Hoxa3 Antisense - T7 NotI Antisense – SP6 Sense – T7 Hoxb3 G. gallus Anthony BamHI Sense – T7 Graham Hoxb4 G. gallus Anthony HindIII G. gallus Anthony Graham 55 Antisense - T3 Sense – T7 Graham Hoxb5 Antisense - T3 EcoRI Antisense - T7 Sense – T3 2.3.3 Wholemount immunofluorescence (Chick, Dogfish, Mouse, Lamprey) Previously fixed embryos were washed three times in PBSTx for 30 minutes to remove any remnants of fixative before being washed in a blocking solution of 10% goat serum in PBSTx twice for one hour at room temperature. The relevant primary antibody was diluted in the blocking solution with 0.02% sodium azide to prevent unwanted bacterial growth. This was added to the embryos which were incubated at 4˚C for a period of 1-2 weeks. Embryos were then rinsed in blocking solution and washed three times for one hour in blocking solution to remove all traces of primary antibody before adding the secondary antibody diluted in blocking solution with 0.02% sodium azide. This was incubated at 4˚C for 1-2 weeks before washing the secondary antibody off with three one hour washes in blocking solution. Embryos were then washed three times for 10 minutes in PBSTx, then fixed overnight in 4% PFA. Embryos were then washed 3-4 times in PBS to remove traces of the fixative and the embryos prepared for either sectioning (by embedding in gelatin), wholemount imaging, or were bisected and prepared on slides. Appropriate negative controls will detect background fluorescence and non-specific labelling. Immunoglobulin isotype-matched control antibodies with no known specificity tagged with the same fluorophore as the test antibody will detect any non-specific binding. Similarly, applying a secondary antibody only will also detect any non-specific binding of the fluorophore (see Figure 2.1). Table 2.2 lists each antibody used, its source, dilution, and how the embryo was processed. 2.3.4 Zebrafish immunostaining Transgenic sox17:GFP zebrafish were washed for 5 minutes in dH2O to remove remnants of PBST before permeabilising in cold acetone from the freezer at 20˚C for 7 minutes. They were then washed in dH2O at room temperature for 5 minutes, then in PBST for 5 minutes. Embryos were blocked in PBS-SSDT for 1 hour before applying primary antibody α-GFP (see Table 2.2 for details) diluted in PBS-SSDT and incubating at 4˚C overnight. Embryos were washed in PBS-DT 56 for 3-4 hours, changing the wash around 8 times to remove traces of primary antibody before applying secondary antibody diluted in PBS-SSDT and incubating overnight at 4˚C. Embryos were then washed for 3-4 hours in PBST and the wash changed around 8 times to remove all traces of secondary antibody. Embryos were fixed in 4% PFA for 2 hours at room temperature and washed in PBS 3-4 times before preparing for sectioning. 57 Figure 2.1. Wholemount immunofluorescence control Representative control experiment for wholemount immunofluorescence using stage 17/18 chick embryos. (A, C) Negative control embryos were treated with an AlexaFluor 488 goat anti-mouse IgG coupled secondary antibody only. (B, D) Primary antibody anti-NF-M was applied followed by detection with AlexaFluor 488 goat anti-mouse IgG coupled secondary antibody. Specific staining is only detected in samples treated with both the primary and secondary antibodies. 58 59 Table 2.2 Antibodies Primary antibody Source Dilution Secondary Species used on antibody mouse Zymed 1:10,000 AlexaFluor α-neurofilament 488 goat α- (Rmo270) mouse IgG rabbit Abcam 1:500 AlexaFluor Chick/Lamprey Dogfish/Mouse 488 goat α- α-neuron- mouse IgG specific-β-III tubulin mouse Sigma 1:200 AlexaFluor Chick 488 goat α- α-β-catenin mouse IgG rabbit α-laminin Sigma 1:100 AlexaFluor Chick/Dogfish/Lamprey 568 goat αrabbit IgG mouse α-GFP Roche 1:500 AlexaFluor 488 goat αmouse IgG 60 Zebrafish 2.3.5 Lysotracker staining for detection of cell death Fertilised hen eggs were incubated until the desired stage was reached. Embryos were dissected from their membranes in PBS and transferred into individual wells within a 12-well plate, with each well containing 2ml PBS. 250µl per embryo being used was warmed to 37˚C and Lysotracker Red diluted into the warmed PBS at 1:100. 1.75ml of PBS was carefully removed from each well while taking care not to damage the embryo, and 250µl of the Lysotracker/PBS solution was added. The plate was covered in foil to shield from the light and incubated at 37˚C for 30 minutes. Embryos were then gently rinsed in PBS 4-5 times to remove all medium, then fixed in 4% PFA overnight at 4˚C. Embryos were then rinsed once in PBS and dehydrated in 100% methanol twice to reduce background. Embryos were stored in MeOH at -20˚C until ready for sectioning and imaging. 2.3.6 Cell death inhibition Fertilised hen eggs were incubated until the desired stage was reached. Sticky tape was applied to the superior surface of each egg and 1ml albumen removed using a syringe through a small hole punched into the more flattened end of the egg. A window was cut into the taped surface using curved scissors to expose the embryo. A 0.5mm needle attached to a 5ml syringe was used to inject ink underneath the embryo for easier visualisation and using a Zeiss Stemi SV6 dissecting microscope the embryo was staged. Forceps and a 0.5mm needle were used to peel back the vitelline membrane. 20µl of z-VAD-fmk or pifithrin-α working solutions, or DMSO for control experiments, was pipetted into a 1ml syringe with a 0.5mm needle attached, and the needle carefully guided underneath the opening of the amnion up to the pharyngeal region were the solution was injected directly onto the external surface of the embryo. 0.5-1ml PBS-PenStrep was then pipetted into the egg (away from the solutions so as not to dilute them) and the egg sealed with tape before replacing in the incubator at 37°C for 6, 12 or 24 hours. Embryos were then dissected out of their membranes into PBS and immediately processed for Lysotracker staining as described above. 61 2.3.7 CCFSE ectoderm labelling Fertilised hen eggs were incubated until the desired stage and embryos exposed and visualised in ovo using the same method as described above. 20µl of CCFSE working solution was injected inside the amnion as described above over the pharyngeal region also. PBS-PenStrep was then pipetted into the egg and the egg sealed with sellotape before replacing in the incubator at 37°C for 24 hours. Embryos were then dissected out of their membranes into PBS and staged once again before fixing in 4% PFA overnight and washing in PBS before preparing for sectioning. 2.3.8 Wholemount LacZ staining (mouse) Freshly dissected embryos were placed into individual wells in a 12-well plate, fixed in 4% PFA for 20 minutes on ice, then washed in PBS for 5 minutes three times while the base solution was warmed to 37°C. X-gal was added to the prewarmed base solution in a 1:40 dilution and applied to the embryos in the dark at room temperature until the stain was developed enough (around 1 hour). The reaction was stopped by removing the x-gal solution and washing the embryos in PBS three times for 5 minutes. Embryos were then fixed again in 4% PFA overnight at 4°C. 62 2.4 Analysing experimental results 2.4.1 Sectioning embryos. 20% gelatin was defrosted at 55-65˚C, applied to relevant embryos and allowed to soak for 1-2 hours at 55-65˚C. Embryos were then pipetted into moulding blocks with gelatin, oriented correctly and allowed to set. Gelatin blocks were then fixed in ice cold 4%PFA for at least 4 days when they were washed 3-4 times in PBS until they were sectioned using a Leica VT1000S Vibratome. All embryos were sectioned at 50µm and sectioned mounted under a cover slip with either 90% glycerol/PBS for in situ hybridisation embryos and Fluoroshield™ with DAPI (Sigma) for fluorescent embryos. Sections were viewed and photographed using a Zeiss Axioscope® compound microscope with mounted Zeiss AxioCam MRc5 digital camera, or a Zeiss Axiophot compound microscope with a Zeiss AxioCam HRc digital camera with AxioVision software. Most fluorescent sections were photographed using an Olympus BX61 confocal laser scanning microscope, which was set to photograph at 2µm sections along the Z plane and the sections assembled with the FluoView FV500 software. 2.4.2 Bisecting embryos In situ hybridisation embryos and chick embryos labelled with α-NF-M were bisected in PBS using Lumsden scissors, transferred to a slide and mounted under a coverslip with 90% glycerol/PBS for non-fluorescent and Fluoroshield™ (Sigma) for fluorescent embryos. All embryos were visualised and photographed with a Zeiss Axioskop compound microscope and attached Zeiss AxioCam MRc5 digital camera using AxioVision software. 2.4.3 Wholemount embryos Wholemount embryos were placed within a small well made in ready-set 10% agarose/PBS with a small amount of PBS to inhibit movement while imaging. Fluorescent embryos (dogfish α-β-III tubulin and lamprey α-NF-M) were 63 photographed using a Zeiss Discovery V.20 dissecting microscope with a Zeiss AxioCam MRm digital camera and AxioVision software, and LacZ stained mouse embryos and wholemount chick and dogfish in situ hybridisations were photographed using a Leica M165 FC with attached QImaging QICam Fast 1394 digital camera and Volocity software. 64 Chapter 3. Pharyngeal pouch/cleft interfaces during pharyngeal segmentation 3.1 Introduction Pharyngeal segmentation occurs when the pharyngeal pouches bud off from pharyngeal endoderm along the A-P axis. The lateral most aspect of the pharyngeal pouch makes contact with an invaginating portion of ectoderm, the pharyngeal cleft, before expanding along the D-V axis. The connection that is made between these two epithelia was described in the 1980s using transmission electron and light microscopy to analyse the ‘closing plates’, or ectoderm/endoderm interface, of the chick (Waterman, 1985). The ectoderm and endoderm of the second pouch makes initial contact via a small and focal intercellular junction where the basement membrane becomes discontinuous, and this junction then enlarges prior to interdigitation of the epithelial cells as well as thinning of the interface area until a small perforation appears (Waterman, 1985). However, little is still known about the cellular interactions responsible for pharyngeal segmentation. Although Waterman (1985) describes epithelial interdigitation of the ectoderm and endoderm at the ‘closing plate’, this has not been examined using cell lineage tracing which would reliably reveal how the epithelial cells interact as the pouches continue to develop. In amniotes, the first pharyngeal pouch contributes to the internal auditory canal and is covered at its lateral surface by the tympanic membrane, or ear drum, which is also the medial border of the external auditory canal that develops from the ectodermally derived pharyngeal cleft. This is the only pharyngeal pouch to persist into adulthood with a ‘pouch’ or ‘canal’ morphology, as the rest of the pouches become extensively remodelled and contribute to internalized structures. The second pharyngeal pouch gives rise to the palatine tonsil in humans (Larsen, 1997), although in chick the second pouch does not give rise to any lymphoid tissue or indeed any distinct tissue at all, and instead contributes to the mesenchyme associated with third arch structures (Hamilton and Hinsch, 1957). 65 The third pouch will develop into the thymus and inferior parathyroid glands, and the fourth pouch gives rise to the superior parathyroid glands. An outpocketing of the endoderm in the posterior pharynx gives rise to the ultimobranchial bodies. These epithelial glands remain separate in the chick, but in humans they migrate toward and become associated with the thyroid gland before differentiating into the parafollicular or C-cells to produce calcitonin (Fagman et al., 2006, Fagman and Nilsson, 2010). Non-canonical Wnt signalling has been shown to be important for the formation and maturation of the pharyngeal pouches. In zebrafish wnt11r mutants there is a reduction in the number of pouches and posterior pouch development is delayed, whereas ectopic expression shows a fragmentation of the endoderm into small ‘rosettes’ (as opposed to the normally bilayered pharyngeal endoderm epithelium), indicating a loss of epithelial integrity (Choe et al., 2013). In wnt4a mutants, the normal number of pouches developed but they displayed inappropriate morphology, while ectopic expression led to the formation of pouch-like structures in incorrect domains of the pharyngeal endoderm (Choe et al., 2013). Therefore, wnt11r is crucial for the initiation of pouch formation, while wnt4a is important for directing the maturation of the pouches with a correct morphology (Choe et al., 2013). wnt4a was shown to direct correct pouch morphology by inducing alcama expression in the endoderm, which is responsible for stabilising apical junctions as the pharyngeal pouch matures (Choe et al., 2013). The mechanism by which the pharyngeal pouches elongate along the D-V axis has been previously shown to be caused by a network of actin cables (Quinlan et al., 2004). These are linked via N-cadherin to adherens junctions to form supracellular cables that act as constraints, thereby directing the growth of the pouches along the D-V axis and generating their complex 3D morphology (Quinlan et al., 2004). Although actin cables are responsible for generating and maintaining the slit-like morphology of the pouches, they are not responsible for their actual formation, as when treated with cytocholasin-D to inhibit new actin filament formation, the pouches were distorted in shape but still present (Quinlan et al., 2004). 66 Interestingly, in Sox3 null mice the first and second pharyngeal pouches are almost completely fused, and the main phenotype is a large expansion of the second pharyngeal pouch. This reduces the proximal connection of the second pharyngeal arch to a thin ‘stem’, limiting the number of the neural crest cells that are able to migrate into the arch and so resulting in second arch-associated craniofacial abnormalities (Rizzoti and Lovell-Badge, 2007). What controls the actin cable network formation in the pharyngeal pouches is not fully understood, but perhaps Sox3 or Wnt signalling has some governance over this process. A complex interaction between genetics, signalling molecules and mechanical forces has been suggested for the generation of tissue morphology within various systems via cytoskeletal remodelling, and further experimentation in this area would reveal the process of pharyngeal pouch ‘budding’ from the endoderm and its elongation (Nelson and Gleghorn, 2012). In the Sox3 mouse mutant, only the anterior pouches are affected (Rizzoti and Lovell-Badge, 2007), while in the vgo mutant zebrafish, where tbx1 is mutated and the pharyngeal endoderm does not segment, only the first pharyngeal pouch can be detected (Piotrowski and Nusslein-Volhard, 2000). This highlights a difference between the development of anterior and posterior pouches, and hints at a possibility that mechanisms utilized across all pouches are regulated differently. This is a complex issue that may be determined by the origin of endodermal cells that form discrete regions of the pharyngeal apparatus. Mapping of the pharyngeal endoderm has revealed that endoderm lining the first pharyngeal arch originates from the axial levels of rhombomeres 1 and 2, endoderm lining the second arch originates from the axial level of rhombomeres 4 and 5, and endoderm from the axial level of rhombomeres 5, 6, and 7 contributes to the endoderm lining the second and third pharyngeal arches and the second and third pharyngeal pouches (Veitch, 2000). This implies anterior to posterior patterning specifically in the pharyngeal endoderm forming the pouches, and could be important for differentiating between the developmental processes of anterior and posterior pharyngeal pouches. Endoderm is likely the key instructive tissue responsible for patterning and organising the pharyngeal apparatus (Veitch et al., 1999, Piotrowski and 67 Nusslein-Volhard, 2000), but relatively little is still known about this tissue, particularly in a morphogenetic sense. It is implied by previous experiments that the endoderm gives instruction to the surrounding tissues, i.e. by giving direction to migrating NCCs and directing their proper fusion and development (Piotrowski and Nusslein-Volhard, 2000, Couly et al., 2002, Ruhin et al., 2003). If the endoderm is the principal instructive tissue in this system, it begs the question of how each pharyngeal pouch gains its own identity to direct the development of distinct structures from surrounding tissue? In other words, does the endoderm have autonomous capabilities and therefore is it able to pattern itself, or is it responding to external cues? Each pouch gives rise to distinct derivatives and so presumably has its own molecular signature, and this will be discussed in later chapters. An important factor involved in patterning the endoderm itself is retinoic acid (RA), the generation of which is controlled by the enzyme Raldh2. Inactivation of this gene causes major disruption to the pharyngeal region with huge abnormalities (Niederreither et al., 1999). Mice without Raldh2 form only the first pharyngeal arch, a phenotype also present in the zebrafish mutant neckless (nls), which carries a point mutation in Raldh2 disrupting development of the caudal arches (Begemann et al., 2001). Interestingly, RA signalling was shown to be responsible for posterior pouch formation and partially responsible for second pouch formation, but not at all responsible for first pouch formation (Quinlan et al., 2002), emphasising the differences in formation of the anterior and posterior arches. Quinlan et al. (2002) also showed that Raldh2 is expressed by the lateral mesoderm with a fixed anterior limit at the level of the second pouch, indicating a potential signalling pathway controlling differences between anterior and posterior pharyngeal domains. Despite differences in the way the anterior and posterior pouches develop, the pharyngeal pouches have not been considered separately before. In a study by Waterman (1985), the second pharyngeal pouch ‘closing plate’ (ectoderm/endoderm interface) is analysed and described, and this process is assumed to be identical across all the pouches. Similarly, in a study of differential cell proliferation in perforation of the chick ‘closing plates’, Miller et al. (1993) 68 did not consider pouch number at all, instead combining all of their data and generating an overall result for a common method of ‘closing plate’ rupture across all pharyngeal pouches. Given the different structures that arise from each of the pharyngeal arches and pouches, it would make sense for each pharyngeal pouch to have its own identity and molecular signature, and for its morphology to reflect this. In this study I decided to address an individual analysis of each pharyngeal pouch, and how the morphology of each pouch changes and matures as development proceeds. Another important aspect of pharyngeal arch segmentation which is due to the growth of the pharyngeal pouches is the interaction between the endoderm of the pouches and the ectoderm of the pharyngeal clefts when they meet. This interaction has not been looked at in any specific detail before, but is important for understanding fully how the arches segment and how particular structures are formed. I have used immunofluorescence to highlight the cellular morphology of the ectodermal and endodermal epithelial sheets, revealing not only a distinct difference in the general morphology between each pharyngeal pouch, but also differences in the way the ectoderm and endoderm interact at their interface. I have also used cell lineage tracing to reveal how individual cells of the endoderm and ectoderm interact. This revealed that no intercalation or interdigitation occurs and that the ectoderm and endoderm remain separate at all times, even once the basement membrane separating them has broken down. I have also showed that the ectoderm undergoes short bursts of apoptosis at the same stage as when there is a breakdown in the basement membrane, allowing the endodermal pouch to continue growing out toward and into the external environment. 69 3.2 Results 3.2.1 Location of the chick ectoderm/endoderm interface during pharyngeal segmentation To visualise the relationship of the ectoderm and endoderm at the pharyngeal pouch/cleft interface, I used β-catenin and laminin immunofluorescence to outline cellular morphology and the basement membrane, allowing identification of interactions between these epithelial sheets. The aim of this was to determine whether the sheets intercalate or fuse together, or if some other mechanism is involved. As each pharyngeal pouch develops into unique structures, it is likely that the morphology of each pouch would also be unique, and so I have examined each one separately before comparing and contrasting them. 3.2.1.1 First pharyngeal pouch morphology β-catenin/laminin double immunofluorescence has revealed a distinct morphology of the first pharyngeal pouch and how this alters as the pouch develops. Figure 3.1 shows a series of stages from when the first and second pharyngeal pouches arise at Stage 13 until all four pouches have formed by Stage 19. At stage 13 (n=6) the first two pouches arise simultaneously, and are consequently followed by the second, third and fourth pouch in a temporal fashion along the A-P axis. When the first pouch arises it is a small out-pocketing with close association to the overlying ectoderm. It is flattened in shape and shallow (Figure 3.1. A-D), and displays slightly disorganised apicobasal polarity as evidenced by apical β-catenin and basal laminin labelling, where expression is particular strong at the ectoderm/endoderm interface. However, although much co-localisation is seen here there is more β-catenin labelling along the apical surface and more laminin along the basement membrane, as would be expected (Figure 3.1. B- D). By stage 15 (n=5) the pouch has obtained a more mature shape with a reasonably flattened morphology whereas the cleft has become much deeper (Figure 3.1. EH). The intensity of the laminin labelling is particularly strong across the 70 ectoderm/endoderm interface, although it reveals a clear basement membrane surrounding the epithelium of the rest of the arch. The first pouch is establishing a coherent basement membrane between the ectoderm and endoderm to separate the two tissue layers, which is complete by Stage 17 (n=6) when this disorganisation has disappeared completely (Figure 3.1. I-L). A clear and distinct basement membrane can now be seen between the ectoderm and endoderm tissue layers (Figure 3.1. J, L, Q). By this stage the pouch itself remains flattened with a slight evaginating morphology, and the ectodermal cleft maintains its deep invagination to meet the pouch. At Stage 19 (n=6) the ectoderm and endoderm appear to have separated from one another (Figure 3.1.M). There is clear expression of β-catenin along the apical membranes, as well as laminin along the basement membrane of the pouch itself. However, the basement membrane is disorganised once again at the pharyngeal cleft suggesting a movement of the ectoderm away from the endoderm as opposed to the other way round. Perhaps as the ectoderm moves, some of cells lose proper alignment within the epithelial sheet thereby disrupting their apicobasal polarity to generate a discontinuous basement membrane. Although the ectoderm does appear to be moving away from the endoderm, this occurs in a more ventral location along the D-V axis, and dorsally the epithelia remain in close contact. However, at stage 13 when the pouch first emerges the epithelia are in intimate association with each other throughout their extent, so at later stages even if the epithelia remain in contact dorsally there clearly has been some movement away from one another at certain locations as the pouch has matured. 3.2.1.2 Second pharyngeal pouch morphology The second pharyngeal pouch shows a different, almost opposing, morphology to that of the first pouch. As mentioned earlier the first and second pouches simultaneously bud off from the pharyngeal endoderm at stage 13. The second pouch endoderm is in contact with the overlying ectoderm, and a somewhat disorganised basement membrane is beginning to be laid down between the two (Figure 3.2. A-D). As the pouch continues its development, an intact basement 71 membrane is visible by stage 14 (n=3) at the interface where the epithelia make contact, while the ectoderm has begun to invaginate slightly towards the pouch (Figure 3.2. E- H). Although this basement membrane is mostly continuous, there are a couple of patches bilaterally to the central portion where the basement membrane is not present, suggesting a breakdown in the basement membrane has begun to occur (Figure 3.2. H). At stage 15/16 (n=3 stage 16), a discontinuous basement membrane is visible indicating that the breakdown is spreading from the lateral portions of the centrally located interface (Figure 3.2. I- L, Y). This breakdown in the basement membrane is clearly allowing some kind of mingling between the two epithelial sheets, although the exact nature of this interaction cannot be determined from the data presented here and will be discussed later in the chapter. Stage 17 embryos reveal a single layer of epithelium connecting the second and third pharyngeal arches, indicating either one tissue layer has replaced the other or the cells have become completely intermingled (Figure 3.2. M-P). The intense β-catenin and laminin labelling in the pouch at this stage is perhaps indicative of growth, remodelling, or merging of the two epithelia (Figure 3.2. O, P). This activity appears to have become mostly quiescent by stage 18 (n=4) where a substantial thinning of this interface is evident, particular at the anterior-most portion directly adjacent to the second pharyngeal arch where the beginnings of a small perforation has appeared (Figure 3.2. Q-T, Z). This perforation has expanded by stage 19 such that the entire interface region is now broken through (Figure 3.2. U-X). 3.2.1.3 Third pharyngeal pouch morphology The third pharyngeal pouch appears at stage 14 posterior to the second pharyngeal pouch (Figure 3.3. A). It is first evident as an out-pocketing of the pharyngeal endoderm with intact apicobasal polarity, indicated by clear expression of β-catenin at the apical surface and laminin expression along the basement membrane (Figure 3.3. B-D, U). At this stage the endoderm is evaginating in a postero-lateral direction, while the ectoderm overlying the third 72 pouch at this stage is flat and has its own complete basement membrane extending parallel to the pouch (Figure 3.3. D). The ectoderm and endoderm have made contact by stage 15 at which point the basement membranes appear to have fused or made close and intimate contact so that a single basement membrane can now be seen extending across their interface (Figure 3.3. F-H). The ectoderm is now no longer completely flat and appears to mould around the growing arches and elongating pouches. Similarly, the third pouch is expanding in a more lateral direction (Figure 3.3. E-H). As the pouch continues its development, it grows toward the external surface of the embryo and causes the ectoderm to bulge outward slightly (Figure 3.3. I-L). The basement membrane at stage 17 has begun to break down, with only patches of laminin seen across the interface (Figure 3.3. J, L, V). By stage 18 the basement membrane has almost disappeared and the bulging of the ectoderm externally is much more prominent (Figure 3.3. M-P). The basement membrane has completely disappeared at the interface by stage 19 indicating this region is composed of a single layer of cells, although it is not possible to tell from this data whether these cells are derived from the ectoderm, endoderm, or some kind of interdigitation of both. This layer has also bulged so much that it extends further laterally than the adjacent pharyngeal arches do (Figure 3.3. Q-T). 73 Figure 3.1. Morphology and maturation at the ectoderm/endoderm interface of the first pharyngeal pouch Confocal sections of embryos following immunofluorescence with β-catenin and laminin antibodies. (A) At stage 13 the first two pouches appear simultaneously as small outpockets. The white arrow is pointing at the first pharyngeal pouch, and (B) shows a magnified view of it. Increased levels of both β-catenin (C) and laminin (D) at the first pouch interface suggests a rearrangement of these proteins as the epithelia are growing and remodelling. (E-H) By stage 15 the two epithelia remain in contact and an increase in the amount of laminin at the junction is evident while the pouch itself has slightly deepened. (I-L) At stage 17 the two epithelia are still in contact but have a distinct basement membrane separating them. The box in (J) is magnified in (Q) to show the continuous basement membrane clearly between the two epithelial sheets as shown by DAPI staining of the cell nuclei. At stage 19 (M-P) they have moved away from each other in this plane, with a rearrangement of the basement membrane in the pharyngeal cleft indicated by more intense laminin labelling. (R-T) show schematic representations of the morphology of the pharyngeal pouches depicted throughout this figure. (R) Shows the embryo from a lateral view with the dotted line representing the coronal section respresented in (S). (S) Represents the morphology seen in (A, E, I, M), and the pouch interface is magnified in the schematic representation in (T). (T) Represents the pouch morphology shown in all other images. White arrows in (A, E, I, M) point to the first pouch and subsequent images in the same row show close up images of the same region. