Making Headway: The Roles of Hox Genes and Neural Crest Cells in Craniofacial Development

Abstract

Craniofacial development is an extraordinarily complex process requiring the orchestrated integration of multiple specialized tissues such as the surface ectoderm, neural crest, mesoderm, and pharyngeal endoderm in order to generate the central and peripheral nervous systems, axial skeleton, musculature, and connective tissues of the head and face. How do the characteristic facial structures develop in the appropriate locations with their correct shapes and sizes, given the widely divergent patterns of cell movements that occur during head development? The patterning information could depend upon localized interactions between the epithelial and mesenchymal tissues or alternatively, the developmental program for the characteristic facial structures could be intrinsic to each individual tissue precursor. Understanding the mechanisms that control vertebrate head development is an important issue since craniofacial anomalies constitute nearly one third of all human congenital defects. This review discusses recent advances in our understanding of neural crest cell patterning and the dynamic nature of the tissue interactions that are required for normal craniofacial development.

Full text

Mini-Review
TheScientificWorldJOURNAL (2003) 3, 240–264
ISSN 1537-744X; DOI 10.1100/tsw.2003.11
Making Headway: The Roles of Hox Genes and
Neural Crest Cells in Craniofacial Development
Paul A Trainor
Stowers Institute for Medical Research, 1000 East 50
th
Street, Kansas City, MO 64110
E-mail: pat@stowers-institute.org; www.stowers-institute.org
Received April 11, 2002; Revised June 11, 2002; Accepted June 19, 2002; Published April 14, 2003
Craniofacial development is an extraordinarily complex process requiring the
orchestrated integration of multiple specialized tissues such as the surface
ectoderm, neural crest, mesoderm, and pharyngeal endoderm in order to generate
the central and peripheral nervous systems, axial skeleton, musculature, and
connective tissues of the head and face. How do the characteristic facial
structures develop in the appropriate locations with their correct shapes and
sizes, given the widely divergent patterns of cell movements that occur during
head development? The patterning information could depend upon localized
interactions between the epithelial and mesenchymal tissues or alternatively, the
developmental program for the characteristic facial structures could be intrinsic
to each individual tissue precursor. Understanding the mechanisms that control
vertebrate head development is an important issue since craniofacial anomalies
constitute nearly one third of all human congenital defects. This review discusses
recent advances in our understanding of neural crest cell patterning and the
dynamic nature of the tissue interactions that are required for normal craniofacial
development.
KEY WORDS: Hox genes, neural crest cells, craniofacial development, mouse, chick,
embryo, branchial arch, pharyngeal arch, plasticity, neural tube, rhombomere,
segmentation, mesoderm, ectoderm, endoderm, evolution
DOMAINS: genetics (mouse), embryology, neuroscience, developmental biology,
evolutionary genetics
ORGANISATION OF THE HINDBRAIN AND ITS INFLUENCE ON
CRANIOFACIAL DEVELOPMENT
The vertebrate hindbrain is one key source of patterning information and it exerts a profound
influence on craniofacial development. During early vertebrate embryo development, the
*Corresponding author.
©2003 with author.
240

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
hindbrain becomes transiently subdivided into seven contiguous cell lineage restricted
compartments called rhombomeres (r) (Fig. 1)[1]. Each rhombomere adopts a distinct set of
molecular and cellular properties including restrictions in cell mixing and gives rise to unique
regions of the mature adult brain[2,3,4,5,6,7,8,9]. The segmental organization of the hindbrain
presages the establishment of an anatomical and functional registration between individual
rhombomeres, cranial ganglia, branchiomotor nerves, and the migration pathways of cranial
neural crest cells into the branchial arches[10,11,12].
In chick embryos, the formation and disposition of motor neurons conforms to a two-
segment periodicity. The trigeminal (Vth), facio-acoustic (VIIth), and glossopharyngeal (IXth)
motor nerves innervate the first three branchial arches, respectively. Although each nerve is
derived from neurons born in multiple rhombomeres, the axons leave the hindbrain through exit
points contained only within the even-numbered rhombomeres (Fig. 1)[10,11]. Similar to its
influence on cranial nerve patterning, the rhombomeric organization of the hindbrain also
influences the pathways of cranial neural crest cell migration. The hindbrain gives rise to the
majority of cranial neural crest cells which migrate ventrolaterally in three discrete segmental
streams adjacent to the even-numbered rhombomeres. The streams of cranial neural crest cells
follow a subectodermal route over the surface of the cranial mesoderm and populate the first,
second, and third branchial arches, respectively, in keeping with their craniocaudal origins (Fig.
1)[13,14,15,16]. Although almost the entire cranial neural tube generates neural crest cells, it
appears that substantially less neural crest cells delaminate from the odd rhombomeres compared
with the even-numbered rhombomeres. In chick embryos, this phenomenon is believed to be
associated with odd rhombomere specific apoptosis of neural crest precursors[17,18,19]. Rather
than delaminating and migrating laterally like the rest of the hindbrain neural crest, lineage
tracing and time-lapse analyses in numerous vertebrates, show that r3- and r5-derived neural crest
cells migrate both rostrally and caudally joining the even-numbered streams as they contribute to
the proximal most regions of the branchial arches[12,14,15,16,20,21,22,23]. Gene mutations—
which disrupt the segmental patterning of the hindbrain—result in fusions of the cranial ganglia,
branchiomotor defects, and abnormal patterns of cranial neural crest cell
migration[24,25,26,27,28,29,30,31,32,33]. Therefore, the segmental organization of the hindbrain
is essential for establishing the foundations of head development and is a conserved strategy used
by vertebrates to ensure proper craniofacial morphogenesis.
MECHANISMS FOR SEGMENTATION OF THE HINDBRAIN
The establishment of segment identity and the maintenance of organized patterns of gene
expression during hindbrain development requires the restriction of cell intermingling between
adjacent segments. Cells with similar adhesive properties such as those from odd or even
rhombomeres display a preferential association[34,35]. Consequently, it was hypothesized that
segregating the distinct cell populations was facilitated by a hierarchy of cell adhesion
interactions[36]. The appearance of restricted domains of gene expression in the hindbrain
coincides with rhombomere partitioning and the adoption of neural character (Fig. 1). Numerous
genes including transcription factors, signalling molecules, membrane and nuclear receptors are
all dynamically expressed in segmentally restricted patterns during hindbrain development
(reviewed in [37,38,39]). Some genes are expressed in single rhombomeres, a few are confined to
pairs of rhombomeres, while others are found only at rhombomere boundaries. Regulatory and
mutational analyses in mice and other vertebrate species have helped to uncover the molecular
and cellular mechanisms involved in subdividing the hindbrain into rhombomeres.
Restrictions in cell mixing are crucial for establishing the unique segmental identities of each
rhombomere and the Eph family of receptor tyrosine kinases and their ligands, the ephrins, play
key roles in this process[40]. The Eph receptors and their ephrin ligands are expressed in
241

