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Development, morphogenesis and evolution of pharyngeal segmentation in
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Shone, Victoria Louise
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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.
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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).
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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.
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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
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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).
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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.
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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
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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
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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,
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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
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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
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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
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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
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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).
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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.
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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.
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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
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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
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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.
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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.
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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
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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
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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
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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
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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).
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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.
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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.
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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.
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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.
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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
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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
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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
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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.
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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).
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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.
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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.
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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).
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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
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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
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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.
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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.
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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
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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
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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.
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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
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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.
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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,
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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
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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.
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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).
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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
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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.
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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
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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
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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.
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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)
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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
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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
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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
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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
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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
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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.
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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.
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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
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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