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 74 75 Figure 3.2. Morphology and maturation at the ectoderm/endoderm interface of the second pharyngeal pouch Confocal sections of embryos following laminin and β-catenin immunofluorescence reveal the morphology of the second pharyngeal pouch and how this alters as the embryo develops. (A) At Stage 13 the first and second pouches develop at the same time. The white arrow points to the second pouch which is zoomed in on in (B-D). A magnified view of the second pouch reveals a clear separation between the ectoderm and endoderm with a disorganised basement membrane between the two (B-D). At stage 14 a clear and intact basement membrane is seen separating the epithelia which remain in close contact (E-H). This basement membrane is in the process of breaking down by Stage 15/16 (I-L). The boxed region in (I) is magnified in (Y) to more clearly show the discontinuous basement membrane. By Stage 17 no clear basement membrane can be seen, with bright laminin labelling indicating the basement membrane is still in the process of breaking down (M-P). This breakdown appears to have ceased by Stage 18 (Q-T) when the portion of the pouch epithelium directly adjacent to the second arch has thinned to two cells thick. This is more clearly seen in (Z), a magnified view of the boxed region in (R), showing no basement membrane in the pouch interface and DAPI staining of cell nuclei to highlight individual cells. Stage 19 (U-X) embryos show this small perforation has led to the opening of the entire pouch region, allowing the pharyngeal lumen to communicate with the external environment. White arrows point to the second pouch in (A, E, I, M, Q and U) and subsequent pictures in the same row show magnified views of it. (a-c) Schematic representations of the morphology shown throughout this figure. (a) Lateral view of a whole embryo with the dotted line representing the coronal section represented in (b). (b) Representation of the morphology seen in (A, E, I, M, Q, U) with anterior to the left, and (c) represents a magnification of the pouch interface region in (b). (c) Representation of the morphology of the pouch interface region shown in all other images. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 76 77 Figure 3.3. Morphology and maturation at the ectoderm/endoderm interface of the third pharyngeal pouch Confocal images of coronal sections through the pharyngeal arches following βcatenin and laminin immunofluorescence reveals the morphology of the pharyngeal pouches. The third pouch is first evident at stage 14 (A, indicated by white arrow), and close up views of this pouch reveal distinct and separate basement membranes along the basal surface of both the ectoderm and endoderm tissue layers (B-D). The region within the box in (B) is magnified in (U) to clearly show the separate basement membranes. (E-H) The epithelia make contact and fuse by stage 15 when a single basement membrane is evident between the epithelia. (I-L) This basement membrane begins to break down at stage 17, as evident by the spotted laminin labelling. The endoderm also appears to be pushing against the ectoderm, causing the ectoderm to bulge outward. The boxed region in (J) is magnified in (V) to clearly show the degrading basement membrane. (M-P) By stage 18 this bulging is well pronounced with still a few spots of laminin visible between the epithelial layers. (Q-T) At stage 19 the basement membrane has broken down and the epithelial cells directly interact, resulting in a thinning of the ectoderm/endoderm interface to a single layer with no basement membrane, and with a bulged morphology toward the external surface. (W) Schematic representation of a lateral view of an embryo. The dotted line represents the coronal section depicted in (X). (X) Representation of the morphology seen in (A, E, I, M and Q), and (Y) represents a magnified view of the pouch interface region in (X) and all remaining images. White arrows point to the third pouch in all images. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 78 79 3.2.1.4 Fourth pharyngeal pouch morphology The fourth pharyngeal pouch buds off from the anterior portion of the developing fourth arch endoderm around stage 15 (Figure 3.4. A-D). At this stage the epithelium has apicobasal polarity and a clear basement membrane can be visualized (Figure 3.4. B-D). This pouch continues to deepen in a posterior direction, but a change in orientation to move toward the external surface can be seen at stage 17 (Figure 3.4. E- H). The basement membrane is still intact and distinct to the endoderm. However, by stage 19 the endoderm has clearly moved in a more lateral direction to meet the overlying ectoderm when a single basement membrane is visible at the interface between the two epithelia (Figure 3.4. I-L). By this stage the fourth arch is also much more substantial. However, due to the increase in depth of the pharyngeal lumen along the D-V axis (and as a result, an increase in size of the entire embryo), sections must be taken in a more ventral plane in order to visualise the fourth pouch clearly. This is also due to the curvature of this region and the fact that each pharyngeal arch will grow to a different size, for example, the second arch becomes much bigger than any of the others, therefore affecting the overall location of the other arches and consequent curvature of the pharyngeal region. 80 Figure 3.4. Morphology and maturation at the ectoderm/endoderm interface of the fourth pharyngeal pouch Confocal images of coronal sections immunofluorescence through the pharyngeal arches following β-catenin and laminin to reveal the fourth pouch morphology. (A-D) The fourth pouch begins to bud off the anterior endoderm of the developing fourth arch around stage 15 and to grow posteriorly. (E-H) At stage 17 the pouch has deepened in a posterior direction and starts to expand laterally with a clear and intact basement membrane. The endoderm makes contact with the ectoderm at stage 19 after a more lateral expansion and the basement membranes of each epithelial sheet have fused (I-L). (M) Schematic representation showing the lateral view of a chick embryo. The dotted line represents the coronal section represented in (N). (N) Representation of the morphology depicted in (A, E, I). The pouch interface region is magnified in (O) which represents the morphology seen in all other images. White arrows in A, E and I indicate the location of the fourth pouch, and the rest of the images in relevant rows are close up photographs of this region. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 81 82 3.2.1.5 Comparative morphology across all pharyngeal pouches Comparing the morphology of all pouches reveals some important distinctions and similarities which have not been addressed previously. It is apparent that the first pharyngeal pouch is very different from the rest, with an almost inverted morphology when compared with the others. The pouch itself remains relatively flattened during the stages of development examined, and the overlying ectodermal cleft gradually invaginates to meet it (Figure 3.5. I- M). By comparison, in the adjacent second pharyngeal pouch the exact opposite is seen. The pharyngeal cleft remains relatively flat and the pouch evaginates laterally to contact it (Figure 3.5. N-R). This pouch also has a narrow rectangular shape to it which gradually increases in length as development progresses. In comparison, the third pharyngeal pouch has a more triangular morphology (Figure 3.5. S-V) which also gradually gets narrower as it matures until the tip of the triangle appears to bulge out into the external environment (Figure 3.5. V). Inspection of the fourth pharyngeal pouch reveals a dramatic change in morphology and direction of growth as the embryo develops. It begins as a small budding in a posterior direction from the posterior portion of the third pouch (Figure 3.5. W), then as it deepens it begins to grow in a more postero-lateral direction (Figure 3.5. X) until it is entirely laterally oriented and the endoderm of the pouch is making contact with the overlying ectodermal cleft (Figure 3.5. Y). This huge change in orientation of the pouch is a feature not seen in any of the other pharyngeal pouches, and could be due to the growth of the fourth pharyngeal arch causing compression of the pouch as it enlarges. Although the morphology of each of the pharyngeal pouches is very different, there are some similarities in the way they behave. However these similarities are generally restricted to the posterior pouches, as the first pouch develops in a very different way to any of the others. As described earlier, the second pharyngeal pouch begins as a small out-pocketing of the pharyngeal endoderm with a distinct basement membrane separating it from the overlying ectoderm (Figure 3.5. N, O). As its development progresses, a breakdown in this basement membrane is 83 Figure 3.5. A comparison of the morphology of all pharyngeal pouches Confocal images of coronal sections through the pharyngeal arches following βcatenin and laminin immunofluorescence at various developmental stages. (A, B, C) Schematic representation of pharyngeal morphology. (A) Lateral view of a chick embryo. The dotted line represents the plane of the coronal section represented in (B). (B) Representation of morphology seen in (D, E, F, G, H). (C) A magnified view of a pouch/cleft interface representing the morphology seen in images (I-Y). (D-H) An overall view of the embryo at particular developmental stages, and associated pouches at the same stage are found in the same column. Rows show how each pouch changes its morphology as the embryo continues to develop and mature. Distinct differences can be seen in the morphology of each pharyngeal pouch as it develops, and also between the different pouches. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 84 85 evident from patchy laminin labelling (Figure 3.5. P), resulting in intercalation of the epithelial cells, a thinning of the interface, and an eventual perforation (Figure 3.5. Q-R). This is similar to the morphogenesis of the third pharyngeal pouch. The third pouch endoderm and third cleft ectoderm are distinct and separate epithelial sheets with their own basement membranes (Figure 3.5. S), until the endodermal pouch elongates toward the overlying ectoderm, eventually making contact resulting in the fusion of their basement membranes (Figure 3.5. T). A breakdown in this basement membrane then occurs (Figure 3.5. U), allowing interaction of the cells so only one layer is evident (Figure 3.5. V). However, while the second pouch perforates the third pouch does not, and a similarity between the third and fourth pouches is seen with regards to this behaviour. The fourth pouch buds off the posterior portion of the third pouch (Figure 3.5. W), and this pattern of budding is not seen during development of the other pouches. The fourth pouch also undergoes a more dramatic change in growth direction and morphology than the other pouches, as described above. However it still has its own basement membrane, as does the overlying ectoderm at this point (Figure 3.5. X), and when the two epithelia make contact these fuse to leave a single basement membrane separating the two (Figure 3.5. Y). 86 3.2.2 CCFSE cell lineage tracing to track ectoderm and endoderm cells at their interface during intercalation The results above detail the morphology of each individual pouch, as well as their morphogenesis as each pouch continues to mature once the ectoderm and endoderm are in apposition at the pharyngeal pouch/cleft interface. There are significant differences in the morphology of each pouch, and although the anterior pouch behaves differently from the rest, the posterior pouches appear to follow a similar morphogenetic program to one another despite their morphological differences. To determine the extent of these similarities I wanted to look in more detail at the cellular interactions between these two epithelia. The observations above have suggested that once these epithelia meet their basements membranes are closely associated with one another so that they cannot be distinguished, suggesting they may have fused, followed by its subsequent degradation. I therefore wanted to investigate how the cells of the epithelia relate to each other once they are in direct contact. To highlight the relationship between these two cell types I applied CCFSE to the external surface of the embryos to label the ectoderm, allowing lineage tracing of this epithelium and differentiation from the endoderm it interacts with (Richardson et al., 2012). CCFSE (carboxyfluorescein diacetate succinimidyl ester) is a lipid-soluble dye that passively diffuses into cells it comes into contact with. It is not fluorescent until its acetate groups are cleaved by intracellular esterases, whereby it is chemically altered into carboxyfluorescein succinimidyl ester which is highly fluorescent. As it is chemically altered by the cell it cannot diffuse any further, therefore specifically labelling only the cells it has come into contact with. These cells retain the fluorescent dye conjugates during development and mitosis to pass on to daughter cells, thus making it useful as an in vivo cell tracer. Lineage tracing of ectodermal cells will reveal how this epithelium interacts with the endoderm at the pouch/cleft interface, helping to resolve if the ectoderm and endoderm interdigitate or if they remain as separate tissues. Interdigitation of these epithelia has been reported during primary mouth formation in some 87 vertebrate species (Soukup et al., 2013), and so one expectation might be that the two epithelial sheets behave the same way in order to achieve the thinning of the interface. Instead my results have shown that the epithelia always remain as distinct layers until the endoderm eventually displaces the ectoderm. The first pharyngeal pouch has been described above as having a relatively flat morphology, with the overlying ectodermal cleft invaginating to meet it. As development progresses I have shown the two epithelial sheets move further away from each other at certain points along their D-V axis, and this cell lineage tracing confirms that no swapping or sharing of cells has occurred and both sheets remains intact at all times (Figure 3.6. E-H). Cell lineage tracing of the second pouch interface also reveals no intercalation of the ectoderm and endoderm with each other. At stage 17 (n=4) the interface has begun to thin considerably but rather than the expected interdigitation of cells, the epithelial sheets remain distinct from one another with each one appearing to thin out independently by a currently unknown mechanism (Figure 3.6. I, T). The endoderm at the interface disappears or disperses prior to the ectoderm, leaving a thin layer that connects the second and third pharyngeal arches at stage 18 (n=12; Figure 3.6. J). By stage 19 (n=6), this thin layer perforates at its most anterior point adjacent to the expanding second pharyngeal arch (Figure 3.6. K) and the now exposed endoderm of the pouch is in contact with the external environment (Figure 3.6. L, U). The third pharyngeal pouch interface also shows no interdigitation of the epithelial layers. Surprisingly, the endodermal pouch pushes against the overlying ectoderm as it expands toward the external surface causing the ectoderm itself to bulge outwards (Figure 3.6. M, N). By stage 19 it appears to be displacing the ectoderm by growing into it, yet still never mixing with the ectodermal cells (Figure 3.6. O). The pouch then pushes through the ectoderm so it is bulging out of the embryo (Figure 3.6. P, V). This experiment shows for the first time that the endoderm of pharyngeal pouches breaks through the overlying ectoderm to make contact with the external environment. This is different to what is seen at the second pharyngeal pouch interface, as here the endoderm does not bulge through the ectoderm but instead thins significantly before disappearing. The ectoderm 88 then perforates, perhaps because this sheet is no longer stable without a basement membrane to attach to. This perforation is not seen in the third pouch up to the stages that have been examined, although it is possible this perforation may occur later. However at later stages the second arch will grow posteriorly and cover the third pouch, thereby internalising any perforation that may or may not occur (Richardson et al., 2012). The fourth pharyngeal pouch interface shows a similar morphology to that of the third. The pouch is still expanding toward the ectoderm at stage 18 (Figure 3.6. Q), but by stage 19 the epithelia have made contact (Figure 3.6. R). At stage 20 (n=8) it is apparent that the endoderm of the pouch has pushed through the ectoderm in a similar manner to that seen in the third pouch, therefore making contact with the external environment (Figure 3.6. S). 89 Figure 3.6. Ectodermal cell lineage tracing using CCFSE In vivo CCFSE injection on to the external surface of the chick embryo labelled the pharyngeal ectoderm for cell lineage tracing as the embryo continued development for 24 hours. Columns show each pouch at that particular stage of development, and rows show a particular pouch as it matures during development. (A-D) Overview of the morphology of all pouches present in the embryo at that particular developmental stage with white arrows pointing to each pouch and labelled with its number. (E-S) Magnified images of the indicated pouch as seen in the overviews in (A-D). Labelling confirms the first pharyngeal pouch/cleft interface do not exchange or share any cells as labelling is only seen in the overlying ectoderm and ectoderm-derived placodal cells (pink asterisks). (E-H). The second pouch ectoderm and endoderm have a distinct interface when contact is made (I). The region within the box is magnified in (T) to show how the epithelial cells do not mix with ectoderm always remaining on the external edge of the pouch interface. (J) The endoderm of the anterior and posterior portions of the pouch appears to grow into the more lateral aspects of the overlying ectoderm as the ectoderm continues to thin, until it perforates (K) and the two poles of the pouch endoderm can be seen pushing up into the overlying ectoderm (L). The region in the box in (L) is magnified in (U) to show the perforated pouch interface. The third pouch interface also reveals each epithelial sheet retains its own territory (M), with the ectoderm thinning as the endoderm continues to grow ventrally (N) until the endoderm begins to displace the ectoderm (O) and bulges through entirely to make contact with the external environment (P). The region in the box in (P) is magnified in (V) to clearly show the pouch bulging through the ectoderm. The fourth pouch can also be seen to grow toward the ectoderm (Q), make contact with it while not intercalating (R), then also pushes through the ectoderm to contact the external environment (S). (W) Schematic representation of a lateral view of a chick embryo. The dotted line represented the plane of the coronal section depicted in (X). (X) Representation of section through the arches seen in (A-D). (Y) Representation of the magnification of a pouch/cleft interface depicting the morphology in (E-S). 90 91 3.2.3 Lysotracker Red staining reveals bursts of cell death in the ectoderm Following breakdown of their fused basement membranes, the ectoderm and endoderm do not intercalate at their interface yet the ectoderm is somehow displaced at most of these junctures, so I wanted to investigate the mechanism responsible for this phenomenon. As cell death is evident during many epithelial remodelling events, including neural tube closure (Lawson et al., 1999, Weil et al., 1997), eye formation (Silver and Hughes, 1973, Laemle et al., 1999) and digit formation (Garcia-Martinez et al., 1993), I decided to test whether it is involved here by using Lysotracker Red to stain cells undergoing programmed cell death. Lysotracker Red is a fluorescent acidotropic probe, with a fluorophore linked to a weak base that is partially protonated at neutral pH. It is highly selective for acidic environments and accumulates in cellular organelles with a low pH, such as lysosomes, after freely diffusing through the cell membrane. It is also well retained within these compartments (although how it is retained is not fully understood) and so it is ideal for labelling and tracking live cells undergoing cell death. Staining at various developmental stages revealed cell death is associated with the displacement of the ectoderm. At stage 15 (n=3) the epithelia at the second pouch/cleft interface are in apposition with each other, but it is the ectoderm that shows most cell death activity (Figure 3.7. A, D, G). This cell death lasts only for a short burst of time, as by stage 17 cell death is no longer evident at the second pouch/cleft interface (Figure 3.7. B, E). This results in some cells of the ectoderm dying allowing this epithelium to thin, and although some ectodermal cells are still present, the two tissue layers remain distinct (Figure 3.6. I, T). The short bursts of time this cell death is activated correlates with the time period following the breakdown of the basement membrane, which allows direct contact between the epithelial cells. This morphogenetic program appears to initiate a process that results in ectodermal cell death. At stage 19 (n=3) cell death is seen again specifically in the ectoderm of the third pouch/cleft interface (Figure 3.7. C, F, H). A similar morphogenetic program is 92 employed for pharyngeal pouch/cleft interface maturation across the posterior pharyngeal pouches, and ectodermal cell death appears to be dependent on the timing of this epithelial interaction. Once the ectoderm overlying the third pharyngeal pouch has gone, the pouch can continue elongating laterally toward the external environment, as is seen in the CCFSE experiments (Figure 3.6. P, V). These bursts of apoptosis may allow the ectoderm to make way so the endoderm can push through at these specific regions. To investigate whether cell death is required for this morphogenetic event, I employed inhibitors of cell death. 93 Figure 3.7. Lysotracker Red staining reveals cell death in the ectoderm at the pouch interface Embryos were sectioned and imaged with a confocal microscope following Lysotracker Red staining. (A) A stage 15 embryo with the white arrow pointing to the second pharyngeal pouch that is enlarged in (D). It is evident that cell death is occurring mostly in the ectoderm at the interface, which is marked by a white dotted line. This region (boxed) is further magnified in (G) to show dying cells among non-dying cells whose nuclei are stained with DAPI. (B) A stage 17 embryo with the arrows pointing to the first, second and third pharyngeal pouches that are enlarged in (E). At this stage, no apoptosis is evident and cell death appears to have halted at this time point. (C) The pouch morphology at stage 19 with a white arrow pointing to the third pouch enlarged in (F). The dotted line once again demarcates the interface showing the majority of apoptotic cells detected here are also located in the ectoderm. This region is further magnified in (H) where dying cells can be seen located nearest the external edge of the pouch interface and at the anterior-most region closest to the second arch. (I) Schematic representation of a lateral view of a chick embryo. The dotted line represents a coronal section through the embryo as seen in (J). (J) Representation of a section through the arches as seen in images (A-C). (K) Representation of the cleft/pouch interface from (J) magnified to depict morphology seen in images DH). 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 94 95 3.2.4 Cell death inhibition To investigate whether cell death is required for proper morphogenesis and development of the pharyngeal pouches, I inhibited cell death using chemical inhibitors. To increase the probability of inhibition I used two different inhibitors: z-VAD-fmk and pifithrin-α. z-VAD-fmk is a pan-caspase inhibitor. It is cellpermeable and irreversibly binds to the catalytic site of caspase proteases, thereby preventing cleavage of pro-caspase proteins that would activate the caspase pathway. Pifithrin-α inhibits p53. It too is cell permeable and reversibly inhibits p53-dependent transactivation of p53-responsive genes, thereby blocking p53mediated apoptosis. I tested the application of the inhibitors individually and combined but unfortunately obtained no clear effects. When examining the embryos following treatment with the cell death detector, Lysotracker Red, all experimental embryos looked identical to control embryos (see Figure 3.8). 96 Figure 3.8. Cell death inhibitors Stage 15/16 embryos were sectioned and imaged with a compound microscope. Cell death inhibitors z-VAD-fmk and α-pifithrin were applied individually and together to embryos. Following treatment with Lysotracker red no inhibition of cell death was observed in any experiment. Only images from z-VAD-fmk/αpifithrin combined experiments are shown as these provide an adequate representation of all experiments performed; if either one of these chemicals had worked there would have been at least a moderate inhibition compared with controls. (A) Control embryo showing cell death within the pharyngeal arches. Cell death is slightly more pronounced on one side of the embryo (shown to the top). (B) The treated embryo displays similar levels of cell death to the control, also with one side showing more pronounced cell death (shown at the top). (C) A zoomed in view of the white box around the second pouch in (A). Cell death is evident throughout the second and third arches and in the second pouch of the control embryo. (D) A zoomed in view of the white box around the second pouch in (B). Cell death is equally evident in the second and third arches and second pouch of the treated embryo as compared with the control. 97 98 3.3 Discussion The pharyngeal arches are formed when the pharyngeal pouches bud off the pharyngeal endoderm and expand laterally toward the overlying ectoderm. Although the arches and pouches have been described as segmented, this implies they are all similar or even identical in size and shape, but in fact just looking at the arches from the external surface reveals distinct differences between them, and it is well documented that each arch and pouch will develop into distinct structures. As the endoderm is a key instructive tissue in patterning the pharyngeal region, the morphology of each pouch likely underpins how and what it develops, as well as appropriately directing the development of surrounding tissues. This chapter has provided a detailed description of the differences in the morphology of each pouch in amniotes, and links how these differences in pouch morphology reflects the structures that will later develop. 3.3.1 Each pharyngeal pouch has a unique morphology, reflecting their development into unique structures Through the use of immunofluorescence, I have shown that each pharyngeal pouch has a unique morphology. β-catenin is part of the protein complex constituting adherens junctions found at cell-cell junctions of epithelial tissue. It acts at the apical membrane to attach the transmembrane cadherin protein to the actin cytoskeleton, therefore making it a useful marker for outlining cellular morphology within a tissue. Laminin is a protein found in the basement membrane of epithelial tissues. It is connected to epithelial cells via integrin receptors and other plasma membrane molecules. Using antibodies against both of these proteins allows visualisation of the apical and basal surfaces of the epithelial sheets revealing how they interact with one another at their interface. The first pharyngeal pouch forms the internal auditory canal proximally and tympanic cavity distally and the first pharyngeal cleft forms the external auditory canal. The proximal part of the external auditory canal contacts the distal portion 99 of the tympanic cavity, although they are kept separated by the mesenchymal tympanic membrane, or eardum. This is responsible for converting soundwaves into mechanical oscillations that are translated by the adjoining ear ossicles (Grevellec and Tucker, 2010). The early morphology of the first pharyngeal pouch and cleft reflects its later morphology and function. As described earlier, the cleft invaginates to meet the barely evaginating pouch, reminiscent of an external ear canal. The ectoderm and endoderm begin their development in this region by contacting each other at the first pouch/cleft interface, before proceeding to separate by the intervention of mesenchymal cells that will eventually form the eardrum (Grevellec and Tucker, 2010). Whether the mesenchymal cells migrate here as a result of the epithelia separating or whether they are the cause of this separation is not clear. However, no such movement has been reported before and it has always been presumed the ectoderm and endoderm migrate toward each other but never make contact because of the intervening mesenchyme (Grevellec and Tucker, 2010). However, this initial contact at the interface and eventual separation is very different to what is seen at the other pharyngeal pouch interfaces. The second pharyngeal pouch has an opposing morphology to that of the first, where the ectodermal cleft barely invaginates and the pouch itself evaginates greatly to meet it. This pouch also begins its development in contact with the overlying ectoderm, and a basement membrane can be seen to separate them. However over time, this basement membrane is broken down and the ectoderm and endoderm are allowed to interact. The exact nature of this interaction is surprising in that the epithelial sheets never interdigitate or mix in any way, always remaining separate. Following this interaction some cell death is seen mostly in the ectoderm layer, which may be responsible for thinning of the interface, followed by perforation at later stages forming an opening where the interface once was. When compared with the third pouch morphology, the second pharyngeal pouch has a rectangular shape whereas the third pouch has a more triangular shape. As the antero-ventral portion of the third pouch gives rise to the thymus and the postero-dorsal aspect to the inferior parathyroid glands in mammals (Grevellec 100 and Tucker, 2010, Larsen, 1997), it appears the pouch is composed of two distinct sides that will eventually develop into two distinct structures. In contrast to the first and second pouches, the third pouch is not in contact with the overlying ectoderm when it first buds off the posterior pharyngeal endoderm, and the immunofluorescence reveals the two epithelia are distinct sheets with apicobasal polarity and their own basement membranes. As the pouch endoderm continues to elongate, it makes contact with the overlying ectoderm and their basement membranes fuse together, at which stage the epithelial interaction becomes more similar to that seen at the second pharyngeal pouch. A breakdown of the basement membrane allows interaction of the epithelial cells resulting in a thinning of the interface, leaving a single epithelial sheet. This layer appears to be bulging out towards the external environment and grows beyond the border of the adjacent pharyngeal arches. Unlike the second pouch though, no external opening is observed at the stages examined. As the pharyngeal pouches arise along the A-P axis, the anterior pouches are effectively more mature than those posteriorly at any given time. Therefore later examination of the third pouch interface may reveal a perforation, although the caudal expansion of the second pharyngeal arch would subsequently enclose this area. Alternatively, perhaps the internalisation of this structure by the enlarged second pharyngeal arch means the third pouch does not have time to break through. It would be interesting to dissect away the second pharyngeal arch prior to its caudal expansion and observe what happens to this posterior pouch, but this is technically tricky. An alternative experiment using cyclopamine beads to block the proliferation-inducing signal of Shh to specifically inhibit second arch growth and allowing the embryo to develop until pharyngeal maturity at around stage 2930 would be an interesting future experiment (Richardson et al., 2012). If the embryo had pharyngeal fistulas, it will be apparent that these posterior pouches do indeed perforate. The fourth pharyngeal pouch differs again from the first, second or third pouch in its origins by budding off the posterior aspect of the third pouch, before expanding posteriorly and eventually laterally to form the posterior border the fourth pharyngeal arch. The fourth pouch begins as a distinct epithelial sheet, 101 similar to the third pouch, and grows toward the overlying ectoderm to make contact where the two layers will share a basement membrane. This basement membrane is intact in the latest staged embryo examined, but it would not be unreasonable to hypothesise that as this interface matures, the basement membrane will degrade and a fusion of the epithelia will occur in a similar manner to that seen during second and third pouch maturation. The fourth pouch also has a different shape to that seen in the second or third with a U-shaped morphology. This pouch will give rise to the superior parathyroid glands, but it also gives off a posterior bud similar to how the fourth pouch arose from the posterior aspect of the third arch endoderm, and this bud or controversial ‘fifth pouch’ (Dudley, 1942, Hilfer and Brown, 1984) gives rise to the ultimobranchial bodies. 