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
mutually exclusive domains in the hindbrain such that each receptor and its associated ligand is
expressed in an alternate, nonoverlapping pattern in adjacent rhombomeres (Fig. 1)[40,41,42,43].
For example, during the 8.0–10.5 days postcoitum (dpc) period of mouse embryonic
development, EphB receptors are expressed in odd-numbered rhombomeres and their
corresponding ephrinB ligands are expressed in the adjacent even-numbered rhombomeres. In
addition to the EphB receptors, the receptors EphA2 and EphA4 are also segmentally expressed in
hindbrain rhombomeres (Fig. 1). The mosaic activation of Eph receptors or their ephrin ligands in
the hindbrain leads to cell sorting at the boundaries of odd and even rhombomeres, respectively.
This suggests that bidirectional signalling at rhombomere boundaries restricts cell
intermingling[44,45]. Therefore bipartite Eph/ephrin contact-mediated repulsion maintains inter-
rhombomeric boundaries which are essential for establishing the hindbrain compartments as units
of cell lineage restriction.
SPECIFYING SEGMENTAL IDENTITY IN THE HINDBRAIN
Genetic analyses in several vertebrate species have shown that Hox genes are essential for
specifying segmental identity in the developing hindbrain. The Hox gene family comprises a set
of developmentally regulated transcription factors that are characterized by a conserved DNA
binding motif called the homeobox. Hox genes are organized into a single chromosomal cluster in
invertebrates, in contrast to higher vertebrates such as the mouse which have 39 Hox genes,
organized into four distinct chromosomal clusters (Hoxa-Hoxd) located on different
chromosomes[46]. This arrangement arose during evolution from a single ancestral homeobox as
a consequence of duplication and divergence[47]. The most striking feature of the organization of
Hox gene family members is the spatial and temporal colinearity, which confers positional
information along the body axis[46,48,49,50]. Genes located nearer to the 3’ end of the cluster
are expressed earlier and more anteriorly during development than those located nearer the 5’
end, such that Hox genes exhibit nested domains of expression along the anterior-posterior (A-P)
axis of the neural tube. Only the 3’ members of each complex (paralogue groups 1-4) are
expressed in the vertebrate hindbrain and these Hox genes exhibit dynamic patterns of expression
during hindbrain development (Fig. 1)[51].
Hoxa1 and Hoxb1 are initially expressed in the neural tube up to the presumptive r3/4
boundaries, but as Hoxa1 expression regresses, Hoxb1 expression becomes confined to r4 by
8.5dpc during mouse embryo development[52,53,54]. Hoxa2 expression is initiated in the neural
epithelium at 7.5dpc and by 8.5dpc it is uniformly expressed up to the r1/r2 boundary[51].
Shortly thereafter Hoxa2 becomes expressed at significantly higher levels in r3 and r5[55]. In
contrast, Hoxb2 is expressed up to the r2/r3 boundary around the same time, but its expression
levels are subsequently upregulated in r3, r4, and r5[56,57]. Group 3 (Hoxa3, Hoxb3, Hoxd3)
genes are expressed anteriorly up to the r4/r5 boundary, with Hoxa3 exhibiting upregulation in r5
and r6 and Hoxb3 being similarly upregulated in r5[58,59]. In contrast, Hoxd3 is expressed at
lower levels in r5 than more posterior regions. Group 4 genes (Hoxa4, Hoxb4, Hoxd4, and
Hoxc4) are generally expressed up to the r6/r7 boundary by 9.5dpc, however their precise anterior
limits vary slightly (Fig. 1)[60,61].
242

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
FIGURE 1. Segmentation of the hindbrain, motor nerves, and pathways of neural crest cell (ncc) migration. The vertebrate hindbrain
becomes subdivided into 7 transient rhombomeres (r) which presages the segmental organization of the branchiomotor nerves, III
(trigeminal), V (facial), and VII (glossopharyngeal) and their innervation of the first (ba1), second (ba2), and third (ba3) branchial
arches, respectively. Although the cell bodies (small red circles) that make up each branchiomotor nerve are derived from multiple
rhombomeres, their axons project and leave the neural tube only from exit points contained within the even numbered rhombomeres
(large red circles). Neural crest cells derived from the hindbrain (large green arrows) migrate ventrolaterally in three segmental
streams adjacent to the even-numbered rhombomeres and into the branchial arches. Neural crest cells derived from the odd-numbered
rhombomeres do not migrate laterally, rather they migrate anteriorly or posteriorly to join the even-numbered neural crest streams.
The hindbrain segmentation process which establishes each rhombomere as a unit of cell lineage and the subsequent imposition of cell
fate identity corresponds with the activation of expression, of genes from multiple families, such as Hox, Eph/ephrin, RAR and
CRABP families. The patterns of gene expression are extremely dynamic and only the primary pattern for each gene between 8.5–
10.5dpc is shown here. The only exception is Hoxa1 which is expressed up to the presumptive r3/4 boundary at 7.5dpc but is
subsequently switched off before rhombomere formation. OV, otic vesicle.
GENETIC CONTROL OF HOX GENE EXPRESSION AND HINDBRAIN
PATTERNING
Hox genes are an integral part of the process specifying regional variation in the developing
hindbrain, however little is known of the cascade of events that regulate their expression or the
process of segmentation itself. In general, Hox gene expression patterns are generated in two
distinct phases: establishment followed by maintenance. The mechanism by which Hox genes
become expressed at higher relative levels in specific rhombomeres is independent from the
process that generates the more generalized expression patterns along the length of the neural
tube. Positional values along the A-P axis appear to be conferred on the rhombomeres by
differential Hox gene expression however it is not entirely clear how appropriate levels of
expression are attained within individual rhombomeres. The observation that other genes were
243

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
expressed in the hindbrain in segmentally restricted patterns overlapping with the patterns of Hox
gene expression implied that these genes were involved in regulating Hox gene expression and
ultimately controlling hindbrain development. Currently, there are three upstream candidate
regulators of Hox gene expression during hindbrain development and they are the transcription
factors Krox20 and kreisler and the vitamin A derivative, retinoic acid.
KROX20 REGULATION OF HOX GENES
Krox20 encodes a protein with three C
2
H
2
-type zinc fingers and was initially identified as an
immediate early response gene in serum stimulated fibroblasts[62]. During early embryonic
development, Krox20 is expressed in r3 by the 8.0dpc stage, followed by r5, which occurs prior to
the appearance of lineage-restricted compartments in the hindbrain (Fig. 1)[2]. Krox20 acts as a
transcription factor by binding to a specific DNA sequence and directly regulates Hoxa2, Hoxb2,
Hoxb3, and EphA4 through cis-acting sequences in the 5’ flanking region of these genes[63,64].
Krox20 therefore is an essential component of the upstream regulatory cascade governing
hindbrain segmentation. The overlapping expression patterns of Krox20 and EphA4 expression
suggests that transcriptional control and cell-cell signalling could be coupled together.
KREISLER REGULATION OF HOX GENES
The kreisler mouse was identified due to its circling behavior, a phenotype, which is usually
associated with inner ear defects[65]. Kreisler expression is initiated at 8.5dpc in the prospective
r5 territory and is later expressed in r5 and r6 after which it is quickly downregulated in both
rhombomeres (Fig. 1)[66]. The gene responsible for the kreisler mutation has been positionally
cloned, and is a member of the Maf oncogenic family of b-zip (basic domain-leucine zipper)
transcription factors[66]. The analysis of gene expression in kreisler mutant embryos strongly
suggested it might play a direct role in the transcriptional regulation of genes important for
hindbrain and inner ear development, particularly the Hoxa3 and Hoxb3 genes that are
upregulated in r5 and r6. Analyses of the regulatory regions of Hoxa3 and Hoxb3 in mouse and
chick embryos have shown that both these genes are under the direct control of the product of the
kreisler gene[32].
RETINOID REGULATION OF HOX GENE EXPRESSION
Retinoids constitute a group of vitamin A–derived signalling molecules that are involved in
specifying the CNS[67,68]. Exogenous vitamin A metabolites such as retinoic acid (RA), when
applied to embryos during the early stages of development, cause severe craniofacial
malformations, particularly within the hindbrain and branchial arch region[67,69,70,71,72].
Cellular retinoid-binding proteins (CRBPI and II, CRABP I and II) are thought to control regional
RA concentration and hence facilitate its function[73]. CRBP expression is confined primarily to
the floor plate, where it binds retinol taken up from the blood and CRABPI is expressed in the
mouse hindbrain in r4-6 and at lower levels in r2 at 9.5dpc[74,75]. CRABPII is also expressed at
high levels in the hindbrain and its expression extends posteriorly along the length of the neural
tube (Fig. 1)[76,77]. CRABPs therefore may be responsible for sequestering RA and thus limiting
the amount of RA available to the nuclear RA receptors (RARs and RXRs) that recognize
particular sequence motifs within target genes called r
etinoic acid response elements
(RAREs)[78,79]. There are three genes in each of the RAR and RXR families (a, b, and g). RARa
is expressed up to the r3/4 border while RARb is expressed in the neural tube to an anterior limit
244