3.3.2 Direct interaction of ectoderm and endoderm forms an opening following basement membrane degradation and apoptosis of ectodermal cells Direct contact between the ectoderm and endoderm is not commonly seen in the developing embryo, although it does occur during pharyngeal pouch formation and elongation, as well as during the development of other structures such as the primary mouth (Soukup et al., 2013). Although this has been reported previously, a comprehensive analysis of each individual pouch and the exact nature of this interaction has not been fully elucidated (Cordier and Haumont, 1980, Piotrowski and Nusslein-Volhard, 2000, Xu et al., 2002). The first pouch behaves differently to the others. It is in contact with the overlying ectoderm as soon as it buds off the pharyngeal endoderm and then the epithelia move away from each other, which is not seen in any other pouch. This pouch and cleft will go on to form the internal and external auditory canals respectively, with the distal end of the pouch and proximal end of the cleft contributing to the epithelium surrounding the mesenchyme-derived centre of the eardrum (Grevellec and Tucker, 2010), which acts as a boundary separating the two. It is clear that whatever governs this 102 process is not responsible for regulating the development of the other pouches, but it would be interesting to inhibit this mesenchymal intervention and see if the pouch develops in a similar way to the others. The second, third and fourth pouches all have interfaces where the ectoderm and endoderm make direct contact resulting in the contact of their basement membranes. The basement membrane is composed of two layers: the lamina fibroreticularis and basal lamina. The basal lamina itself is composed of two layers: the lamina lucida and lamina densa. These layers are tightly adhered to one another, with the basal lamina being attached to the lamina fibroreticularis by reticular fibres. The basal lamina portion of the basement membrane contains laminin proteins. When the basement membranes from the two epithelia make contact with one another, they may either directly fuse resulting in a single basement membrane or they may intimately associate with one another so that they cannot be seen to be separate membranes from my images. Following this contact, the basement membrane degrades, allowing interaction of the epithelia which may induce localised cell death in the ectoderm tissue layer at the interface. The basement membrane at the second pouch/cleft interface breaks down at stages 14-15 (see Figure 3.2. E-L, Y), and cell death is seen at stage 15 (see Figure 3.7. A, D). No cell death is evident at stage 17, but following the breakdown of the basement membrane at the third pouch/cleft interface in stage 17-18 embryos (see Figure 3.3. I-P, Y), localised cell death is again evident here in the ectoderm by stage 19 (see Figure 3.7. C, F, H). This focal cell death appears to be initiated following the breakdown of the basement membrane, and the interface thins to a single layer. This layer eventually breaks through at the second pouch/cleft interface and will potentially break-through in the third and fourth pouch/cleft interfaces also. The mechanisms that control this process are unknown, although as cell death is seen in the ectodermal layer it is possible the endoderm is emitting a cue causing the basement membrane to degrade. It is also possible however that the ectodermal cells undergo cell death following the breakdown of the basement membrane because they no longer have attachment to the extracellular matrix (ECM). The ECM consists of proteins and polysaccharides to support and anchor the cells it surrounds. The basement 103 membrane is part of the ECM and nearby cells attach to it, providing signals to prevent apoptosis, or ‘anoikis’ (Meredith et al., 1993, Frisch and Francis, 1994). ‘Anoikis’ is the term given to the apoptosis program when not induced by any specific molecular signal, but rather from a lack of attachment to the ECM (Frisch, 1994). Receptors of the integrin family link the ECM to the actin cytoskeleton of cells and are essential for preventing the apoptotic program within it (Gilmore, 2005). This cause of cell death is seen during neural tube closure, where extensive apoptosis during this process is a consequence of remodelling following neural fold fusion rather than a cause of it, and inhibition of apoptosis in this region shows no effect on neural tube closure (Massa et al., 2009a, Massa et al., 2009b). I also tested whether inhibiting cell death would have an effect on the development of the pharyngeal pouches. Unfortunately these experiments were unsuccessful; when comparing the results of experimental embryos with controls there was no difference (Figure 3.8). I identified known regions of cell death, such as the lens and neural tube, where cell death should have also been inhibited following application of the inhibitors to the head region but in this instance were not. This was likely due to my method of application, as my experiments had to be performed on chick embryos in ovo. When applying the inhibitors I added fast green to their solution so I was able to visualise where the solution was in relation to the embryo. At the time of application, the solution was injected within the amnion covering the region where the chick head was, but following examination after a further 24 hours of incubation, the solution had diluted out and very little fast green was visible in its intended region. To try and combat this I incubated the embryos for shorter lengths of time, trying 12 hours and 6 hours, but to no avail. Another method that may prove successful would be to culture the chick embryos ex ovo and apply the inhibitors to the culture medium. Few systems have been described for whole-chick culturing, with limited success reported for most of these. These systems have also generally been set up for ex ovo imaging, electroporation, microinjection and microsurgery experiments (Endo, 2012, Yalcin et al., 2010, Chapman et al., 2001). The chicks are also cultured on agar or 104 in albumen (Chapman et al., 2001, Yalcin et al., 2010), so adding inhibitors to the culture media may not work as they may not be able to diffuse adequately within viscous substances. Contact between ectoderm and endoderm is seen in a few other instances during development, and in each case appears to trigger a signalling cascade resulting in perforation to create an opening. During nasal cavity formation, the ectodermal nasal pit deepens and invaginates until it meets the underlying endoderm, whereupon these epithelia fuse to form the oronasal membrane. This will then rupture via apoptosis, allowing free passage between the oral cavity and the external environment via the nasal cavity, or nostril (Cole and Ross, 2001, Larsen, 1997). This contact is also seen during mouth and anus formation, yet very little research has been conducted into these processes, with mouth formation being the most studied area. Dickinson and Sive (2006) proposed a model for primary mouth formation in Xenopus embryos whereby an initial breakdown of the basement membrane separating the ectoderm from the endoderm allows direct contact of these epithelia, followed by ectodermal invagination and apoptosis of ectodermal cells while the epithelia intercalate. The combination of these events results in a ‘thinning’ of the primary mouth area followed by perforation. The exact mechanism for perforation is still unknown, although Dickinson and Sive (2006) hypothesize it could be due to either tension generated by the growth of surrounding facial regions, or the loss of cell adhesion. In the mouse a similar finding of apoptotic bursts were recorded hours prior to rupture of the primary mouth (Poelmann et al., 1985), and although studies in other vertebrate species have not detected apoptosis in this region, they did report the presence of lysosomes which could be a sign of cell death (Waterman, 1977, Waterman and Schoenwolf, 1980, Watanabe et al., 1984). Cell death has been reported elsewhere to be responsible for the ‘thinning’ of tissues, such as during the formation of the hindbrain roofplate in chick (Lawson et al., 1999) and during formation of the urethral and anal opening (Qi et al., 2000a, Qi et al., 2000b, Qi et al., 2000c). While investigating cloacal septation in rats, Qi et al. (2000a,b,c) uncovered direct interaction between ectoderm and endoderm at the anal and urethral membranes resulting in a marked thinning of 105 the interface and apoptosis of the ectoderm. This was described as a gradual process with a few gaps appearing in the epithelium, before the stress of continued proliferation of the surrounding tissue caused the membrane to rupture. On close inspection of the second pouch Lysotracker staining (Figure 3.7. D), it is apparent that only the cells nearest the more anterior portion of the interface where the perforation will occur have undergone apoptosis, and presumably this perforation is assisted by the great proliferation and expansion of the second pharyngeal arch. 3.3.3 Apicobasal polarity is not maintained during growth and morphogenesis of the pharyngeal pouch/cleft interface Apicobasal polarity of epithelial sheets ensures the integrity of the sheet is maintained while each cell retains contact with its neighbour. The connection between a cell and the extracellular matrix is found at the basal end via a focal adhesion (FA). This FA consists of a transmembrane protein, in this instance an integrin, which binds to proteins in the basement membrane such as laminin, forming a connection to the intracellular actin cytoskeleton (Nelson and Gleghorn, 2012). Apically, tight junctions maintain cell-cell contact and prevent unwanted molecules from crossing the sheet. Just basal to these are adherens junctions, which also maintain cell-cell contact via transmembrane proteins, cadherins, which homophilically bind to the corresponding cadherin on the cell membrane of its neighbour. These cadherins are attached via various proteins, including β-catenin, to the actin cytoskeleton of the cell, thereby having the capacity to cause apical constriction of the cell. ‘Budding’ occurs during many morphogenetic process, including lung formation, and is caused by coordinated apical constriction of epithelial cells resulting in their cellular morphology becoming more wedge-like, causing a U-shaped evagination (Metzger et al., 2008, Martin et al., 2010, Sawyer et al., 2010, Nelson and Gleghorn, 2012). This mechanism is also responsible for the onset of pharyngeal pouch elongation (Quinlan et al., 2004). 106 BMP signalling has been shown to modulate epithelial polarity during neural tube closure in chick (Eom et al., 2011), and given the expression of BMPs in pharyngeal pouch endoderm (Veitch et al., 1999, Graham, 2001), apicobasal polarity in this region may well be governed by the endoderm, supporting it as being the instructive tissue driving pharyngeal arch segmentation and patterning. However, I have revealed several instances throughout pharyngeal pouch development where both β-catenin and laminin proteins are labelled concurrently, and this does not reflect true apicobasal polarity. There have been reports of epithelial sheets temporarily losing their apicobasal polarity during times of morphogenesis to allow growth and remodelling (Ewald et al., 2008, Bryant and Mostov, 2008). Ewald et al. (2008) found this was the case when examining mammary branch morphogenesis and coined the term ‘morphogenetically active epithelial state’ to describe the state of the epithelium during a period of morphogenesis. Although in this particular case the epithelial cells which transiently lost their polarity were part of a multilayered sheet, they suggest this mechanism is seen across other areas of epithelial remodelling and growth making it possible that this is what is happening during remodelling of the pouch/cleft interface. 107 3.4 Summary Pharyngeal pouches bud off the pharyngeal endoderm and elongate along the DV axis to form the anterior and posterior borders of each pharyngeal arch, thereby segmenting them, with each pouch subsequently giving rise to specific structures. The differing morphology of each pouch shown in this chapter is a clear reflection of the different structures each pouch will go on to develop. The interaction of the pouch endoderm and overlying ectoderm has not been accurately examined before. I have shown that their interaction is different across the pouches, with the first pouch epithelia being separated by mesenchyme whereas the rest of the pouch interfaces behave differently. Generally, contact between the ectoderm and endoderm results in a breakdown of the intervening basement membrane, followed by a ‘thinning’ of the interface. This ‘thinning’ was not caused by intercalation of the epithelia and cell death was detected mostly in the ectoderm, providing a potential mechanism for this ‘thinning’. At the second pouch interface this results in an eventual perforation, possibly partially caused by tension exerted from the caudally expanding second pharyngeal arch. The third pharyngeal pouch expands toward the external surface of the embryo following apoptosis of the overlying ectoderm, and the fourth pouch interface shows similar interactions although was not examined late enough to determine if it would follow suit. These results answer basic yet important questions regarding the development of the pharyngeal apparatus, and provide a platform for the examination of similar mechanisms seen during development of this region across other vertebrate species. 108 Chapter 4. Conservation of pharyngeal pouch/cleft interfaces during pharyngeal segmentation across vertebrates 4.1 Introduction During early stages of embryonic development, different vertebrate species bear a striking resemblance to one another at a particular phase, termed the ‘phylotypic stage’ (Slack et al., 1993, Duboule, 1994, Hall, 1997). The pharyngeal arches are a key feature present at this stage, forming a segmented series on the lateral surface of the head, although they vary in their number between different species. Despite the similarity in the morphology of these structures across vertebrate embryos at this stage, each species has a very different morphology when fully developed, for example, fish pharyngeal arches develop in to gills while mammals form a neck. Basal vertebrates are jawless (agnathan), and jawed vertebrates (gnathostomes) evolved from this group (see Figure 1.4). Extant agnatha are represented by the hagfish and lamprey. Hagfish have between 6-14 arches while lamprey have 9, but fossils of extinct agnathans, the ostracoderms, revealed jawless species had up to 30 pairs of gill arches (Janvier and Arsenault, 2007). The first pharyngeal arch in hagfish and lamprey contributes to the velum, a structure found in their oral region that has adapted to filter feeding and parasitism, and their lower lip (Kuratani et al., 2001, Shigetani et al., 2002), while the gnathostome first arch will develop into a jaw. Chondrichthyans and osteichthyans represent basal gnathostomes. Extant chondrichthyans are characterized by species with a cartilaginous skeleton, although some extinct agnathan, primitive gnathostome (placoderms) and chondrichthyan species had bony skeletons, so extant chondrichthyans are derived and have secondarily lost their bony skeleton (Donoghue and Sansom, 2002, Eames et al., 2007). Most chondrichthyans have 7 pharyngeal arches, although this number varies between 6 and 9 arches. However the number of 109 arches in osteichthyans is stable, so the variability within chondrichthyans is probably specific to this group. Osteichthyans have 7 pharyngeal arches, and a change in the external appearance of the arches once fully formed is apparent between these clades. While chondrichthyans retain open gill slits clearly visible on their external surface, the osteichthyan gill apparatus is covered, but not enclosed, by the posterior growth of the second gill arch, known as the operculum. Osteichthyans are comprised of two groups, the actinopterygians and sarcopterygians (Figure 1.4). Basal sarcopterygians, such as the coelacanth and lungfish, retain gills and an operculum, but more derived sarcopterygians, the tetrapods, do not and their pharyngeal arch number is reduced to 5, all of which are internalized during development, except the first arch which will develop the jaw. It is within this clade that the gill apparatus evolved from being the primary respiratory organ to becoming remodelled into structures associated with a neck during the transition from water to land. Therefore through vertebrate evolution, although pharyngeal arch number can vary within a particular clade, there has been a general trend toward a reduction in their number in more derived species as the function of the pharyngeal apparatus has become more refined following its adaptation to new environments. Anuran species within the tetrapod clade of Amphibia show interesting intermediate features of the pharynx allowing the fully grown organism to adapt from a life based in the water to a land-dwelling life. When metamorphosing species hatch they have gills covered by an ‘opercular flap’, a caudal extension of the second arch, which during tadpole development will fuse with their ventral surface internalising the gills (Callery and Elinson, 2000, Callery et al., 2001). Even in direct-developing anuran species ‘opercular folds’ are present during embryogenesis, internalising the pharyngeal arches prior to hatching (Callery and Elinson, 2000). This is similar to the embryogenesis of amniotes, including birds and mammals, which undergo no such metamorphosis but retain a similar program during their early development of the pharyngeal arches. This process involves the posterior expansion of the second pharyngeal arch until it makes contact and fuses with the epithelium of the cardiac eminence to internalize all posterior arches (Richardson et al., 2012). This is reminiscent of anuran tadpole110 developing species metamorphosing stage, but without the arches ever functioning as gills. While it is apparent that the pharyngeal arches form into different structures dependent on the clade, it is also hard to ignore the homology of the structures the arches give rise to, regardless of the final form they have. The gills of waterbased species function not only for respiration but also for calcium homeostasis. In more derived, land-dwelling vertebrates that do not have gills, this function is accomplished by the parathyroid glands. These glands are unique to tetrapods, having evolved to regulate calcium internally after the move from water to land (Okabe and Graham, 2004). In tetrapods, Gcm-2 expression is detected in the developing parathyroid glands and endodermal pharyngeal pouches they develop from (Okabe and Graham, 2004, Gordon et al., 2001, Gunther et al., 2000). To identify the evolutionary origins of the parathyroid glands, Okabe and Graham (2004) tested for gcm-2 in zebrafish and found its expression in the developing gills. Expression of casr, which encodes calcium-sensing receptors, and pth, which encodes parathyroid hormone, were also detected, both of which are integral to parathyroid function. This clearly shows that, despite the difference in the anatomy and morphology seen between fish and tetrapod species, these structures are homologous and amniotes simply adapted to their new environment by internalising their ‘gills’ in order to retain this function. On observing the similarity between vertebrate embryos at the phylotypic stage, Ernst Haeckel stated that ‘ontogeny recapitulates phylogeny’ (Haeckel, 1910). Of course this is not true, but it does refer to a conserved stage of development at a certain period of embryogenesis, raising the question of whether the pharyngeal arches follow a conserved developmental program during these early stages despite the fact they will develop into different structures later on. In the previous chapter I used the chick to analyse the relationship between the epithelia at the pharyngeal pouch/cleft interface, revealing that the endoderm of the pharyngeal pouch has the competence to break through the overlying ectoderm to make contact with the external environment. I therefore wanted to analyse the relationship between the endoderm and ectoderm at the pharyngeal pouch/cleft interfaces in basal vertebrates, which are represented in this study by 111 the agnathan lamprey, chondrichthyan shark and osteichthyan zebrafish. I have then compared these results with those of amniotes, the chick (from the previous chapter) and mouse, to determine whether a conserved morphogenetic program is evident during early development of the pharyngeal region. 112 4.2 Results 4.2.1 Location of the shark ectoderm/endoderm interface during pharyngeal pouch formation In the previous chapter I described the relationship of the endoderm with the overlying ectoderm at the pharyngeal pouch/cleft interface in the chick. There are clear differences in their interaction between anterior and posterior pouches, although a general morphogenetic program of development exists across the posterior pouches resulting in their eventual break-through out into the external environment. To determine if this program is also seen in basal gnathostomes, whose posterior pouches will develop into gills, I have looked at the relationship of these epithelia at the pouch/cleft interfaces in Scyliorhinus canicula, the species of shark used throughout this study. In order to look at the interaction of the epithelia at the pouch/cleft interface, I have used laminin immunofluorescence to visualise the basement membrane that separates them. In order to demonstrate changes in the pouch/cleft interface as they mature, I have investigated the development of the pouch/cleft interface over three different stages (n=3 embryos at each stage). Figure 4.1. shows a shark embryo at stage 19 where only three (out of a total of 6) pouches have developed. In sharks, the first pharyngeal pouch will break through when contact is made with the overlying ectodermal cleft to form the spiracle (Figure 4.1. A; pp1), which allows the organism to continue breathing when its mouth is closed (Baker et al., 2008). The second pharyngeal pouch has clearly been evaginating outward and has made contact with the overlying ectoderm, and while a distinct basement membrane is visible on the lateral sides of the pouch, it is not as distinct over the middle portion where it appears to have degraded (Figure 4.1. B, F). The third pharyngeal pouch endoderm has made contact with the overlying ectoderm at a small focal location (Figure 4.1. C, G), while lateral of this point of contact separate basement membranes are evident for each of the epithelia. This suggests this pouch has evaginated to contact the overlying ectoderm but has only just 113 done so, and this point of contact will continue to enlarge as the basement membranes of the epithelia fuse together. Five pharyngeal pouches are present by stage 21 (Figure 4.2. C). The first pharyngeal pouch and cleft is still separated by a basement membrane, which is somewhat disorganised at this stage as evidenced by intense α-laminin labelling (Figure 4.2. D). The intense labelling is present bilaterally and is within the confines of the tissue itself, indicating this observation is real. However there is also some intense localisation of the laminin antibody within the fourth and fifth pharyngeal pouches (Figure 4.2. C), but this is unilateral and appears to be trapping of the antibody at the lateral-most portion of the lumen of the pouch. At this stage the first pouch has not broken through (Figure 4.2. D), even though the second pouch has (Figure 4.2. E, I). Results from the chick data presented in Chapter 3 indicated that anterior pouches are more ‘mature’ than those posteriorly, and so I would have expected the first pouch to break through earlier, again highlighting a difference in the way the first pharyngeal pouch develops as discussed in the previous chapter. The second pouch endoderm has broken through and is now interacting with the external environment, which will function in the adult as a gill slit along with all pouches posterior to here. At this stage the third pharyngeal pouch/cleft interface has thinned greatly and no basement membrane is evident separating the endoderm and ectoderm, indicating it has broken down since stage 19 (Figure 4.2. F). The next posterior fourth pouch has a basement membrane which has started to break down as evidenced by its spotty appearance, although its presence shows a distinct thinning of the ectoderm layer and the endodermal pouch pushing through it toward the external surface (Figure 4.2. G, J). The most posterior fifth pouch/cleft interface present at this stage reveals an intact basement membrane separating the endoderm and ectoderm, with both epithelial layers retaining the same thickness (Figure 4.2. H, K). By stage 22, the second, third and fourth pharyngeal pouches have all broken through, with a fifth pouch beginning to bud off the posterior pharynx (Figure 4.3. A). The first pouch has still not broken through, although the basement membrane is no longer disorganised and does appear to have begun its degradation (Figure 4.3. B (arrowheads) and F). The ectoderm layer is also now 114 markedly thinner than the endoderm. The most posterior pouch/cleft interface at this stage is the penultimate fifth pouch which at this point has a distinct basement membrane separating the epithelia, although it does appear to be starting to degrade at its anterior aspect (Figure 4.3. C). A general morphogenetic program for the interaction of the endoderm and ectoderm appears to apply to all the gill-forming posterior pouches in the shark. The epithelia make contact and their basement membranes fuse, whereupon its breakdown is initiated allowing direct contact between the epithelial cells. The ectodermal layer then thins significantly as the endodermal pouch continues to push through it out into the external environment when it perforates, allowing a connection between the pharyngeal lumen and the external environment. 115 Figure 4.1. Location of the pharyngeal pouch/cleft interface in stage 19 shark embryos Confocal sections of stage 19 shark embryos following laminin immunofluorescence (anterior to the left). The location of the basement membrane can be visualized separating the pharyngeal pouches from their overlying ectoderm. (A) A view of all pharyngeal pouches present at this stage, with white arrows pointing toward each pouch and labelled appropriately. (B) A zoomed in view of the second pharyngeal pouch, showing a continuous basement membrane at the lateral edges of the pouch with less prominent labelling along the middle of the interface indicating a discontinuation (arrowheads). Magnification of this region with DAPI staining to highlight individual cells reveals clear laminin at the edges of the interface but none across the middle, showing this part of the basement membrane has broken down. (C) A magnified view of the third and most posterior pharyngeal pouch at this stage reveals an intact basement membrane separating the endodermal pouch from the ectoderm of the cleft. This image is magnified further in (G) to show a continuous basement membrane separating the epithelia. (D) Schematic representation of the section through the arches as seen in (A), and a close up of the pouch/cleft interface is depicted in (H) to represent the morphology shown in images (B, C, F, G). 