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
coinciding with the r6/7 border[80,81]. RXRa and RXRb are expressed fairly uniformly
throughout the entire hindbrain neuroepithelium (Fig. 1)[82,83]. Not all the family members
however are expressed in the hindbrain
RA is the strongest candidate as an overall mediator of nested Hox gene expression. Excess
RA causes both an anterior shift in Hox gene expression and an anterior to posterior
transformation of regional fate[50,67,71,72,84,85,86,87,88,89,90,91,92]. Conversely, the
suppression of RA signalling by the expression of dominant-negative retinoid receptors, results in
anteriorization[93,94,95,96]. Hox genes contain the molecular machinery for responding directly
to retinoid signalling[97,98,99,100,101,102,103]. The group 1 paralogs Hoxa1 and Hoxb1 contain
3’ cis-regulatory RARE as do the group 4 paralogs Hoxa4, Hoxb4 and Hoxd4. Hoxa1 and Hoxb1
are among the first Hox genes to be activated during development and their expression is initiated
during gastrulation in response to RA, first in the newly formed mesoderm, and then in the
overlying ectoderm up to a sharp anterior limit at the presumptive rhombomere 3/4 hindbrain
border. As Hoxa1 expression subsides, retinoids make a second input into Hoxb1 regulation by
refining its domain of expression to a single rhombomere segment (r4). This second input
involves a negative pathway that is employed to abolish expression in the neighboring odd
rhombomeres r3 and r5[99,100]. The site for repression is located within the 5’ flanking region of
the Hoxb1 gene and includes another RARE, point mutations in which permit expression to
spread into the neighboring rhombomeres. The application of exogenous retinoids at 7.5dpc
results in the ectopic activation of Hoxa1, Hoxb1, and Hoxa2 in more anterior tissues which is
consistent with RAs perceived role as having a posteriorizing effect on the CNS. The group 4
paralogs Hoxa4, Hoxb4, and Hoxd4 genes are also capable of responding to RA but only when
administered between 8.5–9.5dpc[61,92,104]. Therefore retinoids (or more specifically RA acting
via its receptor molecules [RARs and RXRs]) are crucial in the establishment phase of Hox gene
expression and the specification of segmental identity in the hindbrain.
HOX GENE AUTO AND CROSS-REGULATION
A unique property of the group 1 paralogs is displayed by the Hoxb1 gene which after its
establishment phase, maintains high levels of expression in r4[52,54]. The continued expression
or maintenance phase of Hoxb1 in r4 does not involve RA but is achieved through a conserved
autoregulatory loop involving Hoxb1 together with an unspecified Pbx family protein[105].
Despite the similarities in expression and regulation between various Hox gene paralogs, there is
frequently variation in their relative levels within specific segments. For example, of the group 2
homologs simultaneously expressed in r4, Hoxb2 is upregulated, but in contrast, Hoxa2 is not.
This type of differential expression suggests that in even-numbered rhombomeres these two genes
have distinct modes of regulation. Consequently it has been shown in vitro that Hoxb1 is able to
bind to Hoxb2 in a cross-regulatory interaction[57]. A great deal of our understanding of Hox
cross-regulation has come from analyses of the cis-regulatory regions of the group 4 paralogs,
Hoxb4 and Hoxd4, which have domains of expression that partially overlap and map to the r6/7
junction[106]. Similar to Hoxb1, Hoxb4, and Hoxd4 are also capable of auto-regulation. These
studies highlight the importance of auto-regulatory and cross-regulatory mechanisms for the
functional maintenance of Hox gene expression and specification of rhombomere identity during
vertebrate hindbrain development.
245

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
HOX GENE SPECIFICATION OF SEGMENTAL IDENTITY AND MUTATIONAL
ANALYSES
By analogy to their Drosophila counterparts, it was predicted that Hox genes would play essential
roles in regulating the identity of individual hindbrain segments[4,51,52] and this has now been
confirmed through null mutation and ectopic expression analyses for most of the 3’ members of
the Hox gene family. Hoxa1 null mutants die at birth from anoxia and exhibit marked defects in
the inner ear and in specific cranial nerve components[24]. The embryonic phenotype is
characterized by a reduction in the size of r4 and the partial deletion of r5 as revealed by the
diminished expression of Hoxb1 in r4, and of Krox20 and Fgf3 in r5, and this may account for the
absence of facial nerve and abducens nerve motor neurons. A second independently produced
Hoxa1 null mutant exhibits a slightly different phenotype, in which there is a complete absence of
rhombomere segmentation, including the first three rhombomeres where Hoxa1 is never
expressed[25]. Nevertheless, the null mutant phenotypes indicate that the Hoxa1 gene product is
necessary for the proper segmentation of r4-6. In addition the activity of Hoxa1 appears to be
critical for the specification of rhombomeric identity because ectopic expression of this gene in r2
results in the adoption of gene expression typical of r4[107,108]. Hoxa1 is therefore critical to
both hindbrain segmentation and specification.
Hoxb1 mutants exhibit changes in segmental identity; however, in contrast to the Hoxa1
mutants they show no segmentation defects[30,31]. Molecular analyses indicate that the
patterning of r4 is initiated properly but not maintained in Hoxb1 mutant embryos. Rhombomere
4 specific markers such as Wnt8c and CRABP1 fail to be upregulated and genetic markers of r2
are ectopically expressed in r4 suggesting that the identity of r4 has been transformed[31].
Consequently, the facial branchiomotor neurons (FBM) and contralateral vestibular acoustic
efferent (CVA) neurons, which are specific to r4 are incorrectly specified. Both sets of neurons
fail to migrate into the correct position leading to a loss of the facial motor nerve. These results
demonstrate that as part of its role is in maintaining rhombomere identity, Hoxb1 is involved in
controlling migratory properties of motor neurons in the hindbrain. Ectopic expression of Hoxb1
in r2 of chick embryos transforms the character of this segment into that of rhombomere 4[109].
These gain of function experiments suggest that Hoxb1 is involved in specifying individual
segmental identity.
The phenotypes of individual Hoxa1 and Hoxb1 loss of function (null) mutations suggest
that these genes play distinct roles in hindbrain development. However, Hoxa1/Hoxb double
mutants exhibit a surprising range of phenotypes that are absent from the individual mutants,
indicating that establishment of r4 identity and patterning of the VII-XI cranial nerves requires
extensive synergy between Hoxa1 and Hoxb1[110,111]. In the absence of both genes, a territory
appears in the region of r4, however EphA2 (which is one of the earliest markers of r4) fails to be
activated. Not only does EphA2 lie downstream of Hoxa1 and Hoxb1 in the genetic cascade that
regulates hindbrain segmentation and specification, but it suggests that in the double mutants
there is a failure to initiate rather than maintain the specification of r4 identity. The double
mutants also revealed that Hoxa1 and Hoxb1 work synergistically to initiate the r4 restricted
expression domain of Hoxb1. Hoxa1 seems to play a role during the establishment phase of Hox
gene expression by activating the r4 enhancer of Hoxb1. This is achieved through 3’ RARE
pararegulatory interactions. Hoxa1 however, is unable to participate in the long-term maintenance
of Hoxb1 expression because it is not expressed at these later stages. Therefore in concert with
RA, pararegulatory interactions between Hoxa1 and Hoxb1 initiates Hoxb1 expression in r4, after
which time, the Hoxb1 autoregulatory mechanism kicks in to ensure that high Hoxb1 expression
levels are maintained. Hoxb1 also functions in a direct cross-regulatory manner to upregulate
Hoxb2 expression in r4. Therefore Hoxb1 utilizes auto-, para-, and cross-regulatory mechanisms
as part of its role in maintaining r4 identity and regulating facial motor neuron patterning.
246

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
Hoxa2 is the only Hox gene to be expressed in r2 and is the most anteriorly expressed of all
the Hox genes[112]. Null mutations in Hoxa2 result in homeotic transformations of the second
arch neural crest derived elements into first arch derivatives and subsequently perinatal
lethality[113]. The hindbrain is also affected to the extent that the segmental identities of r2 and
r3 are altered. EphA4 expression is selectively abolished in r2, indicating that Hoxa2 is required
for the maintenance of EphA4 expression in r2. Consequently, the alar territories of r2 and r3 are
reduced and there is a concomitant expansion of r1. In addition, the trigeminal motor axons
derived from r2 and r3 migrate caudally and exit the hindbrain from the r4 facial nerve exit point
rather than their normal exit point in r2. Since Hoxa2 is the only Hox gene expressed in r2, these
results suggest that the identity of r2 has been transformed and that it now resembles r4. Hoxa2
therefore not only acts as a selector gene for second arch mesenchymal neural crest cells but also
plays a fundamental role in rostral hindbrain patterning by establishing the identity of r2 and by
influencing the migration of r2/3 derived motor axons.
In contrast to Hoxa2 mutants, the loss of Hoxb2 locus does not result in abnormalities to the
segmental anterio-posterior organization of the developing hindbrain[114]. Rhombomeres 3 and 5
develop normally, despite the absence of Hoxb2 upregulation in these rhombomeres. Phenotypic
changes affecting dorsoventral (D-V) patterning however were observed in the hindbrain such
that r4 specific neural precursors now closely resembled those originating in r2. This suggests
that a subset of ventral motor neuron precursors may be incorrect specified. In addition,
development of the somatic motor component of the facial nerve is impaired in Hoxb2 null
mutants, which is reminiscent of the phenotype described in the Hoxb1 mutants. Hoxb2 therefore
plays an important role in regulating the generation and/or fate of subsets of r4 motor neuron
progenitors.
The generation of Hoxa2/Hoxb2 double mutants revealed that synergistically these genes
play an important role in coupling A-P and D-V patterning[115]. Hoxa2 and Hoxb2 play distinct
roles in mediating hindbrain neurogenesis. Hoxa2 regulates development in the alar and dorsal
basal plates of r2 and r3 whereas Hoxb2 is essential for motorneuron development in the ventral
region of r4. In the double mutants, functional synergy between Hoxa2 and Hoxb2 in r3 has been
suggested based on the absence of a ventral interneuron subtype, which is developmentally
normal in both the single mutants. In addition there also appears to be a joint requirement for
Hoxa2 and Hoxb2 in patterning inter-rhombomeric boundaries. Despite the fact that the normal
number of hindbrain segments form in the double mutants, boundary formation between r1-4 is
nonexistent. Therefore these results indicate that Hoxa2 and Hoxb2 differentially control alar and
basal plate development within distinct rhombomeres. Perhaps more importantly this provides a
link between Hox mediated patterning along the A-P axis and neurogenesis along the D-V axis
and can account mechanistically for the generation of neurons at reproducible positions within
hindbrain segments.
A targeted mutation in the Hoxa3 gene results primarily in mesenchymal neural crest
defects[116]. The Hoxa3 null mutant mice are athymic, aparathyroid, and have throat cartilage
malformations. In addition, the loss of Hoxa3 leads to defects in the formation of the IXth cranial
nerve and also fusions between the IXth and Xth nerves. Examination of the cranial nerves in
Hoxb3-/- embryos revealed similar cranial ganglia defects but at a lower penetrance than in the
Hoxa3 mutants[117]. In contrast, the cranial nerves in the Hoxd3-/- embryos are completely
normal[118,119]. Therefore at least two of the group 3 paralogs play essential roles in formation
of the IXth cranial ganglion. Double mutants (Hoxa3/Hoxb3, Hoxa3/Hoxd3, and Hoxb3/Hoxd3)
were subsequently generated and in Hoxb3/Hoxd3 double mutants there was a marked increase in
the penetrance of the IXth cranial ganglion defects relative to that observed in the Hoxb3 single
mutant[118,119,120]. This increase in penetrance shows that Hoxd3 plays a synergistic role with
Hoxb3 in cranial nerve development even though the Hoxd3 single mutant does not show a defect
in these structures.
247

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
KROX20 NULL MUTATIONS
The necessity for coupling cell identity and cell lineage restrictions in order to generate sharply
restricted and specified segmental domains during hindbrain development is highlighted in the
phenotype of Krox20 null mutants. Mice homozygous for a targeted mutation in Krox20 die
shortly after birth and exhibit fusions of the trigeminal ganglion with facial and vestibular ganglia
as a consequence of the profound perturbation of hindbrain morphogenesis[26,27,121]. Although
the presumptive territories of rhombomeres 3 and 5 do form during the very early stages of
hindbrain development, these two rhombomeres are not maintained and their structural
derivatives are subsequently eliminated. Consistent with the interactions unveiled by transgenic
analyses between Krox20 and Hoxa2, Hoxb2, Hoxb3, and EphA4, no upregulation of these three
genes was observed in Krox20 mutants. Therefore Krox20 is a key regulator of gene expression
and segmentation in the developing hindbrain.
KREISLER mutants
The kreisler mutation is not a null allele, but a regulatory mutation affecting those enhancer
elements responsible for the hindbrain domain of expression of the gene[66]. The primary defect
in kreisler mutant embryos is a disruption of segmentation in the otic region of the hindbrain,
whereby the rhombomeric borders that normally demarcate r4, 5, and 6 are absent[32,65,122].
Consequently the normal expression domains of Fgf3 and CRABPI and the upregulation of
Hoxa3 in r6 are absent. Although Krox20 expression in presumptive r3 is present, the posterior
band associated with r5 is missing. Similarly, the normal expression domains of Hoxb2, Hoxb3,
and Hoxb4 in r5 are also abolished. Analysis of patterns of expression of EphA7 and ephrinB2
indicate that only a single segment correlating with r5 is missing. Therefore the segmentation
defect associated with the kreisler mutant is a specific loss of r5 and although an r6 territory does
form, it fails to mature[32]. Through direct regulation of Hox genes, kreisler controls segmental
identity and this is further evidenced by the ectopic activation of kreisler in r3 which transforms
the character of this territory into that of r5[123]. The role played by kreisler in hindbrain
patterning is therefore analagous to Krox20 because rather than being restricted to a single
patterning aspect, kreisler is involved at several different levels including establishment and
maintenance of A-P identity in addition to segment specification.
Clearly hindbrain segmentation in the form of cell lineage restrictions and
compartmentalization is coupled with the acquisition of specific cellular identities. The studies
described above illustrate that Hox genes function in many steps of hindbrain segmentation
process particularly in specifying segmental identity. Consequently hindbrain segmentation and
rhombomere specification plays important roles in patterning the craniofacial structures of neural
crest origin.
CRANIAL NEURAL CREST CELLS AND THE PREPROGRAMMING MODEL
FOR CRANIOFACIAL DEVELOPMENT
Neural crest cells were so named because of their formation along almost the entire axis of
vertebrate embryos at the crest of the closing neural folds. This region corresponds to the
interface or junction between the non-neural ectoderm (presumptive epidermis or surface
ectoderm) and the neural plate (neuroepithelium), a region commonly referred to as the neural
plate border (Fig. 2). Neural crest cell induction requires contact-mediated interactions between
the surface ectoderm and neuroepithelium and importantly, each of these tissues contributes to the
neural crest cell lineage[124]. BMP signalling is thought to play critical roles in both positioning
248

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
the border of the neural plate during gastrulation and then also later in inducing migration of
neural crest cells[125]. Cranial neural crest cells give rise the neurons and glia of the PNS and
most of the cartilage, bone, and connective tissue of the face. The cartilage and bone forming
properties of cranial neural crest cells distinguishes this population from trunk neural crest cells.
Hence the cranial neural crest is a pluripotent population that is vital to vertebrate head
development (Fig. 2)[126,127,128,129,130,131]. It is essential to understand the mechanisms by
which cranial neural crest cells are patterned since many craniofacial malformations can be
attributed to defects in the formation, proliferation, migration, and/or differentiation of this cell
population.
The generation of regional diversity in the vertebrate head is believed to be a consequence of
patterning information provided by neural crest cells during their migration and neural fold
transplantation experiments have provided much of the basis for our current understanding of
neural crest and craniofacial development[132,133,134,135,136]. First arch neural crest cells give
rise to Meckel’s cartilage, the articular, quadrate, and squamosal bones. In contrast the second
arch neural crest cells develop into the hyoid or tongue cartilage and the retroarticular
process[9,136,137]. In posterior transplantations of first arch (mandibular) neural crest primordia
in place of second (hyoid) or third (visceral) arch neural crest, the transplanted neural crest cells
migrated into the nearest arch but therein formed ectopic duplicated proximal first arch skeletal
elements[136]. Not only were these duplicated crest-derived structures inappropriate for their new
location but the muscle cell types and attachments associated with the duplicated structures were
also characteristic of a first arch pattern. This suggested firstly that neural crest cell fate may be
preprogrammed prior to their emigration from the neural tube and secondly, that myogenic
populations and other cell types receive spatial cues from the invading neural crest-derived
connective tissue. The vast majority of cranial neural crest cells are derived from the hindbrain
and molecular evidence supporting this scenario was provided by the observation that the same
restricted domains of Hox gene expression in the hindbrain are emulated in the migrating neural
crest cells and then later in the ganglia and branchial arches, reflecting the origins of the neural
crest cells contributing to these tissues[4,51]. Under this preprogramming model, it was thought
that positional information encoded by the Hox genes was carried passively from the hindbrain to
peripheral tissues and branchial arches by the neural crest, where it was elaborated to form the
characteristic head structures. Therefore it was hypothesized that the spatial organization of
structures within the vertebrate head was determined by the neural crest and furthermore that the
positional information imparted by the neural crest was irreversibly set before the neural crest
emigrates from the neural tube.
CRANIAL NEURAL CREST PLASTICITY AND INDEPENDENT HOX GENE
REGULATION MODEL
The neural crest prepatterning model predicts that molecular and cellular alterations to the
segmental organization of the hindbrain would have profound and detrimental consequences to
craniofacial development via disruptions to the patterns of Hox gene expression and neural crest
cell migration. The chick embryo has been the primary species for testing this model via
rhombomere transplantations, rotations, and ablations owing to the ease of tissue manipulation in
this species[112,138,139,140,141,142]. These analyses have yielded conflicting results regarding
the degree of autonomy of Hox gene expression[143]. The recent application of techniques for
transposing cells to the hindbrains of mouse[144] and zebrafish[145] embryos has significantly
advanced our understanding of hindbrain and neural crest development. In the mouse, cells from
r3, r4, or r5 were heterotopically grafted into r2. The majority of the transplanted cells remained
as a cohort and maintained their Hox gene A-P identity. A few transplanted cells however,
became separated from the primary graft and dispersed becoming intermingled with the
249