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 116 117 Figure 4.2. Location of the pharyngeal pouch/cleft interface in stage 21 shark embryos Confocal sections following laminin immunofluorescence reveals location of basement membrane at the pharyngeal pouch/cleft interface (anterior to the left). (A) Schematic representation of a coronal section through the arches to show the morphology seen in (C). A close up view of the pouch/cleft interface is depicted in (B) to represent the morphology seen in images (D-K). (C) An overview of all 5 pouches seen at this stage. (D) A zoomed in view of the first pouch reveals a discontinuous basement membrane separating the endoderm and ectoderm at their interface. (E) The second pharyngeal pouch has broken through following a breakdown in the basement membrane. (F) Barely any basement membrane is visible separating the epithelia, and the interface has become remarkably thin (arrowheads mark juncture between endoderm and ectoderm cells). (G) The interface at the 4th pharyngeal pouch is also beginning to thin, particularly in the ectodermal layer, and a visible but discontinuous basement membrane is present (arrowheads). (H) An intact and distinct basement membrane can be seen separating the two epithelia, with both epithelial layers each retaining a 2-3 cell deep thickness. (I-K) Magnified images of particular pouches with DAPI staining to show individual cells. (I) A magnified view of the second pouch in (E) showing this pouch has broken through and perforated and the two adjacent arches are separate from each other. (J) A magnified view of the fourth pouch shown in (G) to clearly show the discontinuous basement membrane. (K) The fifth and most posterior pouch magnified from (H), showing a continuous basement membrane. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 118 119 Figure 4.3. Location of the pharyngeal pouch/cleft interface in stage 22 shark embryos Confocal images of sectioned embryos following laminin immunofluorescence reveals the basement membrane separating the pharyngeal pouch/cleft (anterior to the left). (A) This image gives a general overview of all pouches present at this stage, with (B) and (C) showing zoomed in images of pp1 and pp5 where a basement membrane is still intact. As the dogfish embryo matures, the basement membranes separating the ectoderm and endoderm of middle pouches breaks down, followed by interaction of the epithelial cells, a thinning of the interface and perforation. (B) The first pharyngeal pouch retains a distinct basement membrane separating the pouch and cleft, although this has now begun to break down (arrowheads). (C) The fifth pouch basement membrane will break down (arrowhead) and perforate as more posterior pouches form with continued development. (D) Schematic representation of a coronal section through the arches as seen in (A). (E) A schematic representation of a pharyngeal cleft/pouch interface magnified and seen in images (A, B, C, F). (F) Magnification of (B) to show more clearly the basement membrane that has only just begun to break down in the first pouch. 1-6 – pharyngeal arches; ov – otic vesicle; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 120 121 4.2.2 Comparing pharyngeal endoderm/ectoderm pouch interface development between at amniotes the and anamniotes. The previous chapter described the morphology of each pharyngeal pouch in the chick embryo during development, while this current chapter has so far focused on pharyngeal pouch development in the shark. A comparison of pharyngeal development between these two species reveals significant similarities considering the different structures that will eventually form. However, one obvious difference between these two species is the development of the first pharyngeal pouch. The first pouch in the chick remains mostly separated from the overlying ectoderm with mesenchyme separating these two tissues, and will eventually develop into the internal auditory canal so therefore never perforates. However, this is not the case in the shark. The shark first pouch does remain mostly in direct contact with the overlying ectoderm with a distinct basement membrane separating the epithelia, but will eventually break through at a later stage to form the spiracle. This emphasises a difference of first pharyngeal pouch development not only within a species but also across different species. The rest of the pharyngeal pouches appear to follow a similar developmental program. As mentioned already, the more anterior the pouch, the better developed it is. When comparing an anterior, middle and posterior pouch of the chick and shark at developmental stages when the chick has all 4 pouches present and the shark has at least the same amount (Figure 4.4. A and E), a resemblance in the morphogenetic program is seen across both species. The most anterior (aside from the first) pouch has broken through entirely in both species (Figure 4.4. B, F). The shark pouch/cleft interface will remain broken through into adulthood and forms a gill slit, whereas the chick second pharyngeal arch will expand caudally to cover and close all broken through or potentially broken through posterior pouches. Examination of ‘middle’ pouch interfaces in both species reveals a basement membrane between the epithelia in the process of degrading (Figure 4.4. C, I, G, J). At the shark interface, the endoderm is pushing into the ectoderm as the ectoderm layer thins significantly in comparison. In the chick, the endoderm is pushing against the ectoderm causing it to bulge outward, and a 122 significant thinning of the ectoderm layer is also seen. At the most posterior pouch/cleft interface of both species, a distinct basement membrane is evident separating the epithelia which are at this point of equal thickness (Figure 4.4. D and H). 123 Figure 4.4. Comparison of the pharyngeal pouch/cleft interface during shark and chick pharyngeal development This figure gives an overview of how the pouches develop along the anterior to posterior axis in shark and chick, with the anterior pouches being more mature than those posteriorly (anterior to the left). (A-D) Confocal sections following laminin immunofluorescence of stage 21 shark embryos. (A) Overview of all pouches zoomed in on in (B-D). (B) An ‘anterior’, and therefore more mature, pouch present at this time point, which has broken through. (C) A ‘middle’ pouch whose basement membrane is in the process of degrading as the interface begins to thin, particularly at the ectodermal layer. (D) A ‘posterior’, or less mature, pouch which is distinct from the overlying ectoderm with an intact basement membrane separating them. (E-H) Confocal sections following β-catenin and laminin immunofluorescence of stage 19 chick. (E) This image shows the general morphology of all pouches present in the chick, with the white arrows labelling each pouch. (F) The second pharyngeal pouch, or the ‘anterior’ pouch, which at this stage has broken through, reminiscent of how the ‘anterior’ pouch in the shark behaved. (G) The ‘middle’ pouch shows a breakdown in the basement membrane separating the endoderm and ectoderm, while the ‘posterior’ pouch in (H) shows an intact basement membrane still present. (I, J) Magnification of boxed regions in (C and G) respectively clearly representing the ‘middle’ pouches with degrading basement membranes. How each pouch behaves in the chick as it develops shows a remarkable similarity to how those in the shark develop, hinting at a conserved developmental program in this region. (K, L, M) Schematic representation of the pharyngeal morphology shown in this figure. (K) Lateral view of a chick embryo. The dotted line represents the coronal section through the arches seen in (L). (L) Representation of the morphology seen in images (A and E). (M) Magnified pouch/cleft interface as seen in images (B-D and F-J). 124 125 4.2.3 Pharyngeal pouch development in the lamprey. To further explore the homology of this morphogenetic program of pouch formation, I decided to look at the relationship of the epithelia at the pouch/cleft interface in a basal vertebrate species, and have used the agnathan lamprey for this study (see Figure 1.4. for its location in vertebrate phylogeny). In this species, the first pharyngeal pouch will contribute to musculature of the posthypophyseal process, velum, and velar chamber, and the rest will develop into gill pores. By stage 26 (n=3) all eight pharyngeal pouches are present (Figure 4.5. A). As the anterior pouches are better developed than those posteriorly, it is evident that a change in general pouch morphology occurs over time. The most posterior, or least mature, pouches look similar to those in other vertebrate species with a curved-V shape morphology and a basement membrane present separating the endoderm from the ectoderm (Figure 4.5. A, white asterisks). Looking at the more anterior pouches however reveals a drastic change in their morphology, showing a much more pointed triangular shape with the tip of the triangle, where the interface of the pouch/cleft should be, curving posterolaterally. The pouch in this region also appears to be a single layer bulging out through the external surface of the lamprey pharyngeal region (Figure 4.5. A, blue asterisks), although whether this is ectoderm, endoderm, or a mixture of the two cannot be derived from this data. By stage 27 (n=3) the posterior pouches remain distinct from anterior ones with a clear basement membrane still separating the epithelia (Figure 4.5. B, white asterisks). Anteriorly, a basement membrane cannot be seen, and although the pouches seem to continue to push against the overlying ectoderm they are not as bulged as they were at the previous stage, appearing to have flattened out somewhat, and none of the pouch/cleft interfaces have broken through. The arches themselves have thinned a lot since the previous stage and are triangular in shape. The posterior pouches in stage 28 (n=4) embryos have started to lose such strong expression of laminin (Figure 4.5. C, white asterisk), and these pouches have adopted a similar morphology to that seen in the anterior pouches at stage 26, confirming that they do develop later than the anterior pouches. 126 Figure 4.5. Location of pharyngeal pouch/cleft interface in lamprey embryos Confocal images of lamprey sections following laminin immunofluorescence (anterior to the left). (A) A stage 26 lamprey embryo reveals laminin labelling at the posterior pouch/cleft interfaces (marked by white asterisks). Anteriorly no laminin labelling is evident at the interfaces and the same developmental program seen in chick and dogfish is not apparent either, although there is an outpocketing at the location of the interfaces as seen in the chick third pouch. (B) A stage 27 embryo reveals laminin labelling present still in the posterior pouches (marked by white asterisks) but none in the anterior pouch/cleft interfaces. (C) At stage 28 there is an unusual pouch morphology as the embryo progresses with development, and laminin labelling in the posterior pouches is not as marked any longer (white asterisk). (D) By stage 30, the pouches have completely altered their morphology from an earlier state and look entirely different from the other species examined. Gill openings are present (white arrowheads) but no apparent link to a basement membrane breakdown has been recorded. (E) Schematic representation of the head of a lamprey embryo from a lateral viewpoint. The dotted line represents the plane of the coronal section represented in (F), depicting the pharyngeal morphology seen in (A-D). pp – pharyngeal pouch. 127 128 By stage 30 (n=3), no laminin expression is visible at all between the ectoderm and endoderm and it is difficult from this data to determine the relative positions of each epithelium. However, the distinct change in morphology of the pharyngeal arches is evident as they are now a greatly reduced scalene triangular shape with long thin bars of internal gills protruding off them into the pharyngeal cavity (Figure 4.5. D). The pouch region is now distinctly square in shape, and the posterior edge at the interface appears to have ‘opened up’ rather than broken through due to a ‘flap’ remaining that appears as though it has just been lifted off the anterior surface of the posterior arch (Figure 4.5. D, white arrows). This is reminiscent of the morphology seen when the chick second pharyngeal pouch breaks through initially at the anterior edge of the interface nearest the posterior border of the second pharyngeal arch and suggests this perforation may have occurred due to tensile stress caused by the drastic changes in the morphology over a short period of time. 129 4.2.4 Endoderm cell lineage tracing reveals the location of the pharyngeal pouch/cleft interface in transgenic mouse and zebrafish lines. 4.2.4.1 Sox 17 transgenic zebrafish My investigation of the relationship between the ectoderm and endoderm at the pouch/cleft interface in the shark suggest that, following fusion of the basement membranes and its subsequent degradation, the ectoderm layer thins greatly as the endoderm pushes against it, presumably to make contact with the external environment prior to its perforation for the development of the gills. To reliably determine whether the endoderm does push out into the external environment, I have used transgenic zebrafish to cell lineage trace the endoderm of the pharyngeal pouches and determine its exact location during pouch development in order to define its relationship to the ectoderm and adjacent pharyngeal arches. In order to achieve this I used a Tg(sox17:GFP) zebrafish line (Chung and Stainier, 2008), in which the Sox17 regulatory region directs expression of a GFP transgene allowing visualisation of the endoderm. At 48hpf (n=4) the second arch has begun growing posteriorly, covering the third and fourth arches (Figure 4.6. A; pink asterisks). Strong Sox17 expression is evident at the external surface of each pharyngeal arch posterior to the operculum (Figure 4.6. A). A magnified view reveals this is likely because the Sox17expressing pouch endoderm begins development internally as normal, but as development proceeds the pouches break through the overlying ectoderm (Figure 4.6. C), as shown previously in the chick and suggested in the shark data. To be confident this is part of normal development, a later stage of 72hpf (n=6) was also examined revealing a similar result of strong Sox17 expression along the external surface of the pharyngeal arches (Figure 4.6. B; white asterisks). In addition, the inner surface of the operculum strongly expresses Sox17 indicating this part of the flap is of endodermal origin, which has not been reported before (Figure 4.6. A and B; pink asterisks). This is true in all embryos examined and is 130 evident both at 48hpf and 72hpf, indicating the second pharyngeal pouch endoderm pushes through the overlying ectoderm to continue expanding posteriorly with the second arch as the inner lining of the operculum and will directly cover the pharyngeal arches located beneath. A magnified view of the 72hpf pharyngeal pouches reveals the anterior half of the pouch contributes to the posterior portion of the anterior adjacent pharyngeal arch, revealing the endoderm does become part of the external surface of the arches (Figure 4.6. D). 4.2.4.2 Sox17 transgenic mice I have shown that the gills develop in zebrafish following the outward migration of pharyngeal pouch endoderm to contribute to the external surface of the pharyngeal arches. Studies in the chick suggest that this morphogenetic program is similar, so I decided to examine endodermal movement at the pharyngeal pouch/cleft interface in a mammalian species, the mouse, to confirm whether or not a conserved morphogenetic program for pouch maturation exists in vertebrates. To analyse this interaction I wanted to lineage trace endodermal cells of the pharyngeal pouches in order to define their relationship with the overlying ectoderm and adjacent pharyngeal arches in the mouse, so therefore utilised a transgenic Sox17 mouse line for this study. Sox17-2A-iCre mice were crossed with R26R (R26 reporter) mice, which allows visualisation of Sox17-expressing endodermal cells and their descendants during early stages of development (Engert et al., 2009). A lateral view of wholemount embryos shows Sox17 expression in the pharyngeal pouches (Figure 4.6. E and G, white asterisks). A coronal section through the arches reveals the morphology of each pouch and the nature of its interaction with the overlying ectoderm. At E9.5 (n=3), the ectoderm and endoderm are separate from one another at the first pouch/cleft interface, as was described in the chick in the previous chapter (Figure 4.6. F; pp1). The second pouch at this stage has evaginated toward the external surface and leaves only a thin layer of endoderm connecting the second and third pharyngeal arches (Figure 4.6. F; pp2). This is different to what is seen at the chick second 131 pouch/cleft interface, where some ectoderm cells contribute to the thin layer connecting the arches before breaking through at its anterior aspect. By E10.5 (n=3) the pouches are seen to behave in a similar way as has been described for the continued development of chick, shark, and zebrafish pharyngeal pouches. The first pharyngeal pouch remains separate from the overlying ectoderm (Figure 4.6. H; pp1), while the second pharyngeal pouch has broken through, allowing the endoderm contact with the external environment (Figure 4.6. H, pp2). At this stage the second pharyngeal arch has grown significantly since E9.5, and its posterior portion is now beginning to expand over the anterior part of the third pharyngeal arch. This is perhaps the reason for the break-through of the second pouch, similar to that seen during second pouch perforation in the chick, which has been suggested to be due to increased stress exerted by the growing second arch. At E10.5 the third pharyngeal pouch has become more triangular in shape, and has pushed through the ectoderm to make contact with the external environment (Figure 4.6. H; pp3). This is homologous with the behaviour of the third pouch in the chick, and poses the question of whether it too would break through at a later stage if the expansion of the second arch was inhibited. This data shows that a similar morphogenetic program does appear to exist across the vertebrates, with amniotes attempting to form ‘gills’ early in development only to have this region remodelled later on. 132 Figure 4.6. The pharyngeal pouch/cleft interface of transgenic Sox17 mice and zebrafish (A-D) sox17:gfp zebrafish (anterior to the top). (E-H) Sox17iCre;R26R mice (anterior to the left). (A) Confocal image of a coronal section through a 48hpf zebrafish embryo. The endoderm of all pharyngeal pouches has pushed out toward the external surface of the embryo. The posterior pharynx is blocked from view because of the still enlarged yolk (orange asterisk). The operculum (pink asterisk) covers the first two posterior arches and its inner lining expresses Sox17. The boxed region is magnified in (C) showing the pouch has grown outward past the adjacent pharyngeal arches. (B) Confocal section of a 72hpf zebrafish reveals the pharyngeal arches (white asterisks) with contribution to their external lining by endoderm. The boxed region is magnified in (D) showing the anterior half of the pouch extends over the posterior portion of the anterior adjacent arch. (E) A lateral view of an E9.5 wholemount embryo shows Sox17 expression in all pharyngeal pouches (white asterisks). (F) The first pouch is separated from the overlying ectoderm by mesenchyme. The second pouch interface has thinned so only a narrow band of endoderm connects the second and third arches. The third pouch is in contact with the overlying ectoderm and appears to bulge through it, with the ectoderm much thinner than the endoderm. (G) A lateral view of a wholemount embryo at E10.5. (H) A coronal section through the arches at E10.5 reveals the morphology of the pouches at this later stage. The first pouch is still separate from the cleft. The second pouch has now broken through with endoderm from the anterior border of the pouch beginning to extend posteriorly over the anterior surface of the third arch as the second arch expands caudally. The third pouch endoderm has broken through and is now in contact with the external environment. (I) Schematic representation of a coronal section through zebrafish arches showing the second arch expanded caudally as the operculum. (J) Schematic representation of the side of the embryo head seen in (E and G). The dotted line represents the plane of the coronal section seen in (K). (K) Schematic representation of the arch morphology seen in (F and H). ov – otic vesicle; PA – pharyngeal arch; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 133 134 4.3 Discussion Despite the different adult form of species within each vertebrate group, they all bear pharyngeal arches during early stages of development. Following their development, fish species will retain these segmented structures once they have broken through to form gills that function in respiration, while amniote species will extensively remodel this region so no evidence of an early segmented series is visible. Despite these differences in final morphology, my results have shown that all vertebrates follow a similar early morphogenetic program for gill formation, whether they develop them as adults or not, contributing to existing evidence that these structures are indeed homologous across all vertebrates. 4.3.1 Epithelial interactions at the pharyngeal pouch/cleft interface are conserved in fish and amniotes Comparing the movement of epithelia between the shark, which develops gills from its posterior pouches, and the chick, whose posterior pouches become internalised and contribute to neck structures, reveals there is a conserved program during early pharyngeal development. As shark pharyngeal segmentation occurs an iterated series is evident with all posterior pouches retaining a similar morphology. This is different to what was described in the chick during the previous chapter, where each pouch had a different shape. This probably reflects the structures those pouches will develop into and as all shark posterior pouches will form gills, it is not surprising they all start with a similar morphology. Despite the differences in the morphology of the pharyngeal pouches between these two species, there are surprising similarities in the way their epithelia interact with one another. In both species, the pharyngeal pouches arise along the anterior to posterior axis, meaning the anterior pouches are developmentally more mature than those posterior. In both the chick and shark, once the pouch endoderm made contact with the cleft ectoderm their basement membranes fused before its degradation 135 was initiated. This degradation in turn leads to a thinning of the ectoderm as the pouch endoderm continues to grow laterally toward the external surface, eventually breaking through the ectoderm. In the chick the second pouch interface breaks through, reminiscent of what is seen during gill formation as evidenced during the development of all posterior pouches in the shark (Figure 4.4). This implies that a common morphogenetic program is employed during early stages of pharyngeal development across the vertebrates, where the chick is apparently attempting to form ‘gills’ by following the same developmental process as more basal species, before a different program is initiated preventing this from continuing any further. The first pouch however is different, and this difference is apparent when compared with other pouches within the shark and also when compared with the first pouch in the chick. The shark first pouch is in contact with the overlying ectoderm with a basement membrane separating the epithelia at all stages examined. This is different from what occurs in all the other pouches, where as soon as the epithelia make contact a developmental program is initiated causing the breakdown of the basement membrane. However, the first pouch interface morphology at stage 22 reveals a similar program appears to be responsible but with some kind of delay put in place, as the pouch has a thicker epithelium than the overlying ectoderm and a breakdown in the basement membrane is just becoming evident, presumably allowing the pouch to break through shortly after this stage (Figure 4.3. B, F), which it must do in order to form the spiracle. The fact that this is the most anterior pouch and yet is slower in its development indicates a difference in the way this pouch develops, possibly from a different regulatory program than that which controls posterior pouch development. This difference in first pouch morphology and development appears to be conserved across vertebrates, with the chick revealing a similar pattern as discussed in the previous chapter. With regards to the pharyngeal arches, the first and second arches are quite different from the rest, although this is not surprising due to the structures they will form. The first arch will develop into the jaw, although when compared with the morphology of the first arch in chick (Figure 4.4. A and E) a clear difference 136 is seen, with the shark first arch being much larger and more elongated than the chick first arch. Comparatively, the second arch in the chick is much larger than its first arch, whereas the first arch in the shark is much larger than its second arch. The reason is that the chick second arch enlarges to form an ‘opercular flap’ which will enclose all the posterior arches and pouches (Richardson et al., 2012), whereas the shark second arch does not do this because it does not form an operculum. The zebrafish second arch however does form an operculum similar to the opercular flap in amniotes (Figure 4.6. A and B; pink asterisks), but it does not fuse to the body wall. Therefore the largest arch in osteichythyans is the second arch, whereas this is not the case in chondrichthyans and is representative of the evolution of such structures as seen from their phylogeny (Figure 1.4). Lamprey pouch development is quite different from that seen in any other species examined, and highlights some key differences that are involved in pharyngeal development between agnathan and gnathostome species. The shape of the arches and pouches are quite different from other species, and may be because they form gill pores rather than gill slits, and therefore require a more extensive internal gill filament developmental program. However, the posterior pouches in the lamprey pharynx do show laminin expression similar to what is seen in posterior pouch development across the other species, with an initial fusion of the endoderm and ectoderm evident. Following this, a breakdown of the basement membrane followed by thinning of the pouch/cleft interface is not however seen, with the gill pores not breaking through until quite a while after initial epithelial contact. Even then the gap is restricted to one end of the interface by opening as a flap rather than the entire pouch/cleft interface region breaking through. The fact that there is some similarity in the initial epithelial interaction within the posterior pouches hints at some homology of this developmental process, even though it does not appear to be maintained throughout pharyngeal segmentation. The number of pharyngeal arches differs also between lamprey, sharks and chick. Lamprey have nine arches, the shark has seven, and the chick has only five. The posterior pouches in the lamprey and shark will break through to form gills in the adult, whereas in the chick the pouches may break through during their embryonic development, but will be enclosed eventually so no opening is evident 137 in their fully developed form. It can be assumed that as all the posterior arches and pouches are the same in the lamprey and shark, giving rise to a metameric series of gills, it is posteriorly that the number of arches has been reduced to reach the five seen in amniotes. This will be discussed in more detail in the next chapter. 4.3.2 Cell lineage tracing reveals conservation of pharyngeal pouch out-pocketing 4.3.2.1 Sox17 as an endoderm marker Sox17 belongs to the Sry-related HMG box (Sox) transcription factor family which play a role during various developmental processes (Pevny and LovellBadge, 1997, Wegner, 1999). Sry is a testis-determining factor in mammals (Gubbay et al., 1990, Sinclair et al., 1990), and carries a characteristic high mobility group (HMG) domain which binds DNA in a sequence-specific way. 20 different Sox genes have been identified in humans and mice (Schepers et al., 2002), and these have been further divided into groups A-J depending on the homology of the sequence identity of their HMG domains (Bowles et al., 2000). Sox17 falls within group F, alongside Sox7 and Sox18, and was initially identified as a transcription factor involved in controlling mouse spermatogenesis (Kanai et al., 1996). Sox17 also functions in the maintenance of fetal hematopoietic stem cells and plays a redundant role with Sox18 in vasculogenesis (Kim et al., 2007, Matsui et al., 2006, Sakamoto et al., 2007), as well as being a key regulator for endoderm formation in zebrafish (Alexander and Stainier, 1999), Xenopus (Clements and Woodland, 2000, Hudson et al., 1997) and mice (Kanai-Azuma et al., 2002), indicating a conserved function across vertebrates. Mouse Sox17 null embryos display severe endoderm deficiencies with hugely disrupted gut formation, showing Sox17 is essential to the maintenance and differentiation of the endoderm (Kanai-Azuma et al., 2002). As Sox17 is crucial to endoderm 138 formation in the gut and is expressed during early development, it is an ideal marker to trace endodermal cells. 4.3.2.2 Pharyngeal pouch endoderm shows conserved development despite differences in epithelial organisation between fish and amniotes Transgenic animals are a useful tool in developmental biology by allowing a specific cell lineage to be traced visually, therefore making it possible to determine the embryonic origins of a particular structure. A Sox17-2A-iCre mouse line was developed for the purposes of tracing Sox17 expressing cells (Engert et al., 2009), and has been utilised in this study for lineage tracing pharyngeal pouch endodermal cells which originate from foregut endoderm. Similarly, a Tg(sox17:GFP) zebrafish line was used to identify endodermal cells in the zebrafish pharyngeal region. Examination of these transgenic embryos has revealed conservation in the way the pharyngeal pouches develop across vertebrate species by revealing that they will break through the ectoderm to the external surface of the embryo irrespective of whether the species will form gills or not. This follows the results seen in the previous chapter using CCFSE to cell lineage trace the ectoderm in chick. This revealed that the second pouch interface breaks through after the endoderm pushes against the ectoderm, and that a distinct enlargement of the third pouch endoderm is seen followed by some cell death of the overlying ectoderm which may be responsible for allowing the pouch to push through the ectoderm. Even though apoptosis could be responsible for the disappearance of ectoderm at the pouch interface in chick, this does not mean this mechanism is conserved across all species, as studies in primary mouth development across different vertebrate species reveals apoptosis is involved in some but not all species during rupture of the oral membrane (Soukup et al., 2013). Sox17 cell lineage tracing of the endoderm has revealed that it too lines the external surface of the pharyngeal arches in the zebrafish. As the pharyngeal pouches posterior to the second arch will contribute to the gills they break through and make contact with the external environment. However, the extent to 139 which they appear to spread over the external surface of the arches is surprising and has not been shown before. Homology between the gill-bearing epithelium of the zebrafish and parathyroid forming epithelium of the chick has been shown by the expression pattern of Gcm-2 (Okabe and Graham, 2004, Hogan et al., 2004). However, studies have claimed that the gills are derived from the ectoderm of pharyngeal arches 3-6 (Hogan et al., 2004). Hogan et al. (2004) suggested that gcm-2 expression is seen in the ectoderm of these arches, and propose that the gills and parathyroid gland have the same evolutionary origins due to their development being dependent on the same molecular regulator, gcm-2, despite being derived from different germ layers and despite serving different physiological functions. However, I have cell lineage traced the endoderm in Tg(sox17:GFP) zebrafish to the location where the ectoderm would be expected. This presents an explanation for the conflicting data of which epithelium expresses Gcm-2 between Hogan et al. (2004) and Okabe and Graham (2004), who state it is expressed in the endoderm of gill-forming pharyngeal pouches. My data shows that it is in fact expressed in the endoderm, but that this epithelium is partially localised to the external surface of the pharyngeal arches in the zebrafish. Combined with the data from the laminin immunofluorescence in both shark and chick, it is evident a homology exists in pharyngeal segmentation and during the interaction of the epithelia at the pouch/cleft interface. Although there are clear differences present still between the species, these are mostly limited to a distinct difference between the development of anterior and posterior pouches, which is also true within each individual species. How each pharyngeal arch is patterned is a topic that has been investigated for a long time (Graham and Smith, 2001), although it has become evident the pharyngeal pouches play a key role in this patterning (Veitch et al., 1999). To understand this, the patterning of the pharyngeal pouches themselves needs to be elucidated, and is something that will be discussed in the following chapter. However, it is becoming apparent that a general program of pharyngeal pouch/cleft epithelial interaction may be responsible for segmentation of the arches, and perhaps this program is modified by relevant signals present in the surrounding areas of each pouch. 140 4.3.3 Outpocketing of the pharyngeal pouches and operculum development By analysing the pharyngeal pouch/cleft interfaces in both mouse and zebrafish, it is clear a pattern of the endodermal pouch pushing through to the external surface of the embryo is a key and conserved feature during pharyngeal development. This must happen in fish as their internal gills are derived from the endoderm of the pharyngeal pouches (Okabe and Graham, 2004), and without contact with the external environment they would not be able to function properly. However, as amniotes do not form gills and all pouch-associated structures are internalised when fully formed, the outpocketing of their endodermal pouches may occur as an evolutionary vestige of how earlier species have developed. Alternatively, if the medial border of the operculum is of endodermal origin, perhaps the outpocketing of the pharyngeal pouches is a necessary process to ensure proper fusion of the opercular flap over the top. Thyroid signalling is required for fusion of the opercular flap with the body wall in chick, as when this is blocked a cyst persists in the region between where the opercular flap covers the posterior arches (Richardson et al., 2012). Thyroid signalling has also long been shown to be important for metamorphosis in frog species (Gudernatsch, 1914). The developing thyroid gland releases thyroid hormone, which induces genetic cascades that initiate metamorphic events. However, the kind of morphogenetic changes that are caused by the same hormone differ greatly, evidenced by regression of the gills and tail as opposed to the growth and differentiation of limbs. Two subtypes of thyroid hormone receptors (TRs) coded by different genes have been isolated in Xenopus laevis: TRα and TRβ (Yaoita et al., 1990). Thyroid hormone receptors have also been isolated in the chick (Sap et al., 1986) and human (Weinberger et al., 1986), and so have been shown to be highly conserved in vertebrates. TRβ expression levels are higher in the gills and tail of Xenopus, structures associated with reabsorption and cell death during metamorphosis, and addition of the TR agonist GC-1 shows preferential binding to this receptor subtype, inducing gill and tail metamorphic events (Furlow et al., 2004). 141 However it is unclear which tissue within the gills, or developing pharyngeal arches contains this receptor subtype and is therefore responding to thyroid hormone signalling. (Tata, 1968) investigated the competence of larval tissue to respond to thyroid hormone but did so at a whole tissue level without investigating individual tissue layers. If the endoderm is the competent tissue in pharyngeal formation, it could help explain how the opercular flap fuses, why the medial border of the opercular flap, or operculum in fish, is composed of endoderm, and why the need for endodermal outpocketing is required and therefore retained in amniote species that do not form gills. 142 4.4 Summary By comparing pharyngeal endoderm segmentation across several vertebrate species, I have shown a conserved morphogenetic program exists for the formation of pharyngeal pouches and their subsequent interaction with the overlying ectoderm of the pharyngeal cleft. Sharks and zebrafish retain their pharyngeal arches into adulthood in the form of gills, while amniotes, represented in this study by the chick and mouse, internalise their arches and extensively remodel this region to form a smooth external appearance. Even though amniotes do not form gills they do possess their early form, yet their developmental program is altered when the caudally expanded second pharyngeal arch fuses with their ventral surface to enclose and internalise all the posterior arches. My data has shown that despite these differences in adult topography, initially a similar developmental program is followed and pharyngeal segmentation in amniotes reflects gill formation in fish, suggesting this process is conserved across the vertebrates. This is further evidenced by the homology of the opercular flap in mouse and chick to the operculum in the metamorphosing frog and the zebrafish. By revealing the epithelial composition of this structure in different vertebrate species, I have also uncovered the embryonic origins of particular structures associated with the operculum and pharyngeal region, as well as some potential mechanisms for opercular fusion in amniotes. 143 Chapter 5. Reduction in the number of pharyngeal segments 5.1 Introduction Over the course of vertebrate evolution, a general trend toward a reduction in the number of pharyngeal arches has occurred. For example, throughout this study I have compared a basal vertebrate agnathan species, the lamprey, which has 9 pharyngeal arches, with basal gnathostomes that have 7 pharyngeal arches, and with amniotes, which have only 5 arches. There can be variation in the number of pharyngeal arches within specific groups, for example hagfish can have between 5-14 arches and sharks can have 6-9. The variability in arch number within these groups is likely due to adaptations that have occurred according to the different environmental requirements each species has adapted to. The reason for a general trend toward a reduction in arch number throughout vertebrate evolution however is similar; with the transition from water to land, land-dwelling animals had no use for a large number of arches as they no longer required gills to breathe. The mechanism by which this reduction occurred is unknown yet would help to uncover how the pharyngeal arches are regionalised and thereby where within the pharyngeal apparatus the arches could have been lost from. 5.1.1 Hox genes in vertebrate body patterning Hox genes pattern the body along the A-P axis, and were first discovered in Drosophila melanogaster (Lewis, 1978). They share a common highly conserved 180bp sequence DNA-binding domain called the homeobox. Most vertebrates have four Hox clusters which evolved from a single ancestral cluster following two whole genome duplication events (Garcia-Fernandez and Holland, 1994), although an additional third rounds of whole genome duplication occurred in actinopterygian fish resulting in seven or eight Hox clusters (Stellwag, 1999). 144 There are 13 paralogous groups of Hox genes, and all share a similar organisation within these clusters, where each gene is oriented in the same 5’ to 3’ direction of transcription (Krumlauf, 1994). Interestingly, paralogous genes have higher sequence similarity, are expressed at more similar times and have more functional overlap than other genes located within the same cluster (Krumlauf, 1994). The protein product of Hox genes are transcription factors, which activate or repress specific genetic cascades leading to the development of distinct structures within the territory that the gene is expressed in. They employ a rule of ‘colinearity’, where their order on the chromosome is often the same as the order in which each Hox gene is expressed along the A-P body plan, and furthermore the anterior genes are expressed earlier in time than those more posteriorly (Gaunt et al., 1988, Graham et al., 1989, Duboule and Dolle, 1989). Vertebrate Hox genes have also been shown to be induced by RA signalling in a colinear fashion, with anterior genes being more sensitive to RA signalling than posterior genes (Dekker et al., 1992, Papalopulu et al., 1991). The mechanisms by which Hox genes pattern the A-P axis are not entirely clear, although it has been proposed they do so either by a unique combination of Hox gene expression at specific regions along the A-P axis (Kessel and Gruss, 1990, Hunt and Krumlauf, 1992), or by a mechanism of posterior prevalence, where posterior Hox genes within a domain are functionally more dominant than those anterior to it (Duboule and Morata, 1994). Previous studies have indicated endoderm is the key instructive tissue patterning the pharyngeal apparatus (Veitch et al., 1999, Piotrowski and Nusslein-Volhard, 2000). The expression of a specific set of Hox genes within the pharyngeal region of vertebrate embryos has been well described previously, although these studies have been restricted to Hox expression within NCCs migrating from the hindbrain to the pharyngeal arches (see Chapter 1 for a more detailed discussion). As Hox genes pattern the embryo along the A-P axis, these are good candidate genes to explore to determine whether they align with and impart an identity to the pharyngeal pouches, thereby regionalising the pharynx and patterning the surrounding pharyngeal arches. As Hox genes are axial markers, they also allow a 145 comparison of the organisation of the pharynx between species to identify whether this regionalisation is conserved across vertebrates. Previously, detailed information on Hox expression within pharyngeal arches was limited to data derived from osteichthyan lineages, but 34 Hox genes were recently identified within the chondrichthyan shark species Scyliorhinus canicula (Oulion et al., 2010) allowing a deeper investigation of conservation within gnathostomes. This data has revealed that the first pharyngeal arch has no Hox expression in mouse, chick and shark (Hunt et al., 1991a, Couly et al., 1998, Oulion et al., 2011), that the second arch expresses Hoxa2, Hoxb2 and Hoxb1 in mouse, chick and shark, and the third arch expresses Hoxa3 and Hoxb3 in shark, plus Hoxd3 in mouse and chick (Hunt et al., 1991a, Hunt et al., 1991b, Hunt et al., 1995, Oulion et al., 2011). The fourth arch is more complicated as early chick and mouse work suggested all Hox4 paralogues were expressed here (Hunt et al., 1991a, Hunt et al., 1995), although further exploration in mouse and shark revealed only Hoxd4 is actually expressed here while Hoxa4 and Hoxb4 expression in seen caudally (Minoux et al., 2009, Oulion et al., 2011). While this data reveals a conservation of the pattern of Hox gene expression in the pharyngeal arches, there has been no analysis of Hox expression specifically in the endoderm of the pharyngeal region, which is likely to be the key tissue responsible for patterning the pharyngeal apparatus. In this chapter I have analysed Hox expression within the pharyngeal endoderm, with particular attention paid to expression that would impart an identity to each pharyngeal pouch, as these structures separate each pharyngeal arch and are therefore most likely responsible for directing their patterning. 5.1.2 Vertebrate evolution has resulted in a loss of pharyngeal arches In agnatha, the first arch contributes to the upper lip and velum and in gnathostomes it develops into the jaw, while posterior arches persist into adulthood as the gills, but become remodelled in tetrapods to contribute to the throat. During the transition in function of the structures that are derived from the 146 pharyngeal arches, both with the emergence of gnathostomes and with the emergence of tetrapods, a general trend toward a reduction in their number has occurred. This modification of the pharynx is distinct between anterior and posterior portions, primarily because all gnathostomes have maxillary and jaw structures that surround their mouths and function in feeding and respiration, whereas the posterior pharynx in basal gnathostomes contributes to respiration but does not in amniote species. Nevertheless, amniote lungs develop from the posterior pharyngeal endoderm so there has not been as huge a shift in the function of this region, rather a change of form following their internalisation. This is also evident by the homologous origins of structures involved in calcium homeostasis, which is regulated by pharyngeal pouch-derived gills in fish and pharyngeal pouch-derived parathyroid glands in amniotes (Okabe and Graham, 2004). Almost nothing is known about how this loss in the number of pharyngeal arches occurred, and a mechanism for variation in the number of segments within a region has been sought after for centuries. Comparisons for the alteration in the number of pharyngeal segments can be drawn from other segmented series found within the body plan. Goodrich (1906) described how changes in the number of segments within the vertebral region can explain a shift in the location of limbs. He used the term ‘transposition’ to portray each segment as an adaptable homologous region that can slide up or down the A-P axis through evolution. It was later demonstrated that Hox genes are likely to be responsible for the transposition of vertebral segments via changes in their expression domains along the A-P axis, wherein the number of segments within a specific vertebral region is variable so long as the correct Hox genes are expressed conveying positional information at the anterior and posterior boundaries of these regions (Burke et al., 1995). It is clear from morphological and molecular data that a distinct difference exists between anterior and posterior pharyngeal arches and pouches, intimating regionalisation of this area. If Hox expression is found within the pouches to mark anatomical boundaries, I can try and determine from which territory the pharyngeal arches were lost from. 147 Despite the iterated arrangement of the pharyngeal arches, each obtains a distinct identity for the formation of distinct structures. Revealing how repetitive elements are specified in basal gnathostomes that develop gills will help further refine the location of where the pharyngeal arches have been lost. A combination of information regarding pharyngeal patterning will allow confidence in describing how this is achieved, and so I have undertaken a comprehensive analysis using several methods to determine where this reduction has occurred. This includes a morphological analysis of the pharyngeal pouches, an anatomical analysis investigating the pattern of innervation of the cranial nerves that innervate each arch, and a molecular analysis, analysing Hox expression within the pouches. I have already looked at the morphology of each pharyngeal pouch, both within an amniote species and across more basal gill-bearing species, and found differences between anterior and posterior pouches within and across all species, but also that all posterior gill-bearing pouches share the same morphology whereas each posterior pouch in the amniote differs, reflecting the final form they will take (see Chapter 3 and Chapter 4). In this chapter I have looked at anatomical data by analysing the nerves that innervate each pharyngeal arch within species from different clades. This information has been described previously but not with reference to the number of arches present, and so a comparison of these species to see if a particular nerve has fewer ganglia/processes in those with fewer arches should help identify the location from where the arches have been lost. To lend further support to this data I have also conducted a molecular investigation by analysing Hox expression within the pharyngeal pouches, which subsequently regionalises the pharyngeal apparatus. A combination of all this data has therefore allowed me to determine exactly where the pharyngeal arches have likely been lost throughout evolution. 148 5.2 Results 5.2.1 Cranial nerve innervation identifies where pharyngeal arch reduction has occurred To begin to identify which pharyngeal arches were evolutionarily lost, an examination of cranial nerve innervation of the arches is useful as specific nerves innervate specific arches and their derivatives. To generate useful data it was necessary to examine arch innervation across vertebrate species from different clades that have varying numbers of pharyngeal arches. All embryos examined were at a similar developmental stage and had developed all of their pharyngeal arches. Amniotes have five pharyngeal arches. Immunofluorescence of developing cranial nerves in the chick (n=3) and mouse (n=3) reveals that the first arch is innervated by the maxillary and mandibular branches of CNV, the second arch is innervated by CNVII, the third arch by CNIX, and the fourth and sixth arches by CNX (Figure 5.1. A, B). The hypoglossal nerve (CNXII) is also evident in both the chick and mouse (Figure 5.1. A, B). It originates from the occipital somites and hooks around the posterior pharynx toward the antero-ventral surface to the pharyngeal arches. This morphology of CNXII is evident across all vertebrate species examined, and can clearly be seen in both the shark and lamprey also (Figure 5.1. C, D). The dogfish (n=4) is a chondrichthyan and has seven pharyngeal arches. The first three arches are innervated by CNV, CNVII and CNIX respectively (Figure 5.1. C). Where in amniotes the posterior two arches are innervated by CNX (Figure 5.1. A, B), the posterior four arches seen in this species are innervated by CNX. The roots of CNIX and CNX are both found posterior to the otic vesicle and each is clearly labelled to differentiate between these two nerves (Figure 5.1.). Examination of the innervation pattern of lamprey (n=3), which have nine pharyngeal arches, reveals a similar result. Again, their first three arches have 149 CNV, CNVII and CNIX innervation respectively, and all six of their posterior pharyngeal arches receive CNX innervation (Figure 5.1. D). The lamprey therefore has six arches with vagal innervation, the shark has four arches with vagal innervation, and amniotes have two arches with vagal innervation. All of these arches are adjacent to one another, and all are located in the posterior pharynx. This provides good anatomical evidence for a reduction in the number of arches to have occurred posteriorly. To determine whether they have been lost from the anterior, middle or posterior part of this posterior region, it is necessary to look at molecular data to establish whether the arches are regionalised. I have therefore analysed Hox gene expression to determine if they impart an identity onto the pharyngeal pouches, thereby demarcating anatomical boundaries within the pharynx. 5.2.2 Hox gene expression in the pharyngeal pouches of amniotes To determine whether Hox gene expression correlates with the pharyngeal pouches, demarcating anterior and posterior boundaries within the pharyngeal endoderm thereby regionalising the pharynx, I have used in situ hybridisation on whole-mount chick embryos prior to sectioning in the coronal plane to visualise Hox gene expression specifically in the endoderm of the pharynx. As Hox genes generally follow a rule of colinearity, I wanted to determine whether this was true within the pharyngeal endoderm. Therefore, I first looked for Hoxb1 expression which, if it follows the rule of colinearity, should be expressed most anteriorly. However, this was not the case as expression of Hoxb1 is evident in the posterior pharyngeal endoderm (Figure 5.2. G, N). I will discuss this further later on in the chapter. I looked next at the expression of paralogous group 2, and found an anterior limit of Hoxa2 expression within the second pharyngeal pouch (n=5; Figure 5.2. A, H). The anterior limit of Hox gene expression within the second pouch provides an axial marker demarcating the border between the anterior and posterior parts of 150 Figure 5.1. Cranial nerve innervation of pharyngeal arches reveals region where pharyngeal arch reduction has occurred in vertebrates All embryos are at the same developmental stage and have developed their full complement of pharyngeal arches. In each species the first arch is innervated by CNV, the second arch by CNVII, the third arch by CNIX, posterior arches by CNX, and CNXII hooks around the posterior pharynx. (A-B) The chick and mouse (amniotes) have 5 pharyngeal arches and the posterior two are innervated by CNX. (C) The shark (a chondrichthyan) has 7 arches, of which the posterior four are innervated by CNX. (D) The lamprey (an agnathan) has 9 pharyngeal arches of which the posterior 6 are innervated by CNX. Therefore CNX innervated arches from the posterior pharynx must be the ones that have been lost, although whether they have been lost from the anterior, middle or posterior part of the this region is indeterminable from this data. Numbers 1-9 represent pharyngeal arch number. 151 152 the pharynx and supporting the differences discussed previously in the way these two regions develop. It may be the absence of Hox gene expression that controls the initiation of a different morphogenetic program in the first pouch when compared with posterior pouches, providing a potential explanation as to why the first pouch/cleft interface does not follow the same general program of development as the others (see Chapter 3 and Chapter 4). I next analysed the expression of paralogous group 3. Hoxa3 expression is evident in the pharyngeal endoderm and has an anterior limit in the third pharyngeal pouch, but only for a short space of time. At stage 17 Hoxa3 expression is seen in the third pouch (n=3; Figure 5.2. B, I), but is subsequently switched off by stage 19/20 (n=3), when the third pouch clearly does not express Hoxa3 (Figure 5.2. C, J). At this stage the endodermal pouch is bulging out through the overlying ectoderm and starting to contribute to the external surface of the embryo, clearly showing up against a background of third arch mesenchymal expression (Figure 5.2. J; black arrows). I also analysed Hoxb3 expression (n=9) but this gene is not seen in the third pouch or any of the pharyngeal endoderm, although some expression is seen in the fourth arch mesenchyme (Figure 5.2. D, K). Paralogous group 4 was analysed next. No expression is seen in the pharyngeal pouches, although Hoxb4 is evident in the endoderm of the posterior pharynx (n=11; Figure 5.2. E, L; black arrows).Therefore the fourth pharyngeal pouch does not act as an anterior boundary for Hox gene expression. Paralogous group 5 member Hoxb5 also did not show any expression in any of the pouches, as well as the pharyngeal endoderm or any of the arches (n=4; Figure 5.2. F, M). As mentioned earlier, Hoxb1 expression is seen in the most posterior pharyngeal pouch (n=9), which at this stage of development is the ‘bud’ elongating from the posterior part of the fourth pouch (Figure 5.2. G, N). Therefore Hoxb1 is marking the most posterior pharyngeal pouch, and thereby acting as the anatomical border for the posterior pharynx. By doing this it breaks its rule of colinearity by being expressed most posteriorly rather than most anteriorly as might be expected given its 3’ position on the chromosome compared with the other Hox genes described 153 here, although it also does this in the hindbrain (Keynes and Lumsden, 1990, Wilkinson et al., 1989). A pattern of Hox gene expression therefore aligns with particular anatomical boundaries to regionalise the pharynx, as opposed to a different Hox gene displaying nested expression within each pharyngeal pouch. These boundaries appear to separate the anterior and posterior pharyngeal arches, with an anterior limit of Hoxa2 expression in the second pouch separating the first two arches and first pouch from those posteriorly. The third pharyngeal pouch expresses Hoxa3 at an earlier stage of development but later switches off in the chick, and so it is unclear what role this gene plays in demarcating an anatomical boundary within the pharynx but will be discussed further below. Hoxb1 however is clearly seen in the posterior pharyngeal pouch, demarcating the posterior border of the pharynx. These results begin to illuminate where the pharyngeal arches may have been reduced. All the posterior pharyngeal pouches fall within the domain flanked anteriorly by Hoxa2 expression and posteriorly by Hoxb1. Therefore, the pharyngeal arches have likely been lost from somewhere within this region. Combined with the arch innervation data, this forms a strong case for a reduction in arch number to have occurred from the posterior pharynx. The basal gnathostome, the shark, has more pharyngeal arches than the chick, and its posterior arches will be retained in their iterated form into adulthood to develop into the gills. In order to determine whether the same pattern of Hox expression is seen in the pouches of a basal gnathostome, I have analysed Hox expression in the endoderm of the shark pharynx. This will reveal whether the same pattern of Hox gene expression is seen at the same anatomical boundaries within the pharynx, regardless of the number of ‘segments’ or arches present within each region, thereby solidifying my hypothesis for posterior arch reduction. 154 5.2.3 Hox gene expression with the pharyngeal pouches in gnathostomes I chose to analyse Hox expression in the pharyngeal endoderm of a basal gnathostome, the chondrichthyan shark, using the same species that I have been throughout this entire study. I collaborated with Patrick Laurenti, corresponding author on the study of Hox expression in the shark by Oulion et al. (2011), who kindly sent me some embryos that had already been stained for Hoxa2, Hoxa3 and Hoxb1 expression. I was therefore able to section these embryos and analyse this complement of Hox gene expression in the shark for my comparative work. Expression was examined at two different stages, and Hox gene expression is higher at the older stages. At St20/21, Hoxa2 expression is very strong in the mesenchyme of the second pharyngeal arch, yet the second pouch endoderm shows an equal amount of expression to the rest of the tissue seen in this image (Figure 5.3. A). The St23/24 embryo is clearer, with no expression at all in the first arch or pouch, strong expression in the second arch mesenchyme, and none in the overlying ectoderm, although expression is seen in the posterior second pharyngeal pouch endoderm and more weakly in the endoderm posterior to here (Figure 5.3. D). This suggests that there is Hoxa2 expression in at least part of the second pharyngeal pouch at earlier stages too. However, this expression at this stage of development does show Hox gene alignment with the second pharyngeal pouch, and is the most anterior limit of Hox gene expression seen in the pharyngeal endoderm matching observations from the chick. This suggests Hoxa2 expression marking the anterior boundary of the posterior pharyngeal pouches is conserved. Hoxa3 expression is evident in the mesenchyme of third pharyngeal arch, yet the labelling of the endoderm is uniform with the rest of the tissue seen at the younger stage examined (Figure 5.3. B). At stage 23/24 expression is clear in the arch mesenchyme of the third arch and those posterior, but there is a lack of Hoxa3 expression in any of the endoderm through the pharyngeal region (Figure 5.3. E). This correlates with the stage 19/20 chick Hoxa3 expression data, 155 Figure 5.2. Hox gene expression in chick pharyngeal pouches All embryos are stage 20. The first pharyngeal pouch is devoid of any Hox expression. (A, H) The second pouch marks the most anterior limit for Hox expression. Hoxa2 is expressed here (asterisk in A, arrows in H), as well as in the endoderm posteriorly. (B, C, I, J) The third pouch expresses Hoxa3 at stage 17 (asterisk in B, arrows in I), although by stage 19/20 this expression is no longer evident (asterisk in D, arrows in K). Hoxb3 is not expressed in the third pouch (asterisk in D, arrows in K), or any of the pharyngeal endoderm. (E, F, L, M) Hoxb4 is evident in the posterior pharynx but not in the fourth pharyngeal pouches (asterisk in E, arrows in L), while no Hoxb5 expression is seen at all in the arches or pouches (F, M). (G, N) Hoxb1 expression is seen in the most posterior fourth pouch (asterisk in G, arrows in N), and breaks the rule of colinearity by being expressed here rather than anteriorly. (O) Schematic representation of a lateral view of a chick embryos shown in (A-G). The dotted line shows the plane of the coronal section shown in (P). (P) Representation of the section seen in images (H-N). ov – otic vesicle; PA – pharyngeal arch; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 156 157 although whether this gene is expressed early in the shark and down-regulated later in development is unclear. Hoxb1 expression is very clear in the shark pharyngeal endoderm. At stage 20/21, Hoxb1 is clearly evident only in the endoderm of the posterior pharyngeal pouch, which at this stage is the third pouch. Similarly, in the stage 23/24 embryo Hoxb1 expression is evident only in the posterior pharyngeal pouch endoderm, which at this stage is the fifth pouch. Previous Hox expression seen in the pouches in the chick or the shark has always aligned with a particular pharyngeal pouch. However, Hoxb1 is aligning with the most posterior pouch present at a given stage, regardless of the number of pouches present. Its expression therefore appears to be dynamic and acts to demarcate the posterior boundary of the pharyngeal endoderm. This has never been shown before, so an analysis of Hoxb1 expression across various different stages of shark and chick development was undertaken to determine whether this is seen throughout pharyngeal development. If so, this dynamic expression pattern of Hoxb1 for demarcating the posterior border of the pharyngeal region must also be conserved. 