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
neighboring populations. These cells consequently altered their identity in their new location by
failing to maintain the Hox gene expression patterns characteristic of their origins, which is
evidence for neuroepithelial plasticity. This implies that single or dispersed rhombomere cells
respond and adapt to their new surrounding environments by altering their gene expression
patterns because they lack the neighboring signals necessary to reinforce or maintain their
original identity[144]. Similarly, single rhombomere cell transplantations in zebrafish provide
further evidence for neural crest plasticity and the influence of cell community effects[145]. A
complete switch in Hox gene expression patterns and ultimately cell fates occurred when single
cells from r2 and r6 were reciprocally transplanted within the zebrafish hindbrain. This contrasts
with the higher degrees of cell autonomy observed in chick embryo manipulations. It is important
to note however that, in analyses performed in avian embryos, pairs of rhombomeres but
generally much larger regions of the neural tube were being manipulated. It is not unreasonable to
expect that cell community effects present in such large tissue grafts could account for the cell
autonomy observed in these avian studies compared to the plasticity observed in mouse and
zebrafish experiments where subrhombomeric or single cell populations were transplanted,
respectively[143]. The degree of plasticity or autonomy is dependent upon not only the size of the
graft but also the time at which the manipulations are carried out. During the later stages of
hindbrain development, when morphological boundaries are established, rhombomere cells are
much more likely to be irreversibly committed and maintain their Hox gene expression
characteristics when transplanted to ectopic locations. Therefore one could argue that over time
cells within the neural tube progressively lose responsiveness to the environmental signals that
specify their segmental identities. Together these chick, mouse and fish studies highlight that cell-
community effects and their associated signals are important for maintaining the axial identity of
individual cells within the hindbrain.
These grafting experiments also revealed the absence of preprogramming in the character or
fate of cranial neural crest cells. In heterotopic transpositions of cells within the mouse and
zebrafish hindbrains, graft-derived neural crest cells migrated into the nearest branchial arch
without any evidence of pathfinding or rerouting to their original axial level. Plasticity in Hox
gene expression in mouse neural crest cells was evident by the complete down regulation of
Hoxb1, Hoxb2, and Hoxa2 in these cells[144]. In zebrafish, experimental embryos raised to larval
stages revealed that the transplanted cells differentiated and contributed to pharyngeal cartilages
appropriate to their new A-P location[145]. Therefore these results show that the genetic and
cellular A-P character of cranial neural crest cells are neither fixed nor passively transferred from
the hindbrain to the branchial arches.
The neural plasticity described above correlates with molecular analyses that have identified
distinct regulatory elements controlling Hox gene expression in different tissues such as the
hindbrain and neural crest. Hoxa2 is expressed in the hindbrain anteriorly up to the r1/2 boundary
and in cranial neural crest cells that migrate into the second branchial arch[112]. Transgenic
regulatory analyses of Hoxa2 have revealed that multiple cis-acting elements are required
independently for hindbrain and neural crest specific activity[55,146,147]. In r3 and r5, Hoxa2
expression is directly regulated by the transcription factor Krox20[55]. In contrast, Hoxa2
expression in second branchial arch neural crest cells is tightly controlled by a number of
elements, one of which binds to AP-2 family members. Mutation or deletion of this site in the
Hoxa2 enhancer abrogates expression in cranial neural crest cells but not in the hindbrain. These
findings clearly demonstrate that at the molecular level, Hoxa2 is independently regulated in
rhombomeres and neural crest cells and this provides a mechanism for how neural crest cells can
respond to the environment through which they migrate independently of the neural tube.
These experiments have revealed a surprising degree of plasticity in cranial neural crest
cells, which is inconsistent with the prepatterning model. Since transposed neural crest can be
reprogrammed it appears that crest cells rely on distinct cues in the branchial arch environments
through which they migrate to elaborate their proper regional identity. Furthermore, the size of
250

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
FIGURE 2. Neural crest cell induction and differentiation. Neural crest cells (ncc; blue circles) are induced at the neural plate border,
which demarcates the junction between the neural plate and surface ectoderm. Both the neuroepithelium and the surface ectoderm
gives rise to neural crest cells and contact between these tissues is essential for neural crest induction. In the mouse as depicted here,
this occurs prior to closure of the neural tube. In contrast, in avian species the neural tube is closed prior to the emigration of neural
crest cells. After migrating along stereotypic routes, the cranial neural crest cells differentiate into cartilage, bone, connective tissue as
well as neurons and glia of the PNS. In contrast, trunk neural crest cells do not give rise to cartilage or bone rather they differentiate
predominantly into the characteristic cells that constitute the trunk PNS.
the cell community is functionally important, indicating that a far more complex balance of
genetic and cellular interactions are involved in hindbrain and neural crest patterning than was
previously thought. The experiments described above therefore provide the basis for a new
model, the neural crest plasticity and independent Hox gene regulation model which has been
proposed as an alternative mechanism for governing neural crest and craniofacial
development[143,144,148].
251

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
EVOLUTION AND PATTERNING OF NEURAL CREST CELLS
The classic neural crest preprogramming model implies that branchial arch formation and
patterning is dependent upon the neural crest. In contrast, the neural crest plasticity and
independent gene regulation model described above implies that branchial arch patterning arises
due to interactions between the arch components and the neural crest. This raises the question of
what happens to the formation and patterning of the branchial arches in the absence of
contributing neural crest cells. This issue has been investigated in chick embryos through
rhombomere ablations[149] and also mouse embryos by genetic manipulation of Hoxa1 and
Hoxb1, which are required for the generation of neural crest cells in r4[33]. In both experimental
situations, despite the absence of any neural crest cell contribution, the branchial arches develop
normally and are properly regionalized. The expression patterns of Bmp7 in the posterior
endoderm, Fgf8 in the anterior surface ectoderm, Pax1 in the pharyngeal pouch endoderm and
Shh in the endoderm of the branchial arches were all normal and unchanged compared to wild
type embryos. In addition there was no evidence for excessive cell death or loss of proliferation in
the arch epithelium, which suggests that the neural crest cells are not the source of any
indispensable branchial arch mitogenic or survival signals[33]. These results clearly demonstrate
that the branchial arches are not dependent upon the neural crest for their formation, nor for their
anterior-posterior and proximo-distal regionalization and hence this provides additional support
for the neural crest plasticity and independent gene regulation model. This is also consistent with
the evolutionary history of the branchial arches and neural crest cells. Pharyngeal segmentation is
characteristic of the phylum chordata whereas neural crest cells are exclusively a craniate
(vertebrates plus hagfish) characteristic[150], which implies that branchial arch segmentation
occurred prior to the evolutionary origin of cranial neural crest cells. Hence it is not surprising
that the branchial arches do not depend upon the cranial neural crest for their initial formation and
regional specification. Additional support for this idea comes from the observation of
regionalized domains of Pax gene expression in Amphioxus (the nearest extant vertebrate
relative), which is indicative of pharyngeal segmentation. Amphioxus lack neural crest cells and
therefore the mechanism for generating pharyngeal pouches clearly predates the evolution of the
vertebrate head[151].
PATTERNING ROLES FOR THE MESODERM, ENDODERM, AND
ECTODERM IN BRANCHIAL ARCH DEVELOPMENT
A crucial issue in craniofacial development is understanding the mechanisms that regulate the
size and shapes of the characteristic facial skeletal structures and that branchial arch segmentation
constitutes one of the first visible steps in this patterning process. Since normal branchial arch
formation and patterning can occur independently of a contribution from the neural crest, this
implies that the branchial arches may rely on either the endoderm, paraxial mesoderm, and/or the
surface ectoderm tissues for their patterning information.
The Mesoderm
Fate mapping studies have shown that mesoderm and neural crest cells derived from the same
axial level contribute to the same branchial arch during embryonic development (Fig.
3)[16,130,152,153]. The cranial mesoderm predominantly gives rise to the myogenic cores of
each branchial arch, which are enveloped by migrating neural crest cells[16,130,131,153,154].
Previously, the cranial mesoderm was not thought to play a patterning role during craniofacial
development[136], however, it has now been shown that the cranial mesoderm provides
252