5.2.4 Transient Hoxb1 expression marks the anatomical border for the posterior pharynx As the pharyngeal arches develop along the A-P axis, with posterior arches forming later than those anteriorly, identifying the gene responsible for specifying the posterior pharynx should help uncover what is controlling when to continue or stop forming arches. As Hoxb1 is only expressed in the most posterior pharyngeal pouch and appears to be expressed here regardless of the number of pouches present at a particular stage of development, this could potentially be what is regulating when to stop forming arches, thereby controlling the number that develop. To determine whether this observation in the shark was real or not, a more comprehensive analysis of Hoxb1 expression was conducted across various different stages. The same analysis was conducted in the chick and show that this expression is conserved. 158 At stage 21 (n=3) the shark has three fully formed pouches while the fourth one is beginning to develop. Here, the third and most posterior fully formed pouch expresses Hoxb1, although the newly developing fourth pouch expresses it more strongly implying expression is being down-regulated anteriorly or transferred posteriorly (Figure 5.4. E). By stage 22 (n=3), the third pouch no longer expresses Hoxb1 at all and the most posterior fully formed fourth pouch expresses Hoxb1 with a gradient; the anterior portion of the pouch shows weak expression, while the posterior part expresses it more strongly with the strongest expression evident in the newly forming fifth pharyngeal pouch and posterior endoderm (Figure 5.4. F). At stage 23 (n=3) the sixth and final pouch has begun elongating toward the ectoderm, and a similar gradient of expression can be seen along the fifth pouch toward the sixth pouch as described between the fourth and fifth pouch at stage 22 (Figure 5.4. G). This expression is further refined at stage 25 (n=3) when Hoxb1 is evident in the endoderm posterior to the sixth pouch where the seventh pouch has begun to form (Figure 5.4. H). Comparing this data with the chick reveals the same expression dynamics are employed in amniotes too. In the chick, the first two pharyngeal pouches arise simultaneously at stage 13, and in situ hybridisation reveals Hoxb1 expression in the most posterior pouch at this stage, which is the second pouch (Figure 5.4. A). Hoxb1 expression has never been reported in the second pouch endoderm previously as it has always been associated with the posterior pharynx. By the time the third pouch has developed at stage 15, expression of Hoxb1 has shifted from the second pouch to the third pouch, the now most posterior aspect of the pharynx (Figure 5.4. B). No expression is evident in the second pouch at this stage, although the mechanism by which transience of Hoxb1 is employed is currently unknown. As the fourth pharyngeal pouch begins developing at stage 17, budding off the posterior pharyngeal endoderm, strong Hoxb1 expression is again shifted to this region (Figure 5.4. C), and by stage 19 no Hoxb1 expression is evident in the third pouch at all (Figure 5.4. D). By this stage however the fourth pouch has fully developed, and the bud off the posterior aspect of the fourth pouch is now strongly expressing Hoxb1 (Figure 5.4. D). 159 Figure 5.3. Hox gene expression in dogfish pharyngeal pouches Embryos were stained using in situ hybridisation by Patrick Laurenti (Oulion et al., 2011) and were sent to me for sectioning and endodermal analysis. (A) Earlier expression of Hoxa2 is particularly strong in the second pharyngeal arch mesenchyme, but expression appears in all tissues making it difficult to determine whether it is in the pouch endoderm. (D) Older stage expression looks more restricted, with expression evident in the posterior second pouch endoderm, as well as the second arch mesenchyme (D). Black arrows point to the second pharyngeal pouches in both stages shown (A, D). (B, E) Hoxa3 expression is evident in arch mesenchyme with an anterior border of the third arch, although background staining makes it difficult again to determine whether any specific expression in the pouch endoderm is seen. Black arrows point third pharyngeal pouches. (C, F) Hoxb1 expression is very clearly in the posterior pharyngeal pouch endoderm at both stages (black arrows). It is expressed in the posterior pouch regardless of the number of pouches present, aligning with the posterior limit of the pharyngeal pouches. (G) Schematic representation of a lateral view of an embryo with a black dotted line showing the plane of coronal section shown in (H). (H) Representation of the pharyngeal morphology depicted in images (A-F). ov – otic vesicle; PA – pharyngeal arch; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 160 161 Figure 5.4. Conserved transient Hoxb1 expression in posterior pharynx of chick and shark (A-D) Sections of chick embryos following in situ hybridisation using a Hoxb1 probe at various stages of development reveals expression in the posterior pouch regardless of the number of pouches present. At St13 in the chick the first two pouches have developed, and Hoxb1 expression is evident in the second pharyngeal pouch which is the most posterior one present at that time (A; black arrows). By stage 15, the third pouch has developed and expression has moved from the second to the third pouch (B; black arrows). At stage 17 the fourth pharyngeal pouch can be seen budding off from the posterior pharyngeal endoderm and expression here is very strong (C; black arrows), although still evident in the third pharyngeal pouch. By stage 19 the fourth pouch has developed properly and the bud off the posterior aspect of the fourth pouch is seen strongly expressing Hoxb1, marking the most posterior border of pharyngeal endoderm (D; black arrows). (E-H) A similar pattern of Hoxb1 expression is seen in the shark. At St21 the third pouch weakly expresses Hoxb1 whereas the developing fourth pouch has strong expression (E; black arrows). By St22 this pouch the third pouch no longer expresses Hoxb1 at all, with expression in the fourth pouch limited to the posterior border while strong expression is seen in the developing fifth pouch (F; black arrows). The sixth pouch is developing at stage 23 and expresses Hoxb1 (G; black arrows), while only the posterior border of the fifth pouch expresses it now. By stage 25, all six pharyngeal pouches are fully developed and Hoxb1 expression is restricted to the posterior border of the sixth pouch and the posterior pharyngeal endoderm (H; black arrows). Both chick and shark show the same dynamic pattern of Hoxb1 expression, suggesting this mechanism is conserved across vertebrates and acts as the anatomical boundary for the posterior pharynx. (I) Schematic representation of a lateral view of an embryo. The black dotted line represents the plane of the coronal section shown in (J). (J) Representation of arch morphology seen in (A-H). ov – otic vesicle; PA – pharyngeal arch; pc – pharyngeal cleft; pp – pharyngeal pouch; red – ectoderm; green – endoderm; blue - mesoderm 162 163 This data reveals that Hoxb1 is always expressed in the most posterior pouch of the chick, regardless of the number of pouches present at that stage of development. Generally Hox genes are known for demarcating anatomical boundaries regardless of the number of ‘segments’ within each domain. However, Hoxb1 functions with a dynamic mode of expression that ‘moves’ from segment to segment during development. I have also shown that this expression dynamic of Hoxb1 is conserved given that it is seen in both in a basal gnathostome and an amniote, therefore helping to elucidate how the pharynx is patterned and how arches were lost with the emergence of gnathostomes and amniotes. This method of reduction, or possibly addition, is likely also employed to control the varying numbers of arches seen in different species within certain groups, i.e. hagfish. 164 5.3 Discussion In order to understand where a loss of pharyngeal arches has occurred with vertebrate evolution, it is important to first decipher how the pharyngeal arches are patterned. It has long been known a conserved pattern of Hox gene expression exists in the pharyngeal region (Hunt et al., 1991a, Oulion et al., 2011, Hunt et al., 1995). However, an extensive analysis of Hox expression specifically in the endoderm has not been conducted previously, despite mounting evidence this is the key instructive tissue of this region (Veitch et al., 1999, Piotrowski and Nusslein-Volhard, 2000). My study has revealed that Hox genes align with the pharyngeal pouches, and this data can be used to uncover where within the pharyngeal region the number of arches has been reduced. Combined with anatomical evidence of nerve innervation, it is apparent that this reduction has occurred from the posterior pharynx. 5.3.1 A conserved Hox code aligns with the pharyngeal pouches in the vertebrate pharynx Regionalised Hox expression in the pharyngeal endoderm is conserved across vertebrates, as comparison between a basal gnathostome, the shark, and an amniote, the chick, reveals (see Figure 5.5 for schematic). In both the chick and shark, the first pharyngeal pouch has no Hox expression, as has been shown also in the first pharyngeal arch. In amniotes the first pouch contributes to middle ear structures (Grevellec and Tucker, 2010), while in the shark it contributes to the spiracular organ (Barry et al., 1988). The amniote middle ear evolved from the spiracular organ, and these structures have recently been proven to be homologous (O'Neill et al., 2012), therefore sharing a common molecular program of development which is likely regulated by a lack of Hox expression in this region. The anterior limit of Hox expression in the shark and chick pharyngeal endoderm is Hoxa2 within the second pharyngeal pouch, although Hoxa2 is also expressed 165 in the second arch mesenchyme. The first pharyngeal arch will form the jaw and the second arch will form hyoid structures, and studies have shown Hoxa2 is required for the formation of hyoid structures (Rijli et al., 1993). These two arches are different from those posteriorly, which will develop into gills in the shark or the neck in the chick. Therefore Hox gene expression in the second pouch represents the anatomical boundary separating the anterior two arches, splitting the pharyngeal apparatus into anterior and posterior portions. This also provides an explanation as to why the first two arches and first pouch behave so differently to those posteriorly. The third pharyngeal arch expresses Hoxa3 in the mouse (Hunt et al., 1991a, Manley and Capecchi, 1995), chick (Hunt et al., 1995), and shark (Oulion et al., 2011). My results show Hoxa3 expression in the third pharyngeal pouch at stage 17, although it had disappeared by stage 19/20. Following the cell lineage tracing data in Chapter 4, it is evident the pharyngeal pouch breaks through to the external surface and the Hoxa3 negative third pouch shown in the data here displays the same tissue morphology. These data substantiate the finding that Hoxa3 is not expressed in the third pharyngeal pouch of the chick at this later stage. Hoxa3 is important for the formation of third pouch-derived structures, the thymus and parathyroid glands (Manley and Capecchi, 1995, Manley and Capecchi, 1998, Chisaka and Capecchi, 1991), although its expression is possibly only required early on, providing a possible explanation for its later downregulation once downstream pathways for development of these glands have been initiated. Following reports of Hox4 paralogues being expressed in the fourth pharyngeal arch (Hunt et al., 1991a, Hunt et al., 1995), it was expected if any Hox expression was to be detected in the fourth pharyngeal pouch it would be Hoxb4. However, this was not evident in the fourth pharyngeal pouches, although expression is present in the pharyngeal endoderm caudal to here. This data is consistent with what has been reported in mouse and shark, as the only Hox4 paralogue expressed in the fourth arch is Hoxd4, whereas Hoxa4 and Hoxb4 are expressed caudally (Minoux et al., 2009, Oulion et al., 2011). Further analysis for Hoxd4 expression 166 in all these species would reveal whether this gene is expressed in the fourth pouch and whether its expression is therefore conserved in this region. My analysis of Hox gene expression within the pharyngeal pouches reveals a conserved pattern of expression which may be responsible for regionalisation of the pharynx within gnathostomes, although data from an even more basal vertebrate, the lamprey, further indicates conservation of this pattern of Hox gene expression in vertebrate pharyngeal endoderm. Cohn (2002) reported Hox expression in the first arch of the lamprey species, Lampetra fluviatilis, suggesting a loss of Hox expression in this arch is linked to the evolution of feeding apparatus, the jaw, in gnathostomes. The grafting of Hox-expressing neural crest into the chick first arch resulted in no jaw forming, while down-regulating Hox expression in the second arch lead to jaw formation, supporting this hypothesis (Couly et al., 1998, Rijli et al., 1993). However, Takio et al. (2004) pointed out that a different species of lamprey, Lethenteron japonicum, does not have Hox expression in the first arch and therefore the initial finding by Cohn (2002) does not represent a general feature of agnathans. An absence of Hox expression in the first pouch probably indicates a difference in the highly specialised structures formed by the first arch and pouch rather than a link as such to the specific formation of jaws. Takio et al. (2007) then reported a Hox code regionalising the lamprey pharynx. They reported no Hox expression in the first arch, with expression of the paralogue group Hox2 (PG2) forming the anterior limit of Hox expression within the pharynx aligning with the second arch, PG3 expression in the third arch, and LjHox1w expression in the posterior pharyngeal endoderm. This is almost identical to the pattern of expression I have reported in gnathostomes, indicating the pharyngeal apparatus was regionalised prior to the evolution of jaws. 167 Figure 5.5. A conserved pattern of Hox gene expression aligns with the pharyngeal pouches of gnathostomes Schematic representation of Hox gene expression in the pharyngeal pouches of a generalised gnathostome throughout pharyngeal development. (A) Schematic representation of initial pharyngeal pouch formation beginning with the simultaneous budding off the pharyngeal endoderm of the first and second pharyngeal pouches. Hoxb1 aligns with the second, most posterior pharyngeal pouch. (B) Schematic representation of a middle state of pharyngeal development with four pharyngeal pouches present. Hoxa2 is now expressed with an anterior limit in the second pouch and Hoxa3 expression is evident with an anterior limit at the third pouch. Hoxb1 expression is no longer seen with an anterior limit in the second pouch although it is present in the most posterior pouch, which at this stage is the fourth. (C) Once all pouches are developed, the anterior limit of Hoxa2 expression still aligns with the second pouch while Hoxa3 expression is no longer seen in the endoderm, representing its transiency as described in the chick. Hoxb1 expression again no longer aligns with the fourth pouch but does still align with the most posterior pouch, which at this stage is the sixth pouch. This represents a conserved dynamic expression pattern of Hoxb1 in the most posterior pouch present of gnathostomes at any given time point during pharyngeal pouch development irrespective of the number of pouches that have formed at that time. (pp = pharyngeal pouch) 168 169 5.3.2 Dynamic and transient Hoxb1 expression demarcates the posterior pharynx Detection of Hoxb1 in the chick fourth pharyngeal pouch has been reported before (Wendling et al., 2000). However, an analysis of Hoxb1 expression in the shark at two different developmental time points revealed Hoxb1 was expressed in the most posterior pouch present at any particular stage, despite the number of pouches that had formed by that time. To establish whether Hoxb1 is a marker specifically for fourth pouch endoderm in the chick, or whether it aligns with the posterior boundary of the pharynx even when not fully developed, I analysed Hoxb1 expression over various different developmental stages in both the chick and shark revealing that it does indeed align with the most posterior pouch present at any stage of development (see Figure 5.5 for schematic representation). This report describes two previously unknown aspects about Hoxb1 expression during pharyngeal development: 1) its expression dynamic is transient, moving posteriorly with each developing pouch, and 2) it defines the posterior pharynx regardless of the extent of its development. Hoxa1 and Hoxb1 are direct targets of RA signalling, which is a conserved feature in vertebrates (Wendling et al., 2000, Marshall et al., 1996). RA signalling has been shown to be responsible for patterning the endoderm along the entire A-P axis in chick, potentially forming a diffusion gradient across the level of the pharyngeal arches (Bayha et al., 2009). Raldh encodes a retinaldehyde dehydrogenase which synthesises RA, and Raldh2 knockout mice display normal development of the first two pharyngeal arches but impaired development of posterior arches due to a failure of the pharyngeal pouches to bud off the pharyngeal endoderm (Niederreither et al., 2003, Niederreither et al., 1999). Similarly, zebrafish neckless (nls) mutants carry a missense mutation of raldh2 causing a comparable phenotype (Begemann et al., 2001). Treatment of mouse embryos using a pan-RA receptor (RAR) antagonist, which blocks all RAR subtypes, caused similar defects as was also seen in vitamin-A deficient quails (Wendling et al., 2000, Quinlan et al., 2002). It was found that RALDH2 is expressed in the mesoderm flanking the pharyngeal endoderm with an anterior limit at the second pharyngeal pouch, providing a source of RA signalling for the 170 endoderm and offering an explanation as to why the first and second pharyngeal arches are unaffected by reduced RA signalling (Quinlan et al., 2002). Hoxa1 and Hoxb1 contain RA response elements (RAREs) in their regulatory regions, and are therefore regulated by RA signalling (Gavalas et al., 1998, Marshall et al., 1996). Interestingly, response to a reduction of RA is time dependent and appears to coincide with the onset of Hox expression, as when it is reduced later in development no effect is seen (Wendling et al., 2000, Marshall et al., 1996). If no RA signalling is present the caudal arches do not form, Hoxa1 expression is reduced and no Hoxb1 expression is evident (Wendling et al., 2000). The second pharyngeal pouch therefore becomes the most posterior pouch present, but has been shown to retain its second pouch identity as it still expresses Pax1 and Shh, which are not expressed in fourth or posterior pouch endoderm (Wendling et al., 2000, Quinlan et al., 2002). Tbx1 expression in caudal pharyngeal endoderm has also been shown to be crucial for appropriate development of the posterior pharynx. The zebrafish vgo mutation affects the tbx1 gene and results in the failure of the pharyngeal endoderm to segment into pharyngeal pouches 2-6, causing abnormal fusion of neural crest-derived cartilages and impaired thymus gland development (Piotrowski and Nusslein-Volhard, 2000, Piotrowski et al., 2003). Tbx1 knockout mice display ectopic anterior expression of RA signalling causing ectopic anterior activation of Hoxb1 and posteriorisation of the pharyngeal region, indicating Tbx1 somehow regulates RA signalling (Guris et al., 2006). This data suggests that RA is required to activate Hox1 expression in the caudal endoderm, which in turn activates specific genetic cascades to assign a posterior identity to surrounding tissues, and that this posterior domain of expression is restricted by the expression of Tbx1. The mechanism by which Hoxb1 is expressed only in the posterior pouch present at any particular stage is likely due to the levels of RA present. As the embryo grows and elongates, the anterior part moves further away from the more posteriorly located source of RA signalling, ensuring Hoxb1 expression is always expressed only in the posterior pharyngeal endoderm. Hox genes are also known to auto- and cross-regulate one another for the establishment and maintenance of 171 their expression patterns, such as during hindbrain development, and this could also be true in the pharyngeal endoderm (Maconochie et al., 1996, Wong et al., 2011, Fox, 2000). For example, Hoxb1 has been shown to auto-regulate itself to maintain its expression in r4 of the hindbrain (Ferretti et al., 2005, Popperl et al., 1995), and likely acts in concert with repressive regulatory mechanisms to restrict its expression to this appropriate domain. Hoxb3 expression has been shown to repress Hoxb1 in the hindbrain (Wong et al., 2011), and when all Hox3 genes are repressed, the territory of Hoxb1 expands to r6 of the hindbrain (Gaufo et al., 2003). It is therefore possible that up-regulation of Hoxa3 in the third pharyngeal pouch could repress Hoxb1 function here. Hoxb1 has also been shown to directly activate Hoxa2 expression in r4 of the hindbrain of chick and mouse (Tumpel et al., 2007), and ectopic hoxb1a expression in r2 of zebrafish induces ectopic activation of hoxa2 (Hunter and Prince, 2002), providing a potential mechanism for the onset of Hoxa2 activity in the second pharyngeal pouch. Further investigation into cross-regulatory mechanisms between these Hox genes would further elucidate how Hoxb1 expression is controlled so it is only ever seen in the posterior pouch, and how it interacts with other Hox genes to regulate their expression, or indeed how other Hox genes regulate each other within the pharyngeal endoderm. 5.3.3 Pharyngeal arch reduction has occurred from the posterior pharynx Analysing pharyngeal arch innervation in vertebrate species has revealed anatomical evidence for where within the pharynx the number of arches has been reduced. In normal embryos, certain cranial nerves innervate specific pharyngeal arches. Basal vertebrates, the jawless lamprey, have 9 pharyngeal arches the first of which is innervated by CNV, the second by CNVII, the third by CNIX, and the six posterior arches by CNX. When compared with more derived species that have a decreased number of pharyngeal arches, a corresponding decrease in the number of arches innervated by CNX is evident. For example, the shark has 7 172 arches and amniotes have 5, and while both have identical innervation of the first three arches to lamprey, the posterior 4 arches in the shark and 2 arches in amniotes receive CNX innervation. This can be interpreted in terms of pharyngeal arch reduction that the posterior arches have been lost, as amniotes display fewer pharyngeal arches with CNX innervation than those of more basal species, while innervation of anterior arches remains the same. This indicates that the arches were lost from somewhere within this group of posterior arches, although whether they were lost from the anterior, posterior or internal portion of this group is not entirely clear. To help refine the location of arch reduction in the posterior pharynx, the analysis of the pattern of Hox gene expression in the pharyngeal endoderm can provide molecular data contributing to this investigation. This analysis revealed regionalisation of the pharyngeal endoderm which potentially assigns an identity to the adjacent pharyngeal arches within each region to provide the correct environment for appropriate cranial nerve innervation. As described already, the anterior limit of Hox expression lies within the second pharyngeal pouch, generating a unique Hox-negative environment in the first arch. This is likely directed by the Hox-negative first pouch which may be necessary for correct innervation by CNV, as well as the correct development of other structures associated with this arch (Rijli et al., 1993, Gendron-Maguire et al., 1993, Hunter and Prince, 2002). The second arch is bordered by the Hoxnegative first pouch anteriorly and Hoxa2-expressing second pouch posteriorly. This is likely generating the correct environment in the second arch for correct CNVII innervation, as its size is reduced and its location altered in Hoxa2 knockout mice (Rijli et al., 1993, Gendron-Maguire et al., 1993). Similarly, the third pharyngeal pouch expresses Hoxa3, so it is likely the combination of the Hoxa2-expressing second pouch at its anterior border and Hoxa3-expressing third pouch at its posterior border that generates the correct environment in the third arch for CNIX innervation. Support for this also comes from Hoxa2 knockout mice, where the proximal portion of CNIX was absent (Gendron-Maguire et al., 1993), and from Hoxa3 knockout mice who are either also missing the proximal portion of their CNIX or it has fused to CNX (Manley and Capecchi, 1997). The 173 next limit of Hox gene expression is Hoxb1 in the most posterior pharyngeal pouch and endoderm, marking the posterior anatomical border of the pharynx. Each ‘segment’, or arch, within this domain is innervated by CNX, and so these arches are specified by their location within the region flanked anteriorly by the Hoxa3-expressing third pouch and posteriorly by the Hoxb1-expressing posterior pouch. This expression pattern is important for specifying appropriate cranial nerve formation and innervation in the posterior pharynx. Hoxa3 knockout mice often display fused CNIX and CNX neurons (Manley and Capecchi, 1997), while blocking RA signalling, thereby inhibiting Hoxb1 expression, results in fused and much reduced neurons and ganglia of CNIX and CNX (Wendling et al., 2000). It would be interesting to determine whether the anterior and posterior boundaries set up in the anterior pharynx need to lie within directly adjacent pouches, or whether, for example, misexpressing Hoxa3 to a more posteriorly located pouch would result in more arches being innervated by CNIX. This pattern of Hox gene expression and its ability to impart pharyngeal pouch identity is not restricted to the pharynx. Hox genes are responsible for conferring axial positional information in all three germ layers of developing embryos, thereby regionalising the developing body plan by specifying anatomical boundaries. A common pattern of Hox expression is evident within the tetrapod vertebral column, where certain Hox genes are expressed at anatomical boundaries demarcating a transition between regions of the vertebrae irrespective of the species-dependent number of vertebrae present within each section, i.e. Hoxc6 is expressed at the cervical-thoracic boundary even though in chick and mouse these boundaries are separated by 7 somites (Burke et al., 1995). Similarly in the chick and shark pharynx, Hoxb1 is expressed in the posterior pharyngeal pouch even though this region is reduced by two pharyngeal arches in the chick. In the vertebral column, if Hox genes are misexpressed homeotic transformations occur, with paralogous Hoxa6 knockout mice showing a partial posterior transformation of their last cervical vertebra toward a thoracic phenotype (Kostic and Capecchi, 1994). Future experiments would reveal whether similar homeotic transformations would occur in the pharynx as a result of misexpression studies. The only Hox misexpression investigations conducted in this region to date have 174 focused on misexpression in NCCs and arch mesenchyme (Rijli et al., 1993, Pasqualetti et al., 2000, Gendron-Maguire et al., 1993, Hunter and Prince, 2002), so it would be interesting to see what effects misexpression within the pharyngeal endoderm would cause. For example, if Hoxa2 is up-regulated in the first or third pouch endoderm would this posteriorise or anteriorise the pharynx respectively within these regions? Or similarly, if Hoxb1 is constitutively activated in a pouch anterior to the posterior pharyngeal boundary, i.e. in the fourth pouch of a 6pouched shark, would this inhibit the continued formation of pouches and arches posteriorly resulting in a 5-arched shark? It has long been assumed that pharyngeal arches have been lost internally, as evidenced by the nomenclature of amniote pharyngeal arches being numbered one, two, three, four and six (Larsen, 1997). This would imply that Hoxb1 defines a particular pouch as the posterior boundary and extra pouches develop anteriorly to it. However, the combination of my anatomical data with the patterns of Hox gene expression I have detected indicates that new arches develop posteriorly, and therefore in gnathostome species Hoxb1 expression is transient and demarcates the posterior pharyngeal pouch as each new one develops. It is likely something else is acting upstream of Hoxb1 to control its transience within the pharynx and to ultimately restrict it to a particular pouch. This would prevent the development of any more pouches so each species will form the appropriate number, although how this process is controlled remains unknown. Complimentary information for this theory arises from fossil data, where it has been suggested that a pair of small articulating cartilages posterior to the gill arch skeleton in some shark species are rudimentary of an extra gill arch (Compagno, 1999). Therefore, I propose pharyngeal arch reduction has occurred from the posterior pharynx. 175 5.4 Summary In order to determine where a reduction in the number of pharyngeal segments has occurred with vertebrate evolution, I needed to establish what was responsible for patterning the pharyngeal arches. Due to the instructive capacity of the pharyngeal endoderm (Veitch et al., 1999, Piotrowski and Nusslein-Volhard, 2000), it became apparent this tissue is likely responsible for regionalising the pharyngeal apparatus for correct patterning. As Hox genes are responsible for regionalisation of developing embryos and act along the A-P axis, these are good candidate genes for being involved in specification of anatomical boundaries within the pharynx. Following an analysis of various Hox genes within the pharyngeal pouches, I have shown a particular expression pattern exists in this region and that it is conserved across vertebrate species. I have also uncovered a dynamic expression pattern of Hoxb1 demarcating the posterior pharynx throughout its early period of development, and suggested its mechanism of regulation is important in determining the number of pharyngeal arches that are present within a particular species. Combining this molecular data with a comparison of pharyngeal arch cranial nerve innervation across different species from varying phylogenetic groups revealed the pharyngeal arches were likely reduced from the posterior pharynx. This has resulted in the five pharyngeal arches seen in amniote species compared with nine pharyngeal arches seen in the basal lamprey. 176 Chapter 6. Discussion and Conclusions Pharyngeal arch ontogeny is a highly complex process incorporating interactions between various different tissues and molecular signals at different time points throughout the entire development process. However, the processes by which these tissues are properly patterned and formed are not well understood. It has transpired over the last decade or so that endoderm is likely the principal tissue governing development of this region, in particular the pharyngeal pouches which are formed of endoderm and are responsible for segmenting the pharynx. However, little investigation has focused on these structures and how they interact with surrounding tissues. My study has aimed to rectify this gap in the literature by conducting an extensive analysis of the pharyngeal pouches. The morphology of each pharyngeal pouch within the chick, an amniote, have been shown to each be very different from one another, reflecting the unique structures each one will develop into and/or direct surrounding tissues to develop into. This distinct difference in pouch morphology is not apparent in the posterior pouches of gill-bearing species, where each one will develop into a gill slit almost identical in structure. This reflects the true segmentation of the pharyngeal region, which is clearly maintained in basal gnathostomes but modified in amniotes. This segmentation is not at all apparent in adult amniote organisms, yet the early segmentation of the pharynx indicates an evolutionarily conserved process for development of the head and neck regions across vertebrate species. I therefore investigated the morphology of each pharyngeal pouch and how they interact with the overlying ectoderm in the chick, and compared and contrasted these both within this species and across other vertebrate species to assess whether these processes are conserved. I have also looked into whether a pattern of Hox gene expression is present within the pharyngeal pouches, potentially regionalising the pharynx and assigning each pouch and adjacent arches an identity. Finally, I have determined how these processes have regulated an 177 alteration in the number of pharyngeal arches seen across different vertebrate species and where within the pharynx this has occurred. 6.1 Endodermal segmentation is conserved Pharyngeal pouches bud off the pharyngeal endoderm to segment the pharyngeal region, thereby forming the anterior and posterior borders of each pharyngeal arch. Following elongation the pharyngeal pouches make contact with the overlying ectoderm, but a detailed cellular analysis of how these epithelia interact with each other has not been previously conducted. I have shown in Chapter 3 and Chapter 4 that a general program of interaction exists at the endoderm/ectoderm interface and that this mechanism is conserved across vertebrate species. In the shark, a basal gnathostome, when the ectoderm and endoderm make contact their basement membranes fuse and then break down, allowing direct interaction between ectoderm and endoderm cells. However, no tissue intercalation is seen, and instead the interface thins and eventually perforates to form the gill slits. Cell lineage tracing of the endoderm in zebrafish confirmed the location of ectoderm and endoderm cells following perforation of the gills, showing that the endoderm breaks through the ectoderm to lie on the external surface of the embryo. Investigations using endoderm and ectoderm cell lineage tracing in amniote species revealed a similar process, also resulting in an outward movement of the endoderm to make contact with the external surface. It has not been previously shown that the endoderm comes to lie on the external surface of the arches, and the discovery that this occurs even in amniote species, despite their later remodelling of the pharyngeal region, supports the theory that an initial morphogenetic program for gill formation is initiated in all vertebrates and therefore pharyngeal development is conserved. I have also illustrated a distinct difference between the anterior and posterior pharyngeal pouches, both in their morphologies and in their behaviour. As the first pharyngeal arch develops into the jaw, it is not surprising this arch is very different from the others. Similarly, the first pharyngeal pouch is different from 178 those posteriorly, and this remains true across all vertebrate species despite different structures developing from the first pharyngeal pouch in basal gnathostomes and amniotes. However these structures, the spiracular organ and the middle ear, are homologous with one another and I have shown they share a similar initial mechanism of development, despite the fact that the basal gnathostome first pouch will eventually perforate, albeit much later than posterior pouches do, whereas the amniote first pouch does not. The molecular cues governing this continued development of the first pharyngeal pouch within vertebrate species is not well understood, and further investigation into this area will increase an understanding of how formation of this arch and pouch is governed and why it is so different from posterior arches and pouches. One thing that is clear however is that the first two pharyngeal arches and first pharyngeal pouch are not affected by retinoid signalling, whereas caudal pouches and arches are, so this may be a good place to start (Wendling et al., 2000, Quinlan et al., 2002). Another noticeable difference between the anterior and posterior pouches within vertebrate species is the number of posterior pouches and arches present. The first two arches form similar structures in gnathostomes: the first arch forms the jaw while the second arch will form hyoid apparatus for jaw support. Posterior to these two arches are three pharyngeal pouches in amniotes, five pouches in the basal gnathostome, the shark, and seven in lamprey (for schematic see Figure 1.2 in Chapter 1). Although lamprey are jawless vertebrates, their first arch is still very different from those posteriorly and forms the velum, a specialised mouth part. With the emergence of tetrapods, a transition from water to land occurred resulting in a reduction of the number of pharyngeal arches and pouches. This occurred as they no longer contribute to a function in respiration as they do in basal gnathostomes. Despite this difference in the number of pouches and arches present, the initial formation of the posterior pouches follows the same morphogenetic program and supports the conclusion that the pharyngeal pouches across vertebrate species are homologous, despite the huge shift in morphology and function seen in the fully developed organisms within different groups. 179 6.2 A pattern of Hox gene expression in the pharyngeal pouches governs regionalisation of the pharynx To begin to understand how the pharyngeal region became regionalised into anterior and posterior portions, and to determine whether the pharyngeal endoderm is the key tissue involved in patterning the pharyngeal region, I analysed whether a particular pattern of Hox gene expression exists in the pharyngeal pouches, and whether this could be responsible for organising the pharyngeal region. My results revealed that Hox gene expression does indeed align with the pharyngeal pouches and that it may regionalise the pharynx into anterior and posterior portions, rather than assigning a unique identity to each individual pharyngeal pouch. The most anterior limit of Hox expression is Hoxa2 and lies within the second pharyngeal pouch, which is the border between the first two arches and those posteriorly. Hoxa3 is expressed in the third pouch endoderm and posterior, although transiently in the chick, and is important for thymus and parathyroid gland formation (Mulder et al., 1998, Manley and Capecchi, 1995, Manley and Capecchi, 1998), and Hoxb1 is expressed in the posterior pharyngeal pouch endoderm, regardless of the number of pouches present at a particular point of development, thereby marking the posterior boundary of the pharynx. My results therefore show that a common pattern of Hox gene expression is seen across these gnathostome species, and it may well be present within the agnatha too (Takio et al., 2007, Takio et al., 2004). The expression of these Hox genes initiates a cascade of genetic pathways, allowing each region to instruct surrounding tissues for their correct development. This results in an intricately patterned structure where the correct environment is generated within each region for the development or direction of appropriate structures. For example, the first three arches are innervated by specific cranial nerves, CNV, CNVII and CNIX respectively, while all posterior arches, regardless of the species-dependent number present, are innervated by branches of CNX. The corresponding pattern of Hox gene expression in the pharyngeal pouches reveals that the first pouch is devoid of Hox gene expression while the second and third pouches express Hoxa2 and Hoxa3 respectively, and the most posterior pouch expresses Hoxb1. The expression pattern of Hox genes in the 180 pharyngeal pouches clearly reflects the regions where the correct environment is set up so these nerves innervate the appropriate arches. This hypothesis is supported by a study conducted by Wendling et al. (2000) where inhibition of RA signalling, and therefore downstream activation of Hox expression, results in no formation of the caudal arches or pouches and improper formation and innervation of CNIX and CNX in the appropriate arches. This is probably due to a double effect: first, the appropriate pharyngeal pouches were unable to form due to a lack of Hox expression, inhibiting the genetic cascades these initiate, and were therefore unable to impart appropriate signals, including Bmp7 for epibranchial placode formation (Begbie et al., 1999); and second, even if these neurons did still form properly, the correct environment has not been generated to direct appropriate nerves to innervate the correct arches. 181 6.3 A general trend toward a reduction in the number of pharyngeal arches in vertebrates: how and where does this occur? Understanding how the pharynx is regionalised during its early development has assisted in elucidating where within the pharyngeal region segments have been lost over the course of vertebrate evolution, i.e. have they been lost from the anterior or the posterior part of the pharynx? Or has an internal loss occurred? Molecular and embryological analyses have revealed a division of the pharynx into anterior and posterior domains. It is becoming clear that the loss of segments has occurred from the posterior region, although where exactly within the posterior region this has occurred is still a bit unclear. To supplement these investigations, I have looked to anatomical and fossil records to gain insights into the reasons why a reduction in the number of pharyngeal segments has occurred as vertebrates evolved. I have found that this loss has most occurred from the posterior pharynx, and have supported this theory using evidence derived from multiple facets of investigative work including anatomical, embryological, genetic and fossil data, in order to present a well-rounded overview of how these evolutionary changes have occurred. 6.3.1 Why did arch reduction occur? The most basal vertebrate species are jawless and many of these species have a higher number of pharyngeal arches than gnathostome species, with a general trend toward a reduction in arch number occurring with the emergence of the gnathostomes. The reason for this is not clear among species that still form gill structures in later life, as they all serve the same function of respiration. Perhaps the often parasitic agnathans require a larger number of gills because their mouth part functions as a ‘sucker’, creating a vacuum to attach to its food source (often a large fish) and clinging on for long lengths of time. This renders their mouth piece useless for respiration, during which time their gills draw water in as well 182 as expel it (Wilson and Laurent, 2002, Dawson, 1905), and so a higher number of gills would compensate for the loss of the mouth part during feeding. In basal gnathostomes, most species have seven pharyngeal arches and six pouches, with six arches and five pouches developing into gills, although there is some variation in this number. As all these species have a jaw they do not obstruct their mouths for long periods of time during feeding and therefore the mouth can usually participate in respiration, portending a need for fewer pharyngeal arches that will form fewer gills. Additionally, although the first pharyngeal pouch does not form a gill in these species it does form a spiracle which will allow the animal to breathe even when their mouths are occupied during feeding. It is easier to identify a reason for a reduction in arch number with the emergence of tetrapods. As they do not retain their pharyngeal pouches and arches into adulthood as gills, each arch and pouch has undergone extensive modification to form distinct structures, each of which will perform a specific function. This refinement of the pharyngeal apparatus has led to a reduction in the number of arches needed across different vertebrate species. In basal gnathostomes with seven arches, an arrangement of 2+5 is suggested as the first two arches will form the jaw and hyoid apparatus while each of the posterior arches will develop into gills. In tetrapods, Xenopus have a 2+4 arrangement and amniotes have a 2+3 arrangement, as their first two arches still contribute to the jaw and hyoid structures while their posterior arches contribute to throat structures in the adult. In gill-bearing species, as the function of each gill is identical to its neighbour they all bear an identical morphology, and a large number of them is necessary for appropriate respiratory function. It is therefore rational that a conserved pattern of Hox gene expression would align with the anatomical boundaries demarcating this difference between the anterior two arches and the posterior arches, and that another boundary is present in the posterior pharynx indicating the point where it ends, particularly if the number of arches that could potentially be present within this posterior region is variable. Therefore as more derived species evolved this difference in the anterior and posterior domains has been maintained due to the anterior anatomical boundary set up by Hox genes, and modifications to the number of segments has apparently occurred due to ‘moving’ 183 the posterior boundary of the pharynx thereby preventing the formation of more segments. Support for this theory can be found within the fossil record. Palaeontological analyses have linked the spiracle, which assists chondrichthyan species in waterbreathing when their mouths are unavailable, to an air-breathing function in some actinopterygian species, i.e. Polypterus, and sarcopterygian species (Clack, 2007). Strong support for this lies within analyses of tetrapodomorph skeletons, which have a more robust fin skeleton that may have assisted these species in lifting their heads out of the water when in shallow regions, therefore enabling them to breathe air through their spiracle (Clack, 2007). Furthermore, the majority of water-breathing fish species have five gill arches, while the air-breathing basal sarcopterygian species, the lungfish, have only three gill arches of which two do not directly contribute to respiration (Burggren and Johansen, 1987). Therefore, an observation of a general trend toward an increased size of the spiracle alongside a general reduction in the size of the gill chamber has been detected within extant sarcopterygian, tetrapodomorph and early tetrapod species (Coates and Clack, 1991, Clack et al., 2003, Downs et al., 2008, Long et al., 2006, Clack, 2007). Ichthyostega, a basal tetrapod species, has three gills arches and possibly a small fourth one, indicating a reduction in the posterior gill arch size preceded its loss, and that therefore the arches are lost posteriorly as a use for them becomes reduced (Clack, 2007, Clack et al., 2003). 6.3.2 Where did arch reduction occur? To determine whether arch reduction occurred anteriorly, I compared anatomical and developmental data to emphasise that the anterior two arches are conserved in all vertebrates, forming the mandibular and hyoid arches, so called even in agnathan species, due to the conserved neural crest streams that migrate here and the conserved Hox genes these cell express (Horigome et al., 1999, Kuratani et al., 2001). As each of these arches forms distinct and important structures which were not lost during gnathostome evolution, the arches in this anterior region 184 giving rise to these structures also cannot have been lost. Conserved Hox gene expression data in the pharyngeal endoderm regionalising the pharynx also supports this theory. As each pharyngeal pouch lines the anterior and posterior border of each arch, each one of these arches will be patterned by the combination of Hox genes expressed at either side of it. The first pouch is devoid of Hox gene expression, and therefore the first arch receives no information from a Hox-expressing pouch. The second arch is flanked anteriorly by the Hox-negative first pouch and posterior by the Hoxa2-expressing second pouch, so only information from Hoxa2 is imparted onto this arch. The third arch however is lined by the second pouch anteriorly and the Hoxa3-expressing third arch posteriorly, therefore being given an identity via a unique combination of Hox gene expression. The fourth arch is also adjacent to the Hoxa3-expressing third pouch, while all arches posterior to this one are identical until Hoxb1 expression is detected in the posterior pharyngeal pouch, indicating the back of the pharynx. It is unlikely an internal loss has occurred due to the requirement of the fourth arch for expression of Hoxa3 at its anterior border, which could be required to specify the anterior part of this caudal region. A loss of this middle portion would likely prevent the initiation of posterior pharyngeal development, and so the arches also cannot have been lost from this region. My identification of a dynamic mode for the expression of Hoxb1 marking the posterior pharynx indicates that extra arches are added posteriorly during normal development. For example, Hoxb1 marks the posterior pharyngeal pouch in all vertebrate species regardless of pouch number, i.e. the fourth pouch of amniote species or the sixth pouch of a basal gnathostome. Therefore, an earlier halt in arch formation will lead to fewer arches forming, resulting in their general loss. However, what regulates this unique expression pattern of Hoxb1 and controls when its expression should be maintained within a particular pouch to inhibit the formation of further pouches is unknown. As discussed earlier, RA signalling has been shown to induce Hox gene expression, and is crucial for correct patterning and development of the pharyngeal apparatus. It is present from the level of the second pouch and 185 posteriorly but absent from the first two arches and first pouch, therefore if RA signalling is blocked the first two arches and the first pouch form normally but no caudal arches or pouches form (Quinlan et al., 2002, Matt et al., 2003, Wendling et al., 2000). It was shown that no Hoxb1 could be detected in the pharyngeal region of these affected embryos, indicating that they were not able to specify the posterior pharynx and therefore no caudal structures could form (Wendling et al., 2000). It is therefore likely RA plays a key role in regulating this transient Hoxb1 expression to define the posterior pharynx and direct the formation of more arches and further investigation into this area will be key to further understanding how arch numbers are controlled between vertebrate species. This combination of data strongly supports my hypothesis for a general trend toward a reduction in arch number occurring from the posterior pharynx. The smaller size of the posterior gill arch seen in some basal tetrapod species skeletons supports the theory of reduction and eventual loss of this arch in more evolved species. This loss has underpinned a refinement of the structures that develop from each arch, allowing adaptation of the pharynx from functioning in respiration to becoming primarily associated with feeding. 186 6.4 Concluding remarks Haeckel’s Law states that ‘ontogeny recapitulates phylogeny’, and while this is of course not true, it is undeniable that all vertebrate species retain a remarkable similarity in their appearance during the phylotypic stage. This in fact makes perfect sense: evolutionary changes can only be made according to what is initially present, tweaking genetics and altering molecular cues, leading to the reshaping and remodelling of certain structures until eventually they are unrecognisable from the original model. The fact that certain structures are still present at a specific point in embryogenesis emphasises the conservation seen across vertebrate species, and the pharyngeal arches and pouches represent a key feature within this taxon. Throughout this thesis I have uncovered previously alluded to yet unknown theories about how the pharyngeal region is patterned and develops. All the results found in this thesis complement one another, yet for the sake of clarity each has been presented and discussed separately. Determining that a conserved pattern of Hox gene expression exists within the pharyngeal endoderm aligning with specific pharyngeal pouches confirms the previously believed but never proven notion that this is likely the principal tissue regionalising the entire pharynx and is therefore a step forward in understanding how this region develops properly. It also forms the basis of revealing how this region has evolved and changed both structurally and functionally, how the pharynx has become refined depending upon its function in new environments, and how this refinement has affected the morphology and function of each pharyngeal pouch and arch as they become less a part of a segmented structure and more specialised in their own right. It has also further emphasised the difference seen between the anterior and posterior regions of the pharynx, both morphogenetically and functionally, and has shown that the cause of these differences are likely associated with the regionalisation imparted to the pharynx by the pharyngeal endoderm. This analysis has raised a number of important questions for the future study of this region. It is not well understood how the endoderm imparts information to the 187 surrounding tissues and exactly what signals are involved. Further investigation in this area would elucidate the molecular cues and tissue interactions responsible for shaping the pharynx. Uncovering this would in turn provide more evidence for how modification of these components has led to pharyngeal evolution and combined with more anatomical and fossil evidence can help further solidify the theory of a posterior loss for a reduction in the number of pharyngeal segments. 188 Chapter 7. Bibliography ABU-ISSA, R., SMYTH, G., SMOAK, I., YAMAMURA, K. & MEYERS, E. N. 2002. Fgf8 is required for pharyngeal arch and cardiovascular development in the mouse. Development, 129, 4613-25. ALEXANDER, J. & STAINIER, D. Y. 1999. A molecular pathway leading to endoderm formation in zebrafish. Curr Biol, 9, 1147-57. BAKER, C. V., O'NEILL, P. & MCCOLE, R. B. 2008. Lateral line, otic and epibranchial placodes: developmental and evolutionary links? J Exp Zool B Mol Dev Evol, 310, 370-83. BALLARD, W. W., MELLINGER, J. & LECHENAULT, H. 1993. A series of normal stages for development of Scyliorhinus canicula, the lesser spotted dogfish (Chondrichthyes: Scyliorhinidae). Journal of Experimental Zoology, 267, 318-336. BARRY, M. A., HALL, D. H. & BENNETT, M. V. 1988. The elasmobranch spiracular organ. I. Morphological studies. J Comp Physiol A, 163, 85-92. BAYHA, E., JORGENSEN, M. C., SERUP, P. & GRAPIN-BOTTON, A. 2009. Retinoic acid signaling organizes endodermal organ specification along the entire antero-posterior axis. PLoS One, 4, e5845. BEGBIE, J., BRUNET, J. F., RUBENSTEIN, J. L. & GRAHAM, A. 1999. Induction of the epibranchial placodes. Development, 126, 895-902. BEGEMANN, G., SCHILLING, T. F., RAUCH, G. J., GEISLER, R. & INGHAM, P. W. 2001. The zebrafish neckless mutation reveals a requirement for raldh2 in mesodermal signals that pattern the hindbrain. Development, 128, 3081-94. BOWLES, J., SCHEPERS, G. & KOOPMAN, P. 2000. Phylogeny of the SOX family of developmental transcription factors based on sequence and structural indicators. Dev Biol, 227, 239-55. BRYANT, D. M. & MOSTOV, K. E. 2008. From cells to organs: building polarized tissue. Nat Rev Mol Cell Biol, 9, 887-901. BURGGREN, W. W. & JOHANSEN, K. 1987. Circulation and respiration in lungfishes (Dipnoi). Journal of Morphology Supplement, 1, 217-236. BURKE, A. C., NELSON, C. E., MORGAN, B. A. & TABIN, C. 1995. Hox genes and the evolution of vertebrate axial morphology. Development, 121, 333-46. CALLERY, E. M. & ELINSON, R. P. 2000. Opercular development and ontogenetic re-organization in a direct-developing frog. Dev Genes Evol, 210, 377-81. CALLERY, E. M., FANG, H. & ELINSON, R. P. 2001. Frogs without polliwogs: evolution of anuran direct development. Bioessays, 23, 233-41. 189 CHAPMAN, S. C., COLLIGNON, J., SCHOENWOLF, G. C. & LUMSDEN, A. 2001. Improved method for chick whole-embryo culture using a filter paper carrier. Dev Dyn, 220, 284-9. CHISAKA, O. & CAPECCHI, M. R. 1991. Regionally restricted developmental defects resulting from targeted disruption of the mouse homeobox gene hox-1.5. Nature, 350, 473-9. CHOE, C. P., COLLAZO, A., TRINH LE, A., PAN, L., MOENS, C. B. & CRUMP, J. G. 2013. Wnt-dependent epithelial transitions drive pharyngeal pouch formation. Dev Cell, 24, 296-309. CHUNG, W. S. & STAINIER, D. Y. 2008. Intra-endodermal interactions are required for pancreatic beta cell induction. Dev Cell, 14, 582-93. CLACK, J. A. 2007. Devonian climate change, breathing, and the origin of the tetrapod stem group. Integr Comp Biol, 47, 510-23. CLACK, J. A., AHLBERG, P. E., FINNEY, S. M., DOMINGUEZ ALONSO, P., ROBINSON, J. & KETCHAM, R. A. 2003. A uniquely specialized ear in a very early tetrapod. Nature, 425, 65-9. CLEMENTS, D. & WOODLAND, H. R. 2000. Changes in embryonic cell fate produced by expression of an endodermal transcription factor, Xsox17. Mech Dev, 99, 65-70. COATES, M. I. & CLACK, J. A. 1991. Fish-like gills and breathing in the earliest known tetrapod. Nature, 352, 234-236. COHN, M. J. 2002. Evolutionary biology: lamprey Hox genes and the origin of jaws. Nature, 416, 386-7. COLE, L. K. & ROSS, L. S. 2001. Apoptosis in the developing zebrafish embryo. Dev Biol, 240, 123-42. COMPAGNO, L. J. V. 1999. Endoskeleton. In: HAMLETT, W. C. (ed.) Sharks, skates and rays: the biology of elasmobranch fish. Maryland: The Johns Hopkins University Press. CORDIER, A. C. & HAUMONT, S. M. 1980. Development of thymus, parathyroids, and ultimo-branchial bodies in NMRI and nude mice. Am J Anat, 157, 227-63. COULY, G., CREUZET, S., BENNACEUR, S., VINCENT, C. & LE DOUARIN, N. M. 2002. Interactions between Hox-negative cephalic neural crest cells and the foregut endoderm in patterning the facial skeleton in the vertebrate head. Development, 129, 1061-73. COULY, G., GRAPIN-BOTTON, A., COLTEY, P., RUHIN, B. & LE DOUARIN, N. M. 1998. Determination of the identity of the derivatives of the cephalic neural crest: incompatibility between Hox gene expression and lower jaw development. Development, 125, 3445-59. COULY, G. & LE DOUARIN, N. M. 1990. Head morphogenesis in embryonic avian chimeras: evidence for a segmental pattern in the ectoderm corresponding to the neuromeres. Development, 108, 543-58. 