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
maintenance signals for regulating the identity of second branchial arch neural crest cells[144]
and that it may also play a role in patterning the pathways of neural crest cell migration (Fig.
3)[23]. When second arch neural crest cells are transplanted into the first arch, they downregulate
their expression of Hoxb1. In contrast, if second arch neural crest cells are transplanted anteriorly
in combination with second arch mesoderm, then Hoxb1 expression is maintained in the grafted
neural crest cells. The cranial mesoderm therefore provides maintenance signals that elaborate the
programme of Hox expression, but the cranial mesoderm does not appear to initiate Hox gene
expression in neural crest cells[144]. It is important to note however that Hoxb1 is expressed in
the cranial mesoderm at 7.5dpc prior to its induction in the neuroepithelium suggesting that
perhaps the mesoderm is patterning neuroepithelial tissues at much earlier stages of development.
The effects of the mesoderm are consistent with the fact that the fate of the cranial mesoderm is
primarily myogenic and the musculature is inextricably linked to neural crest derived skeletal and
connective tissue patterning. Therefore one of the roles of the cranial mesoderm maybe in
maintaining an A-P register between these different primordial tissues, which is essential for
subsequent craniofacial morphogenesis[16].
The Ectoderm
Similar to the neuroepithelium, it has been suggested that the ectoderm is regionalized into
territories called ectomeres, which contribute to specific regions of the branchial arches (Fig.
3)[155]. Currently, there is no evidence to support the idea that each ectomere represents a
functional developmental unit. In contrast, however, there is evidence suggesting that the surface
ectoderm plays a major role in the induction of odontogenesis during branchial arch
development[156]. The oral ectoderm of the first branchial arch directly regulates the patterning
of the underlying neural crest mesenchyme into teeth and the ability to respond to these
instructive or inducing signals is not confined to first arch neural crest cells[157]. Fgf8, which is
expressed in the anterior surface ectoderm of the first arch, is essential for determining the
polarity of the branchial arch and ectopic applications of FGF8 cause shifts in gene expression
domains as well as repatterning of the craniofacial primordia[158]. Not surprisingly then, in Fgf8
null mutant mice, the branchial arches are severely abnormal[159]. Bmp4, which is expressed in
the ventral region of the first branchial arch ectoderm, appears to restrict the expression domain
of Fgf8 and consequently ectopic applications of BMP4 consistently reduce the size of the
mandibular arch. Hence the surface ectoderm plays important roles in patterning the branchial
arch derivatives particularly through the BMP4 and FGF8 signalling mechanisms (Fig. 3).
The Endoderm
The neurogenic placodes (dorsolateral and epibranchial) form in characteristic positions in all
vertebrates suggesting that conserved localized inductive interactions underlie their
formation[160]. The epibranchial placodes develop near the branchial clefts in close proximity to
the cranial neural crest and the pharyngeal endoderm. Analyses of the nature of the signals, which
underlie epibranchial placode formation, have found that the epibranchial placodes do not require
cranial neural crest cells for their induction[161]. Rather, it is the pharyngeal endoderm that is the
source of the BMP7 inducing signal. It has been suggested that neural crest cells will differentiate
into cartilage only in the presence of pharyngeal endoderm and in amphibians, the endoderm has
been shown to be responsible for promoting the formation of branchial arch components by
directing neural crest cells towards a chondrogenic fate[162]. For example, in the zebrafish
mutant van gogh (vgo), hindbrain segmentation occurs normally, but the endodermal gill slits do
not form and the distinct streams of neural crest cells exiting from the hindbrain are fused.
253

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
Consequently the individual skeletal elements of the viscerocranium do not form[163]. Therefore
the endoderm plays a major role in establishing and patterning the branchial arches (Fig. 3).
Recently the avian neural endoderm was tested in transplantation and ablation studies for its
capacity to specify the facial skeleton[164]. The experiments suggested that the endoderm
instructs neural crest cells as to the size, shape, and position of all the skeletal elements whether
they are cartilaginous or membranous bones. In addition bone orientation was shown to be
influenced by the position of the endoderm relative to the embryonic axes. The nature of the
signals arising from the endoderm are so far unknown and it is not clear whether the effects of the
endoderm manipulations are direct and intrinsic to the tissue itself or whether the effects of the
manipulations indirectly alter local signaling centers or levels of FGF, Shh, or BMP which have
been shown in other analyses to regulate the development of the characteristic craniofacial
structures[159,165,166,167]. What is clear is that segmental characteristics of the endodermal
pharyngeal pouches develop independently of a contribution of neural crest cells (Fig. 3)[149].
FIGURE 3. The influence of mesoderm, ectoderm, and endoderm on the axial identity of neural crest cells and the role of Hoxa2 in
second branchial arch development. The head mesoderm (red and blue ovals) plays a role in maintaining the proper domains of Hox
gene expression in migrating neural crest cells (ncc), whereas the ectoderm and endoderm play roles in possibly influencing the
pathways of neural crest cell migration in addition to patterning the branchial arches. The skeletal derivatives arising from the first
(ba1) and second (ba2) branchial arches are listed. Hoxa2 is expressed in neural crest cells migrating into the second arch but not the
first arch acts and as a selector gene that imposes the unique identity of second arch structures. In the absence of Hoxa2, second arch
derivatives are transformed into a first arch fate and similarly ectopic expression of Hoxa2 transforms first arch structures into a
second arch identity. FGF8 has the capacity to repress Hoxa2 in second branchial arch neural crest cells, transforming the second arch
into a first arch identity. OV, otic vesicle.
254

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
RESOLVING THE ISSUES OF NEURAL CREST PLASTICITY VS.
PREPATTERNING AND SKELETAL DUPLICATIONS
The analyses detailed above provide a wealth of evidence supporting the idea that cranial neural
crest cells are not preprogrammed, but are in fact plastic and that they differentiate in response to
signals from the environment through which they migrate. How then do we reconcile these
findings with the skeletal duplications observed in posterior transplantations of presumptive first
arch neural crest[136]? Two things are often ignored from this classic study. Firstly, in addition to
forming duplicated first arch structures, the transplanted neural crest also contributed to the
development of normal second arch skeletal elements including the paraglossals and basihyoid,
which make up part of the tongue skeleton. This actually provides evidence for neural crest
plasticity. Secondly, when presumptive frontonasal neural crest was grafted posteriorly in place
of second arch neural crest, duplicated first arch skeletal elements (quadrate and proximal region
of Meckel’s cartilage) also developed, providing further evidence in support of neural crest
plasticity[136]. It’s important to note here that the same duplicated skeletal structures were
formed regardless of the origin of the grafted neural crest cells.
What’s intriguing about the skeletal duplications is that they phenocopy the Hoxa2 null
mutant mouse in which the second arch is transformed into a first arch identity and one can
speculate then that perhaps the landmark chick embryo manipulations were in effect creating a
conditional knockout of Hoxa2 in the second branchial arch. Recently, the isthmus was shown to
inhibit Hoxa2 expression in rhombomere 1 via an FGF8 mediated signalling mechanism[168],
and therefore one plausible explanation which could link the results of the two transplantations
together is the possible inclusion of the isthmus in both the first arch and frontonasal neural crest
grafts. This has now been tested directly via posterior transplantations of the isthmus in place of
r4 that inhibits the expression of Hoxa2 in second branchial arch neural crest cells[169]. In the
absence of Hoxa2 expression, as expected these grafted chick embryos develop duplicated first
arch skeletal structures including the quadrate and proximal portion of Meckel’s cartilage similar
to the classic transplantations. FGF8 soaked beads can only transiently block Hoxa2 expression in
second branchial arch neural crest cells indicating that FGF8 cannot recapitulate the entire effects
of the isthmus and that other genes/factors must be involved.
Therefore the possible inclusion of the isthmus in the transpositions and the ability of FGF8
to suppress Hoxa2 in second arch neural crest cells can account mechanistically for the
development of duplicated first arch structures in ectopic posterior locations. Hence rather than
providing evidence for neural crest preprogramming, Noden’s 1983 experiments highlight the
effects of local signaling centers like the isthmus in A-P patterning and regulation of Hox gene
expression by FGFs[136]. Together with recent evidence from mouse, chick, and zebrafish
transplantation studies, this argues as a general principle that cranial neural crest cells are not
prespecified or irreversibly committed prior to their emigration from the neural tube. Rather, that
neural crest patterning is based on plasticity and the ability of neural crest cells to respond to
environmental signals and interactions with the tissues through which they migrate[144,148].
Recently inroads have been made into the mechanisms by which Hoxa2 influences the
morphogenesis of second arch elements[170]. During normal development, Hoxa2 is widely
expressed in the second arch mesenchyme, but it is excluded from the chondrogenic
condensations in the core of the arches (Fig. 3). In the absence of Hoxa2, ectopic chondrogenesis
occurs indicating that Hoxa2 acts upstream very early in the chondrogenic pathway. In addition,
Cbaf1, which is a marker of osteogenic differentiation, is upregulated in the second branchial
arches of Hoxa2 mutant embryos suggesting that the prevention of Cbfa1 induction might
255