190 CRUMP, J. G., MAVES, L., LAWSON, N. D., WEINSTEIN, B. M. & KIMMEL, C. B. 2004. An essential role for Fgfs in endodermal pouch formation influences later craniofacial skeletal patterning. Development, 131, 5703-16. DAVID, N. B., SAINT-ETIENNE, L., TSANG, M., SCHILLING, T. F. & ROSA, F. M. 2002. Requirement for endoderm and FGF3 in ventral head skeleton formation. Development, 129, 4457-68. DAWSON, J. 1905. The breathing and feeding mechanism of the lampreys II. Biol. Bull., 9, 91-111. DEKKER, E. J., PANNESE, M., HOUTZAGER, E., BONCINELLI, E. & DURSTON, A. 1992. Colinearity in the Xenopus laevis Hox-2 complex. Mechanisms of Development, 40, 3-12. DICKINSON, A. J. & SIVE, H. 2006. Development of the primary mouth in Xenopus laevis. Dev Biol, 295, 700-13. DONOGHUE, P. C. & SANSOM, I. J. 2002. Origin and early evolution of vertebrate skeletonization. Microsc Res Tech, 59, 352-72. DOWNS, J. P., DAESCHLER, E. B., JENKINS, F. A., JR. & SHUBIN, N. H. 2008. The cranial endoskeleton of Tiktaalik roseae. Nature, 455, 925-9. DUBOULE, D. 1994. Temporal colinearity and the phylotypic progression: a basis for the stability of a vertebrate Bauplan and the evolution of morphologies through heterochrony. Dev Suppl, 135-42. DUBOULE, D. & DOLLE, P. 1989. The structural and functional organization of the murine HOX gene family resembles that of Drosophila homeotic genes. EMBO J, 8, 1497-505. DUBOULE, D. & MORATA, G. 1994. Colinearity and functional hierarchy among genes of the homeotic complexes. Trends Genet, 10, 358-64. DUDLEY, J. 1942. The development of the ultimobranchial body of the fowl, Gallus domesticus. American Journal of Anatomy, 71, 65-97. EAMES, B. F., ALLEN, N., YOUNG, J., KAPLAN, A., HELMS, J. A. & SCHNEIDER, R. A. 2007. Skeletogenesis in the swell shark Cephaloscyllium ventriosum. J Anat, 210, 542-54. ENDO, Y. 2012. Chick embryo culture and electroporation. Curr Protoc Cell Biol, Chapter 19, Unit19 15. ENGERT, S., LIAO, W. P., BURTSCHER, I. & LICKERT, H. 2009. Sox17-2AiCre: a knock-in mouse line expressing Cre recombinase in endoderm and vascular endothelial cells. Genesis, 47, 603-10. EOM, D. S., AMARNATH, S., FOGEL, J. L. & AGARWALA, S. 2011. Bone morphogenetic proteins regulate neural tube closure by interacting with the apicobasal polarity pathway. Development, 138, 3179-88. EWALD, A. J., BRENOT, A., DUONG, M., CHAN, B. S. & WERB, Z. 2008. Collective epithelial migration and cell rearrangements drive mammary branching morphogenesis. Dev Cell, 14, 570-81. 191 FAGMAN, H., ANDERSSON, L. & NILSSON, M. 2006. The developing mouse thyroid: embryonic vessel contacts and parenchymal growth pattern during specification, budding, migration, and lobulation. Dev Dyn, 235, 444-55. FAGMAN, H. & NILSSON, M. 2010. Morphogenesis of the thyroid gland. Mol Cell Endocrinol, 323, 35-54. FERRETTI, E., CAMBRONERO, F., TUMPEL, S., LONGOBARDI, E., WIEDEMANN, L. M., BLASI, F. & KRUMLAUF, R. 2005. Hoxb1 enhancer and control of rhombomere 4 expression: complex interplay between PREP1-PBX1-HOXB1 binding sites. Mol Cell Biol, 25, 8541-52. FOX, E. A. 2000. The previously identified r3/r5 repressor may require the cooperation of additional negative elements for rhombomere restriction of Hoxb1. Brain Res Dev Brain Res, 120, 151-64. FRANK, D. U., FOTHERINGHAM, L. K., BREWER, J. A., MUGLIA, L. J., TRISTANI-FIROUZI, M., CAPECCHI, M. R. & MOON, A. M. 2002. An Fgf8 mouse mutant phenocopies human 22q11 deletion syndrome. Development, 129, 4591-603. FRISCH, S. M. 1994. E1a induces the expression of epithelial characteristics. J Cell Biol, 127, 1085-96. FRISCH, S. M. & FRANCIS, H. 1994. Disruption of epithelial cell-matrix interactions induces apoptosis. J Cell Biol, 124, 619-26. FURLOW, J. D., YANG, H. Y., HSU, M., LIM, W., ERMIO, D. J., CHIELLINI, G. & SCANLAN, T. S. 2004. Induction of larval tissue resorption in Xenopus laevis tadpoles by the thyroid hormone receptor agonist GC-1. J Biol Chem, 279, 26555-62. GANS, C. & NORTHCUTT, R. G. 1983. Neural crest and the origin of vertebrates: a new head. Science, 220, 268-73. GARCIA-FERNANDEZ, J. & HOLLAND, P. W. 1994. Archetypal organization of the amphioxus Hox gene cluster. Nature, 370, 563-6. GARCIA-MARTINEZ, V., MACIAS, D., GANAN, Y., GARCIA-LOBO, J. M., FRANCIA, M. V., FERNANDEZ-TERAN, M. A. & HURLE, J. M. 1993. Internucleosomal DNA fragmentation and programmed cell death (apoptosis) in the interdigital tissue of the embryonic chick leg bud. J Cell Sci, 106 ( Pt 1), 201-8. GARG, V., YAMAGISHI, C., HU, T., KATHIRIYA, I. S., YAMAGISHI, H. & SRIVASTAVA, D. 2001. Tbx1, a DiGeorge syndrome candidate gene, is regulated by sonic hedgehog during pharyngeal arch development. Dev Biol, 235, 62-73. GAUFO, G. O., THOMAS, K. R. & CAPECCHI, M. R. 2003. Hox3 genes coordinate mechanisms of genetic suppression and activation in the generation of branchial and somatic motoneurons. Development, 130, 5191-201. 192 GAUNT, S. J., SHARPE, P. T. & DUBOULE, D. 1988. Spatially restricted domains of homeo-gene transcripts in mouse embryos: relation to a segmented body plan. Development, 104, 169-179. GAVALAS, A., STUDER, M., LUMSDEN, A., RIJLI, F. M., KRUMLAUF, R. & CHAMBON, P. 1998. Hoxa1 and Hoxb1 synergize in patterning the hindbrain, cranial nerves and second pharyngeal arch. Development, 125, 1123-36. GENDRON-MAGUIRE, M., MALLO, M., ZHANG, M. & GRIDLEY, T. 1993. Hoxa-2 mutant mice exhibit homeotic transformation of skeletal elements derived from cranial neural crest. Cell, 75, 1317-31. GILMORE, A. P. 2005. Anoikis. Cell Death Differ, 12 Suppl 2, 1473-7. GOODRICH, E. S. 1906. Notes on the development, structure and origin of the median and paired fins of fish. Quarterly Journal of Microscopical Science, s2-50, 333-376. GORDON, J., BENNETT, A. R., BLACKBURN, C. C. & MANLEY, N. R. 2001. Gcm2 and Foxn1 mark early parathyroid- and thymus-specific domains in the developing third pharyngeal pouch. Mech Dev, 103, 141-3. GRAHAM, A. 2001. The development and evolution of the pharyngeal arches. J Anat, 199, 133-41. GRAHAM, A. & BEGBIE, J. 2000. Neurogenic placodes: a common front. Trends Neurosci, 23, 313-6. GRAHAM, A., FRANCIS-WEST, P., BRICKELL, P. & LUMSDEN, A. 1994. The signalling molecule BMP4 mediates apoptosis in the rhombencephalic neural crest. Nature, 372, 684-6. GRAHAM, A., HEYMAN, I. & LUMSDEN, A. 1993. Even-numbered rhombomeres control the apoptotic elimination of neural crest cells from odd-numbered rhombomeres in the chick hindbrain. Development, 119, 233-45. GRAHAM, A., OKABE, M. & QUINLAN, R. 2005. The role of the endoderm in the development and evolution of the pharyngeal arches. J Anat, 207, 47987. GRAHAM, A., PAPALOPULU, N. & KRUMLAUF, R. 1989. The murine and Drosophila homeobox gene complexes have common features of organization and expression. Cell, 57, 367-78. GRAHAM, A. & RICHARDSON, J. 2012. Developmental and evolutionary origins of the pharyngeal apparatus. Evodevo, 3, 24. GRAHAM, A. & SMITH, A. 2001. Patterning the pharyngeal arches. Bioessays, 23, 54-61. GRAMMATOPOULOS, G. A., BELL, E., TOOLE, L., LUMSDEN, A. & TUCKER, A. S. 2000. Homeotic transformation of branchial arch identity after Hoxa2 overexpression. Development, 127, 5355-65. 193 GREVELLEC, A. & TUCKER, A. S. 2010. The pharyngeal pouches and clefts: Development, evolution, structure and derivatives. Semin Cell Dev Biol, 21, 325-32. GUBBAY, J., COLLIGNON, J., KOOPMAN, P., CAPEL, B., ECONOMOU, A., MUNSTERBERG, A., VIVIAN, N., GOODFELLOW, P. & LOVELLBADGE, R. 1990. A gene mapping to the sex-determining region of the mouse Y chromosome is a member of a novel family of embryonically expressed genes. Nature, 346, 245-50. GUDERNATSCH, J. F. 1914. Feeding experiments on tadpoles. II. A further contribution to the knowledge of organs with internal secretion. American Journal of Anatomy, 15, 431-480. GUNTHER, T., CHEN, Z. F., KIM, J., PRIEMEL, M., RUEGER, J. M., AMLING, M., MOSELEY, J. M., MARTIN, T. J., ANDERSON, D. J. & KARSENTY, G. 2000. Genetic ablation of parathyroid glands reveals another source of parathyroid hormone. Nature, 406, 199-203. GURIS, D. L., DUESTER, G., PAPAIOANNOU, V. E. & IMAMOTO, A. 2006. Dose-dependent interaction of Tbx1 and Crkl and locally aberrant RA signaling in a model of del22q11 syndrome. Dev Cell, 10, 81-92. GUTHRIE, S. & LUMSDEN, A. 1991. Formation and regeneration of rhombomere boundaries in the developing chick hindbrain. Development, 112, 221-9. HAECKEL, E. 1910. The evolution of man, London, Watts & Co. HALL, B. K. 1997. Phylotypic stage or phantom: is there a highly conserved embryonic stage in vertebrates? Trends Ecol Evol, 12, 461-3. HAMBURGER, V. & HAMILTON, H. L. 1951. A series of normal stages in the development of the chick embryo. . Developmental Dynamics, 195, 231272. HAMILTON, H. L. & HINSCH, G. W. 1957. The fate of the second visceral pouch in the chick. Anat Rec, 129, 357-69. HAWORTH, K. E., HEALY, C., MORGAN, P. & SHARPE, P. T. 2004. Regionalisation of early head ectoderm is regulated by endoderm and prepatterns the orofacial epithelium. Development, 131, 4797-806. HILFER, S. R. & BROWN, J. W. 1984. The development of pharyngeal endocrine organs in mouse and chick embryos. Scan Electron Microsc, 2009-22. HOGAN, B. M., HUNTER, M. P., OATES, A. C., CROWHURST, M. O., HALL, N. E., HEATH, J. K., PRINCE, V. E. & LIESCHKE, G. J. 2004. Zebrafish gcm2 is required for gill filament budding from pharyngeal ectoderm. Dev Biol, 276, 508-22. HOLZSCHUH, J., WADA, N., WADA, C., SCHAFFER, A., JAVIDAN, Y., TALLAFUSS, A., BALLY-CUIF, L. & SCHILLING, T. F. 2005. Requirements for endoderm and BMP signaling in sensory neurogenesis in zebrafish. Development, 132, 3731-42. 194 HORIGOME, N., MYOJIN, M., UEKI, T., HIRANO, S., AIZAWA, S. & KURATANI, S. 1999. Development of cephalic neural crest cells in embryos of Lampetra japonica, with special reference to the evolution of the jaw. Dev Biol, 207, 287-308. HUDSON, C., CLEMENTS, D., FRIDAY, R. V., STOTT, D. & WOODLAND, H. R. 1997. Xsox17alpha and -beta mediate endoderm formation in Xenopus. Cell, 91, 397-405. HUNT, P., FERRETTI, P., KRUMLAUF, R. & THOROGOOD, P. 1995. Restoration of normal Hox code and branchial arch morphogenesis after extensive deletion of hindbrain neural crest. Dev Biol, 168, 584-97. HUNT, P., GULISANO, M., COOK, M., SHAM, M. H., FAIELLA, A., WILKINSON, D., BONCINELLI, E. & KRUMLAUF, R. 1991a. A distinct Hox code for the branchial region of the vertebrate head. Nature, 353, 861-4. HUNT, P. & KRUMLAUF, R. 1992. Hox codes and positional specification in vertebrate embryonic axes. Annu Rev Cell Biol, 8, 227-56. HUNT, P., WHITING, J., NONCHEV, S., SHAM, M. H., MARSHALL, H., GRAHAM, A., COOK, M., ALLEMANN, R., RIGBY, P. W., GULISANO, M. & ET AL. 1991b. The branchial Hox code and its implications for gene regulation, patterning of the nervous system and head evolution. Development, Suppl 2, 63-77. HUNTER, M. P. & PRINCE, V. E. 2002. Zebrafish hox paralogue group 2 genes function redundantly as selector genes to pattern the second pharyngeal arch. Dev Biol, 247, 367-89. ISHIHARA, T., IKEDA, K., SATO, S., YAJIMA, H. & KAWAKAMI, K. 2008. Differential expression of Eya1 and Eya2 during chick early embryonic development. Gene Expr Patterns, 8, 357-67. JANVIER, P. & ARSENAULT, M. 2007. The anatomy of Euphanerops longaevus Woodward, 1900, an anaspid-like jawless vertebrate from the Upper Devonian of Miguasha, Quebec, Canada. Geodiversitas, 29, 143216. JEROME, L. A. & PAPAIOANNOU, V. E. 2001. DiGeorge syndrome phenotype in mice mutant for the T-box gene, Tbx1. Nat Genet, 27, 286-91. KANAI-AZUMA, M., KANAI, Y., GAD, J. M., TAJIMA, Y., TAYA, C., KUROHMARU, M., SANAI, Y., YONEKAWA, H., YAZAKI, K., TAM, P. P. & HAYASHI, Y. 2002. Depletion of definitive gut endoderm in Sox17-null mutant mice. Development, 129, 2367-79. KANAI, Y., KANAI-AZUMA, M., NOCE, T., SAIDO, T. C., SHIROISHI, T., HAYASHI, Y. & YAZAKI, K. 1996. Identification of two Sox17 messenger RNA isoforms, with and without the high mobility group box region, and their differential expression in mouse spermatogenesis. J Cell Biol, 133, 667-81. 195 KESSEL, M. & GRUSS, P. 1990. Murine developmental control genes. Science, 249, 374-9. KEYNES, R. & LUMSDEN, A. 1990. Segmentation and the origin of regional diversity in the vertebrate central nervous system. Neuron, 4, 1-9. KIM, I., SAUNDERS, T. L. & MORRISON, S. J. 2007. Sox17 dependence distinguishes the transcriptional regulation of fetal from adult hematopoietic stem cells. Cell, 130, 470-83. KIM, J., JONES, B. W., ZOCK, C., CHEN, Z., WANG, H., GOODMAN, C. S. & ANDERSON, D. J. 1998. Isolation and characterization of mammalian homologs of the Drosophila gene glial cells missing. Proc Natl Acad Sci U S A, 95, 12364-9. KIMMEL, C. B., BALLARD, W. W., KIMMEL, S. R., ULLMANN, B. & SCHILLING, T. F. 1995. Stages of embryonic development of the zebrafish. Dev Dyn, 203, 253-310. KONTGES, G. & LUMSDEN, A. 1996. Rhombencephalic neural crest segmentation is preserved throughout craniofacial ontogeny. Development, 122, 3229-42. KOSTIC, D. & CAPECCHI, M. R. 1994. Targeted disruptions of the murine Hoxa-4 and Hoxa-6 genes result in homeotic transformations of components of the vertebral column. Mech Dev, 46, 231-47. KRUMLAUF, R. 1994. Hox genes in vertebrate development. Cell, 78, 191-201. KURATANI, S. 1997. Spatial distribution of postotic crest cells defines the head/trunk interface of the vertebrate body: embryological interpretation of peripheral nerve morphology and evolution of the vertebrate head. Anat Embryol (Berl), 195, 1-13. KURATANI, S., NOBUSADA, Y., HORIGOME, N. & SHIGETANI, Y. 2001. Embryology of the lamprey and evolution of the vertebrate jaw: insights from molecular and developmental perspectives. Philos Trans R Soc Lond B Biol Sci, 356, 1615-32. LACLEF, C., SOUIL, E., DEMIGNON, J. & MAIRE, P. 2003. Thymus, kidney and craniofacial abnormalities in Six 1 deficient mice. Mech Dev, 120, 669-79. LAEMLE, L. K., PUSZKARCZUK, M. & FEINBERG, R. N. 1999. Apoptosis in early ocular morphogenesis in the mouse. Brain Res Dev Brain Res, 112, 129-33. LARSEN, W. J. 1997. Human Embryology, New York, Churchill Livingstone Inc. LAWSON, A., SCHOENWOLF, G. C., ENGLAND, M. A., ADDAI, F. K. & AHIMA, R. S. 1999. Programmed cell death and the morphogenesis of the hindbrain roof plate in the chick embryo. Anat Embryol (Berl), 200, 509-19. 196 LEWIS, E. B. 1978. A gene complex controlling segmentation in Drosophila. Nature, 276, 565-70. LONG, J. A., YOUNG, G. C., HOLLAND, T., SENDEN, T. J. & FITZGERALD, E. M. 2006. An exceptional Devonian fish from Australia sheds light on tetrapod origins. Nature, 444, 199-202. LUMSDEN, A. & KEYNES, R. 1989. Segmental patterns of neuronal development in the chick hindbrain. Nature, 337, 424-8. LUMSDEN, A., SPRAWSON, N. & GRAHAM, A. 1991. Segmental origin and migration of neural crest cells in the hindbrain region of the chick embryo. Development, 113, 1281-91. LUMSDEN, A. G. S. 1988. Spatial organization of the epithelium and the role of neural crest cells in the initiation of the mammalian tooth germ. Development, 103, 155-169. MACONOCHIE, M., KRISHNAMURTHY, R., NONCHEV, S., MEIER, P., MANZANARES, M., MITCHELL, P. J. & KRUMLAUF, R. 1999. Regulation of Hoxa2 in cranial neural crest cells involves members of the AP-2 family. Development, 126, 1483-94. MACONOCHIE, M., NONCHEV, S., MORRISON, A. & KRUMLAUF, R. 1996. Paralogous Hox genes: function and regulation. Annu Rev Genet, 30, 529-56. MANLEY, N. R. & CAPECCHI, M. R. 1995. The role of Hoxa-3 in mouse thymus and thyroid development. Development, 121, 1989-2003. MANLEY, N. R. & CAPECCHI, M. R. 1997. Hox group 3 paralogous genes act synergistically in the formation of somitic and neural crest-derived structures. Dev Biol, 192, 274-88. MANLEY, N. R. & CAPECCHI, M. R. 1998. Hox group 3 paralogs regulate the development and migration of the thymus, thyroid, and parathyroid glands. Dev Biol, 195, 1-15. MARSHALL, H., MORRISON, A., STUDER, M., POPPERL, H. & KRUMLAUF, R. 1996. Retinoids and Hox genes. FASEB J, 10, 969-78. MARTIN, A. C., GELBART, M., FERNANDEZ-GONZALEZ, R., KASCHUBE, M. & WIESCHAUS, E. F. 2010. Integration of contractile forces during tissue invagination. J Cell Biol, 188, 735-49. MASSA, V., GREENE, N. D. & COPP, A. J. 2009a. Do cells become homeless during neural tube closure? Cell Cycle, 8, 2479-80. MASSA, V., SAVERY, D., YBOT-GONZALEZ, P., FERRARO, E., RONGVAUX, A., CECCONI, F., FLAVELL, R., GREENE, N. D. & COPP, A. J. 2009b. Apoptosis is not required for mammalian neural tube closure. Proc Natl Acad Sci U S A, 106, 8233-8. MATSUI, T., KANAI-AZUMA, M., HARA, K., MATOBA, S., HIRAMATSU, R., KAWAKAMI, H., KUROHMARU, M., KOOPMAN, P. & KANAI, 197 Y. 2006. Redundant roles of Sox17 and Sox18 in postnatal angiogenesis in mice. J Cell Sci, 119, 3513-26. MATT, N., GHYSELINCK, N. B., WENDLING, O., CHAMBON, P. & MARK, M. 2003. Retinoic acid-induced developmental defects are mediated by RARbeta/RXR heterodimers in the pharyngeal endoderm. Development, 130, 2083-93. MEREDITH, J. E., JR., FAZELI, B. & SCHWARTZ, M. A. 1993. The extracellular matrix as a cell survival factor. Mol Biol Cell, 4, 953-61. METZGER, R. J., KLEIN, O. D., MARTIN, G. R. & KRASNOW, M. A. 2008. The branching programme of mouse lung development. Nature, 453, 74550. MILLER, S. A., FAVALE, A. M. & KNOHL, S. J. 1993. Role for differential cell proliferation in perforation and rupture of chick pharyngeal closing plates. Anat Rec, 237, 408-14. MINOUX, M., ANTONARAKIS, G. S., KMITA, M., DUBOULE, D. & RIJLI, F. M. 2009. Rostral and caudal pharyngeal arches share a common neural crest ground pattern. Development, 136, 637-45. MULDER, G. B., MANLEY, N. & MAGGIO-PRICE, L. 1998. Retinoic acidinduced thymic abnormalities in the mouse are associated with altered pharyngeal morphology, thymocyte maturation defects, and altered expression of Hoxa3 and Pax1. Teratology, 58, 263-75. MULLER, T. S., EBENSPERGER, C., NEUBUSER, A., KOSEKI, H., BALLING, R., CHRIST, B. & 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, 403-17. NELSON, C. M. & GLEGHORN, J. P. 2012. Sculpting organs: mechanical regulation of tissue development. Annu Rev Biomed Eng, 14, 129-54. NICA, G., HERZOG, W., SONNTAG, C., NOWAK, M., SCHWARZ, H., ZAPATA, A. G. & HAMMERSCHMIDT, M. 2006. Eya1 is required for lineage-specific differentiation, but not for cell survival in the zebrafish adenohypophysis. Dev Biol, 292, 189-204. NIEDERREITHER, K., SUBBARAYAN, V., DOLLE, P. & CHAMBON, P. 1999. Embryonic retinoic acid synthesis is essential for early mouse postimplantation development. Nat Genet, 21, 444-8. NIEDERREITHER, K., VERMOT, J., LE ROUX, I., SCHUHBAUR, B., CHAMBON, P. & DOLLE, P. 2003. The regional pattern of retinoic acid synthesis by RALDH2 is essential for the development of posterior pharyngeal arches and the enteric nervous system. Development, 130, 2525-34. NODEN, D. M. 1983. The role of the neural crest in patterning of avian cranial skeletal, connective, and muscle tissues. Dev Biol, 96, 144-65. 198 O'NEILL, P., MAK, S. S., FRITZSCH, B., LADHER, R. K. & BAKER, C. V. 2012. The amniote paratympanic organ develops from a previously undiscovered sensory placode. Nat Commun, 3, 1041. OKABE, M. & GRAHAM, A. 2004. The origin of the parathyroid gland. Proc Natl Acad Sci U S A, 101, 17716-9. OULION, S., BORDAY-BIRRAUX, V., DEBIAIS-THIBAUD, M., MAZAN, S., LAURENTI, P. & CASANE, D. 2011. Evolution of repeated structures along the body axis of jawed vertebrates, insights from the Scyliorhinus canicula Hox code. Evol Dev, 13, 247-59. OULION, S., DEBIAIS-THIBAUD, M., D'AUBENTON-CARAFA, Y., THERMES, C., DA SILVA, C., BERNARD-SAMAIN, S., GAVORY, F., WINCKER, P., MAZAN, S. & CASANE, D. 2010. Evolution of Hox gene clusters in gnathostomes: insights from a survey of a shark (Scyliorhinus canicula) transcriptome. Mol Biol Evol, 27, 2829-38. PAPALOPULU, N., LOVELL-BADGE, R. & KRUMLAUF, R. 1991. The expression of murine Hox-2 genes is dependent on the differentiation pathway and displays a collinear sensitivity to retinoic acid in F9 cells and Xenopus embryos. Nucleic Acids Res, 19, 5497-506. PASQUALETTI, M., ORI, M., NARDI, I. & RIJLI, F. M. 2000. Ectopic Hoxa2 induction after neural crest migration results in homeosis of jaw elements in Xenopus. Development, 127, 5367-78. PETERS, H., NEUBUSER, A., KRATOCHWIL, K. & BALLING, R. 1998. Pax9-deficient mice lack pharyngeal pouch derivatives and teeth and exhibit craniofacial and limb abnormalities. Genes Dev, 12, 2735-47. PEVNY, L. H. & LOVELL-BADGE, R. 1997. Sox genes find their feet. Curr Opin Genet Dev, 7, 338-44. PIOTROWSKI, T., AHN, D. G., SCHILLING, T. F., NAIR, S., RUVINSKY, I., GEISLER, R., RAUCH, G. J., HAFFTER, P., ZON, L. I., ZHOU, Y., FOOTT, H., DAWID, I. B. & HO, R. K. 2003. The zebrafish van gogh mutation disrupts tbx1, which is involved in the DiGeorge deletion syndrome in humans. Development, 130, 5043-52. PIOTROWSKI, T. & NUSSLEIN-VOLHARD, C. 2000. The endoderm plays an important role in patterning the segmented pharyngeal region in zebrafish (Danio rerio). Dev Biol, 225, 339-56. PIOTROWSKI, T., SCHILLING, T. F., BRAND, M., JIANG, Y. J., HEISENBERG, C. P., BEUCHLE, D., GRANDEL, H., VAN EEDEN, F. J., FURUTANI-SEIKI, M., GRANATO, M., HAFFTER, P., HAMMERSCHMIDT, M., KANE, D. A., KELSH, R. N., MULLINS, M. C., ODENTHAL, J., WARGA, R. M. & NUSSLEIN-VOLHARD, C. 1996. Jaw and branchial arch mutants in zebrafish II: anterior arches and cartilage differentiation. Development, 123, 345-56. POELMANN, R. E., DUBOIS, S. V., HERMSEN, C., SMITS-VAN PROOIJE, A. E. & VERMEIJ-KEERS, C. 1985. Cell degeneration and mitosis in the 199 buccopharyngeal and branchial membranes in the mouse embryo. Anat Embryol (Berl), 171, 187-92. POPPERL, H., BIENZ, M., STUDER, M., CHAN, S. K., APARICIO, S., BRENNER, S., MANN, R. S. & KRUMLAUF, R. 1995. Segmental expression of Hoxb-1 is controlled by a highly conserved autoregulatory loop dependent upon exd/pbx. Cell, 81, 1031-42. PRINCE, V. & LUMSDEN, A. 1994. Hoxa-2 expression in normal and transposed rhombomeres: independent regulation in the neural tube and neural crest. Development, 120, 911-23. QI, B. Q., BEASLEY, S. W., WILLIAMS, A. K. & FIZELLE, F. 2000a. Apoptosis during regression of the tailgut and septation of the cloaca. J Pediatr Surg, 35, 1556-61. QI, B. Q., BEASLEY, S. W., WILLIAMS, A. K. & FRIZELLE, F. 2000b. Does the urorectal septum fuse with the cloacal membrane? J Urol, 164, 20702. QI, B. Q., WILLIAMS, A., BEASLEY, S. & FRIZELLE, F. 2000c. Clarification of the process of separation of the cloaca into rectum and urogenital sinus in the rat embryo. J Pediatr Surg, 35, 1810-6. QUINLAN, R., GALE, E., MADEN, M. & GRAHAM, A. 2002. Deficits in the posterior pharyngeal endoderm in the absence of retinoids. Dev Dyn, 225, 54-60. QUINLAN, R., MARTIN, P. & GRAHAM, A. 2004. The role of actin cables in directing the morphogenesis of the pharyngeal pouches. Development, 131, 593-9. RICHARDSON, J., SHONO, T., OKABE, M. & GRAHAM, A. 2012. The presence of an embryonic opercular flap in amniotes. Proc Biol Sci, 279, 224-9. RIJLI, F. M., MARK, M., LAKKARAJU, S., DIERICH, A., DOLLE, P. & CHAMBON, P. 1993. A homeotic transformation is generated in the rostral branchial region of the head by disruption of Hoxa-2, which acts as a selector gene. Cell, 75, 1333-49. RIZZOTI, K. & LOVELL-BADGE, R. 2007. SOX3 activity during pharyngeal segmentation is required for craniofacial morphogenesis. Development, 134, 3437-48. ROEHL, H. & NUSSLEIN-VOLHARD, C. 2001. Zebrafish pea3 and erm are general targets of FGF8 signaling. Curr Biol, 11, 503-7. RUHIN, B., CREUZET, S., VINCENT, C., BENOUAICHE, L., LE DOUARIN, N. M. & COULY, G. 2003. Patterning of the hyoid cartilage depends upon signals arising from the ventral foregut endoderm. Dev Dyn, 228, 239-46. SAKAMOTO, Y., HARA, K., KANAI-AZUMA, M., MATSUI, T., MIURA, Y., TSUNEKAWA, N., KUROHMARU, M., SAIJOH, Y., KOOPMAN, P. & KANAI, Y. 2007. Redundant roles of Sox17 and Sox18 in early 200 cardiovascular development of mouse embryos. Biochem Biophys Res Commun, 360, 539-44. SAP, J., MUNOZ, A., DAMM, K., GOLDBERG, Y., GHYSDAEL, J., LEUTZ, A., BEUG, H. & VENNSTROM, B. 1986. The c-erb-A protein is a highaffinity receptor for thyroid hormone. Nature, 324, 635-40. SAWYER, J. M., HARRELL, J. R., SHEMER, G., SULLIVAN-BROWN, J., ROH-JOHNSON, M. & GOLDSTEIN, B. 2010. Apical constriction: a cell shape change that can drive morphogenesis. Dev Biol, 341, 5-19. SCHEPERS, G. E., TEASDALE, R. D. & KOOPMAN, P. 2002. Twenty pairs of sox: extent, homology, and nomenclature of the mouse and human sox transcription factor gene families. Dev Cell, 3, 167-70. SCHILLING, T. F. & KIMMEL, C. B. 1994. Segment and cell type lineage restrictions during pharyngeal arch development in the zebrafish embryo. Development, 120, 483-94. SCHILLING, T. F., PIOTROWSKI, T., GRANDEL, H., BRAND, M., HEISENBERG, C. P., JIANG, Y. J., BEUCHLE, D., HAMMERSCHMIDT, M., KANE, D. A., MULLINS, M. C., VAN EEDEN, F. J., KELSH, R. N., FURUTANI-SEIKI, M., GRANATO, M., HAFFTER, P., ODENTHAL, J., WARGA, R. M., TROWE, T. & NUSSLEIN-VOLHARD, C. 1996. Jaw and branchial arch mutants in zebrafish I: branchial arches. Development, 123, 329-44. SHIGETANI, Y., SUGAHARA, F., KAWAKAMI, Y., MURAKAMI, Y., HIRANO, S. & KURATANI, S. 2002. Heterotopic shift of epithelialmesenchymal interactions in vertebrate jaw evolution. Science, 296, 13169. SILVER, J. & HUGHES, A. F. 1973. The role of cell death during morphogenesis of the mammalian eye. J Morphol, 140, 159-70. SINCLAIR, A. H., BERTA, P., PALMER, M. S., HAWKINS, J. R., GRIFFITHS, B. L., SMITH, M. J., FOSTER, J. W., FRISCHAUF, A. M., LOVELL-BADGE, R. & GOODFELLOW, P. N. 1990. A gene from the human sex-determining region encodes a protein with homology to a conserved DNA-binding motif. Nature, 346, 240-4. SLACK, J. M., HOLLAND, P. W. & GRAHAM, C. F. 1993. The zootype and the phylotypic stage. Nature, 361, 490-2. SOUKUP, V., HORACEK, I. & CERNY, R. 2013. Development and evolution of the vertebrate primary mouth. J Anat, 222, 79-99. STELLWAG, E. J. 1999. Hox gene duplication in fish. Semin Cell Dev Biol, 10, 531-40. SU, D., ELLIS, S., NAPIER, A., LEE, K. & MANLEY, N. R. 2001. Hoxa3 and pax1 regulate epithelial cell death and proliferation during thymus and parathyroid organogenesis. Dev Biol, 236, 316-29. TAKIO, Y., KURAKU, S., MURAKAMI, Y., PASQUALETTI, M., RIJLI, F. M., NARITA, Y., KURATANI, S. & KUSAKABE, R. 2007. Hox gene 201 expression patterns in Lethenteron japonicum embryos--insights into the evolution of the vertebrate Hox code. Dev Biol, 308, 606-20. TAKIO, Y., PASQUALETTI, M., KURAKU, S., HIRANO, S., RIJLI, F. M. & KURATANI, S. 2004. Evolutionary biology: lamprey Hox genes and the evolution of jaws. Nature, 429, 1 p following 262. TATA, J. R. 1968. Early metamorphic competence of Xenopus larvae. Dev Biol, 18, 415-40. TRAINOR, P. & KRUMLAUF, R. 2000. Plasticity in mouse neural crest cells reveals a new patterning role for cranial mesoderm. Nat Cell Biol, 2, 96102. TUCKER, A. S. & SHARPE, P. T. 1999. Molecular genetics of tooth morphogenesis and patterning: the right shape in the right place. J Dent Res, 78, 826-34. TUMPEL, S., CAMBRONERO, F., FERRETTI, E., BLASI, F., WIEDEMANN, L. M. & KRUMLAUF, R. 2007. Expression of Hoxa2 in rhombomere 4 is regulated by a conserved cross-regulatory mechanism dependent upon Hoxb1. Dev Biol, 302, 646-60. VEITCH, E. 2000. Pharyngeal arch development. PhD, King's College London. VEITCH, E., BEGBIE, J., SCHILLING, T. F., SMITH, M. M. & GRAHAM, A. 1999. Pharyngeal arch patterning in the absence of neural crest. Curr Biol, 9, 1481-4. VITELLI, F., TADDEI, I., MORISHIMA, M., MEYERS, E. N., LINDSAY, E. A. & BALDINI, A. 2002. A genetic link between Tbx1 and fibroblast growth factor signaling. Development, 129, 4605-11. WALL, N. A. & HOGAN, B. L. 1995. Expression of bone morphogenetic protein-4 (BMP-4), bone morphogenetic protein-7 (BMP-7), fibroblast growth factor-8 (FGF-8) and sonic hedgehog (SHH) during branchial arch development in the chick. Mech Dev, 53, 383-92. WALLIN, J., EIBEL, H., NEUBUSER, A., WILTING, J., KOSEKI, H. & BALLING, R. 1996. Pax1 is expressed during development of the thymus epithelium and is required for normal T-cell maturation. Development, 122, 23-30. WATANABE, K., SASAKI, F. & TAKAHAMA, H. 1984. The ultrastructure of oral (buccopharyngeal) membrane formation and rupture in the anuran embryo. Anat Rec, 210, 513-24. WATERMAN, R. E. 1977. Ultrastructure of oral (buccopharyngeal) membrane formation and rupture in the hamster embryo. Dev Biol, 58, 219-29. WATERMAN, R. E. 1985. Formation and perforation of closing plates in the chick embryo. Anat Rec, 211, 450-7. WATERMAN, R. E. & SCHOENWOLF, G. C. 1980. The ultrastructure of oral (buccopharyngeal) membrane formation and rupture in the chick embryo. Anat Rec, 197, 441-70. 202 WEGNER, M. 1999. From head to toes: the multiple facets of Sox proteins. Nucleic Acids Res, 27, 1409-20. WEIL, M., JACOBSON, M. D. & RAFF, M. C. 1997. Is programmed cell death required for neural tube closure? Curr Biol, 7, 281-4. WEINBERGER, C., THOMPSON, C. C., ONG, E. S., LEBO, R., GRUOL, D. J. & EVANS, R. M. 1986. The c-erb-A gene encodes a thyroid hormone receptor. Nature, 324, 641-6. WENDLING, O., DENNEFELD, C., CHAMBON, P. & MARK, M. 2000. Retinoid signaling is essential for patterning the endoderm of the third and fourth pharyngeal arches. Development, 127, 1553-62. WILKINSON, D. G., BHATT, S., COOK, M., BONCINELLI, E. & KRUMLAUF, R. 1989. Segmental expression of Hox-2 homoeoboxcontaining genes in the developing mouse hindbrain. Nature, 341, 405-9. WILSON, J. M. & LAURENT, P. 2002. Fish gill morphology: inside out. J Exp Zool, 293, 192-213. WONG, E. Y., WANG, X. A., MAK, S. S., SAE-PANG, J. J., LING, K. W., FRITZSCH, B. & SHAM, M. H. 2011. Hoxb3 negatively regulates Hoxb1 expression in mouse hindbrain patterning. Dev Biol, 352, 382-92. XU, P. X., ZHENG, W., LACLEF, C., MAIRE, P., MAAS, R. L., PETERS, H. & XU, X. 2002. Eya1 is required for the morphogenesis of mammalian thymus, parathyroid and thyroid. Development, 129, 3033-44. YAGI, H., FURUTANI, Y., HAMADA, H., SASAKI, T., ASAKAWA, S., MINOSHIMA, S., ICHIDA, F., JOO, K., KIMURA, M., IMAMURA, S., KAMATANI, N., MOMMA, K., TAKAO, A., NAKAZAWA, M., SHIMIZU, N. & MATSUOKA, R. 2003. Role of TBX1 in human del22q11.2 syndrome. Lancet, 362, 1366-73. YALCIN, H. C., SHEKHAR, A., RANE, A. A. & BUTCHER, J. T. 2010. An exovo chicken embryo culture system suitable for imaging and microsurgery applications. J Vis Exp. YAOITA, Y., SHI, Y. & BROWN, D. 1990. Xenopus laevis alpha and beta thyroid hormone receptors. Proc Natl Acad Sci U S A, 87, 8684. ZOU, D., SILVIUS, D., DAVENPORT, J., GRIFONE, R., MAIRE, P. & XU, P. X. 2006. Patterning of the third pharyngeal pouch into thymus/parathyroid by Six and Eya1. Dev Biol, 293, 499-512. 203