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
mediate Hoxa2 inhibition of dermal (intramembranous) bone formation during second arch
development[170]. Therefore Hoxa2 is essential for proper patterning of structures derived form
the second branchial arch neural crest cells as it inhibits membranous ossification. Ectopic
expression of Hoxa2 in the first arch of chick and Xenopus embryos results in the suppression of
bone formation in the first arch and transformation of the first arch into second arch
structures[171,172]. In the absence of Hoxa2 the second arch is transformed into a first arch
identity and together these results indicate that Hoxa2 acts as a positive selector gene in
specifying second branchial arch development and identity (Fig. 3).
EVOLUTIONARY IMPORTANCE AND CONCLUSIONS
Craniofacial evolution is considered fundamental to the origin of vertebrates and in evolutionary
terms the vertebrate head is a relatively new structure[173]. This review has detailed the multiple
levels of regulation and the diverse tissue interactions that are involved in generating the
characteristic craniofacial features. Although the hindbrain exerts a profound influence in
establishing the foundations of vertebrate head development, a rigid prepatterning model in which
the programs for head morphogenesis were set in the hindbrain would however offer restricted
opportunities for diversifying head structures[143,144,148]. In contrast the neural crest plasticity
and independent gene regulation model could provide the flexibility and adaptability that
facilitates diversity and we can speculate that it might be one reason for the successful radiation
of vertebrates into new environments. This is because neural crest plasticity and independent gene
regulation offers the potential for generating substantially distinct cranial phenotypes by minor
changes of the primordial pattern. Future studies therefore will be focused on elucidating the
mechanisms governing the formation and patterning of neural crest cells as they are critical to
understanding craniofacial evolution and the origins of vertebrates.
REFERENCES
1. Vaage, S. (1969) The segmentation of the primitive neural tube in chick embryos (Gallus domesticus). Adv.
Anat. Embryol. Cell Biol. 41, 1–88.
2. Wilkinson, D.G., Bhatt, S., Chavrier, P., Bravo, R., and Charnay, P. (1989) Segment-specific expression of a
zinc finger gene in the developing nervous system of the mouse. Nature 337, 461–464.
3. Wilkinson, D. (1989) Homeobox genes and development of the vertebrate CNS. Bioessays 10, 82–85.
4. Hunt, P., Whiting, J., Muchamore, I., Marshall, H., and Krumlauf, R. (1991) Homeobox genes and models
for patterning the hindbrain and branchial arches. Development 112(Suppl.), 187–196.
5. Fraser, S., Keynes, R., and Lumsden, A. (1990) Segmentation in the chick embryo hindbrain is defined by
cell lineage restrictions. Nature 344, 431–435.
6. Birgbauer, E. and Fraser, S.E. (1994) Violation of cell lineage restriction compartments in the chick
hindbrain. Development 120, 1347–1356.
7. Marín, F. and Puelles, L. (1995) Morphological fate of rhombomeres in quail/chick chimeras: a segmental
analysis of hindbrain nuclei. Eur. J. Neurosci. 7, 1714–1738.
8. Wintgate, R. and Lumsden, A. (1996) Persistence of rhombomeric organisation in the postsegmental avian
hindbrain. Development 122, 2143–2152.
9. Köntges, G. and Lumsden, A. (1996) Rhombencephalic neural crest segmentation is preserved throughout
craniofacial ontogeny. Development 122, 3229–3242.
10. Lumsden, A. and Keynes, R. (1989) Segmental patterns of neuronal development in the chick hindbrain.
Nature 337, 424–428.
256

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
11. Clarke, J.D. and Lumsden, A. (1993) Segmental repetition of neuronal phenotype sets in the chick embryo
hindbrain. Development 118, 151–162.
12. Sechrist, J., Serbedzija, G.N., Scherson, T., Fraser, S.E., and Bronner-Fraser, M. (1993) Segmental migration
of the hindbrain neural crest does not arise from its segmental generation. Development 118(3), 691–703.
13. Lumsden, A., Sprawson, N., and Graham, A. (1991) Segmental origin and migration of neural crest cells in
the hindbrain region of the chick embryo. Development 113, 1281–1291.
14. Serbedzija, G., Fraser, S., and Bronner-Fraser, M. (1992) Vital dye analysis of cranial neural crest cell
migration in the mouse embryo. Development 116, 297–307.
15. Osumi-Yamashita, N., Ninomiya, Y., Doi, H., and Eto, K. (1994) The contribution of both forebrain and
midbrain crest cells to the mesenchyme in the frontonasal mass of mouse embryos. Dev. Biol. 164, 409–419.
16. Trainor, P.A. and Tam, P.P.L. (1995) Cranial paraxial mesoderm and neural crest of the mouse embryo-
codistribution in the craniofacial mesenchyme but distinct segregation in the branchial arches. Development
121, 2569–2582.
17. Graham, A., Heyman, I., and Lumsden, A. (1993) Even-numbered rhombomeres control the apoptotic
elimination of neural crest cells from odd-numbered rhombomeres in the chick hindbrain. Development 119,
233–245.
18. Graham, A., Francis-West, P., Brickell, P., and Lumsden, A. (1994) The signalling molecule BMP4 mediates
apoptosis in the rhombencephalic neural crest. Nature 372, 684–686.
19. Ellies, D.L., Church, V., Francis-West, P., and Lumsden, A. (2000) The WNT antagonist cSFRP2 modulates
programmed cell death in the developing hindbrain. Development 127, 5285–5295.
20. Schilling, T.F. and Kimmel, C.B. (1994) Segment and cell type lineage restrictions during pharyngeal arch
development in the zebrafish embryo. Development 120, 483–494.
21. Kulesa, P. (1998) Neural crest cell dynamics revealed by time-lapse video microscopy of whole chick
explant cultures. Dev. Biol. 204, 327–344.
22. Kulesa, P.M. and Fraser, S.E. (2000) In ovo time-lapse analysis of chick hindbrain neural crest cell migration
shows cell interactions during migration to the branchial arches. Development 127, 1161–1172.
23. Trainor, P.A., Sobieszczuk, D., Wilkinson, D., and Krumlauf, R. (2002) Signalling between the hindbrain
and paraxial tissues dictates neural crest migration pathways. Development 129, 433–442.
24. Lufkin, T., Dierich, A., LeMeur, M., Mark, M., and Chambon, P. (1991) Disruption of the Hox-1.6
homeobox gene results in defects in a region corresponding to its rostral domain of expression. Cell 66,
1105–1119.
25. Chisaka, O., Musci, T., and Capecchi, M. (1992) Developmental defects of the ear, cranial nerves and
hindbrain resulting from targeted disruption of the mouse homeobox gene Hox-1.6. Nature 355, 516–520.
26. Swiatek, P.J. and Gridley, T. (1993) Perinatal lethality and defects in hindbrain development in mice
homozygous for a targeted mutation of the zinc finger gene Krox-20. Genes Dev. 7, 2071–2084.
27. Schneider-Maunoury, S., Topilko, P., Seitanidou, T., Levi, G., Cohen-Tannoudji, M., Pournin, S., Babinet,
C., and Charnay, P. (1993) Disruption of Krox-20 results in alteration of rhombomeres 3 and 5 in the
developing hindbrain. Cell 75, 1199–1214.
28. Gassmann, M., Casagranda, F., Orioli, D., Simon, H., Lai, C., Klein, R., and Lemke, G. (1995) Aberrant
neural and cardiac development in mice lacking the ErbB4 neuregulin receptor. Nature 378, 390–394.
29. Meyer, D. and Birchmeier, C. (1995) Multiple essential functions of neuregulin in development. Nature 378,
386–390.
30. Goddard, J., Rossel, M., Manley, N., and Capecchi, M. (1996) Mice with targeted disruption of Hoxb1 fail to
form the motor nucleus of the VIIth nerve. Development 122, 3217–3228.
31. Studer, M., Lumsden, A., Ariza-McNaughton, L., Bradley, A., and Krumlauf, R. (1996) Altered segmental
identity and abnormal migration of motor neurons in mice lacking Hoxb-1. Nature 384, 630–635.
32. Manzanares, M., Trainor, P., Nonchev, S., Ariza-McNaughton, L., Brodie, J., Gould, A., Marshall, H.,
Morrison, A., Kwan, C.-T., Sham, M.-H., et al. (1999) The role of kreisler in segmentation during hindbrain
development. Dev. Biol. 211, 220–237.
257

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
33. Gavalas, A., Trainor, P., Ariza-McNaughton, L., and Krumlauf, R. (2001) Synergy between Hoxa1 and
Hoxb1: the relationship between arch patterning and the generation of cranial neural crest. Development 128,
3017–3027.
34. Guthrie, S. and Lumsden, A. (1991) Formation and regeneration of rhombomere boundaries in the
developing chick hindbrain. Development 112, 221–229.
35. Guthrie, S., Prince, V., and Lumsden, A. (1993) Selective dispersal of avian rhombomere cells in orthotopic
and heterotopic grafts. Development 118, 527–538.
36. Wizenmann, A. and Lumsden, A. (1997) Segregation of rhombomeres by differential chemoaffinity. Mol.
Cell. Neurosci. 9, 448–459.
37. Wilkinson, D.G. (1995) Genetic control of segmentation in the vertebrate hindbrain. Perspect. Dev.
Neurobiol. 3, 29–38.
38. Lumsden, A. and Krumlauf, R. (1996) Patterning the vertebrate neuraxis. Science 274, 1109–1115.
39. Trainor, P., Manzanares, M., and Krumlauf, R. (2000) Genetic interactions during hindbrain segmentation in
the mouse embryo. In Mouse Brain Development: Results and Problems in Cell Differentiation. Vol. 30.
Goffinet, A. and Rackic, P., Eds. Springer-Verlag, Berlin. pp. 51–89.
40. Wilkinson, D.G. (2001) Multiple roles of EPH receptors and ephrins in neural development. Nat. Rev.
Neurosci. 2, 155–164.
41. Nieto, M.A., Gilardi-Hebenstreit, P., Charnay, P., and Wilkinson, D. (1992) A receptor protein tyrosine
kinase implicated in the segmental patterning of the hindbrain and mesoderm. Development 116, 1137–1150.
42. Becker, N., Seitanidou, T., Murphy, P., Mattei, M.-G., Topilko, P., Nieto, M.A., Wilkinson, D.G.,
Charnay, P., and Gilardi-Hebenstreit, P. (1994) Several receptor tyrosine kinase genes of the Eph
family are segmentally expressed in the developing hindbrain. Mech. Dev. 47, 3–18.
43. Gale, N.W., Holland, S.J., Valenzuela, D.M., Flenniken, A., Pan, L., Ryan, T.E., Henkemeyer, M.,
Strebhardt, K., Hirai, H., Wilkinson, D.G., et al. (1996) Eph receptors and ligands comprise two
major specificity subclasses and are reciprocally compartmentalized during embryogenesis. Neuron
17, 9–19.
44. Xu, Q., Mellitzer, G., Robinson, V., and Wilkinson, D. (1999) In vivo cell sorting in complementary
segmental domains mediated by Eph receptors and ephrins. Nature 399, 267–271.
45. Mellitzer, G., Xu, Q., and Wilkinson, D. (1999) Eph receptors and ephrins restrict cell intermingling and
communication. Nature 400, 77–81.
46. McGinnis, W. and Krumlauf, R. (1992) Homeobox genes and axial patterning. Cell 68, 283–302.
47. Kappen, C., Schugart, K., and Ruddle, F. (1989) Two steps in the evolution of antennapedia-class vertebrate
homeobox Genes. Proc. Natl. Acad. Sci. U. S. A. 86, 5459–5463.
48. Duboule, D. and Dolle, P. (1989) The structural and functional organization of the murine HOX gene family
resembles that of Drosophila homeotic genes. EMBO J. 8, 1497–1505.
49. Dollé, P., Izpisùa-Belmonte, J.C., Falkenstein, H., Renucci, A., and Duboule, D. (1989) Co-ordinate
expression of the murine Hox-5 complex homeobox-containing genes during limb pattern formation. Nature
342, 767–772.
50. Kessel, M. and Gruss, P. (1991) Homeotic transformations of murine prevertebrae and concommitant
alteration of Hox codes induced by retinoic acid. Cell 67, 89–104.
51. Hunt, P., Gulisano, M., Cook, M., Sham, M., Faiella, A., Wilkinson, D., Boncinelli, E., and Krumlauf, R.
(1991) A distinct Hox code for the branchial region of the head. Nature 353, 861–864.
52. Wilkinson, D.G., Bhatt, S., Cook, M., Boncinelli, E., and Krumlauf, R. (1989) Segmental expression of Hox-
2 homeobox-containing genes in the developing mouse hindbrain. Nature 341, 405–409.
53. Frohman, M.A., Boyle, M., and Martin, G.R. (1990) Isolation of the mouse Hox-2.9 gene; analysis of
embryonic expression suggests that positional information along the anterior-posterior axis is specified by
mesoderm. Development 110, 589–607.
54. Murphy, P. and Hill, R.E. (1991) Expression of the mouse labial-like homeobox-containing genes, Hox 2.9
and Hox 1.6, during segmentation of the hindbrain. Development 111, 61–74.
258

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
55. Nonchev, S., Vesque, C., Maconochie, M., Seitanidou, T., Ariza-McNaughton, L., Frain, M., Marshall, H.,
Sham, M.H., Krumlauf, R., Charnay, P. (1996) Segmental expression of Hoxa-2 in the hindbrain is directly
regulated by Krox-20. Development 122, 543–554.
56. Sham, M.H., Vesque, C., Nonchev, S., Marshall, H., Frain, M., Das Gupta, R., Whiting, J., Wilkinson, D.,
Charnay, P., and Krumlauf, R. (1993) The zinc finger gene Krox-20 regulates Hoxb-2 (Hox2.8) during
hindbrain segmentation. Cell 72, 183–196.
57. Maconochie, M., Nonchev, S., Studer, M., Chan, S.-K., Pöpperl, H., Sham, M.-H., Mann, R., and Krumlauf,
R. (1997) Cross-regulation in the mouse HoxB complex: the expression of Hoxb2 in rhombomere 4 is
regulated by Hoxb1. Genes Dev. 11, 1885–1896.
58. Manzanares, M., Cordes, S., Kwan, C.-T., Sham, M.-H., Barsh, G., and Krumlauf, R. (1997) Segmental
regulation of Hoxb3 by kreisler. Nature 387, 191–195.
59. Manzanares, M., Cordes, S., Ariza-McNaughton, L., Sadl, V., Maruthainar, K., Barsh, G., and Krumlauf, R.
(1999) Conserved and distinct roles of kreisler in regulation of the paralogous Hoxa3 and Hoxb3 genes.
Development 126, 759–769.
60. Geada, A.M.C., Gaunt, S.J., Azzawi, M., Shimeld, S.M., Pearce, J., and Sharpe, P.T. (1992) Sequence and
embryonic expression of the murine Hox-3.5 gene. Development 116, 497–506.
61. Morrison, A., Ariza-McNaughton, L., Gould, A., Featherstone, M., and Krumlauf, R. (1997) HOXD4 and
regulation of the group 4 paralog genes. Development 124, 3135–3146.
62. Chavrier, P., Zerial, M., Lemaire, P., Almendral, J., Bravo, R., and Charnay, P. (1988) A gene encoding a
protein with zinc fingers is activated during Go/G1 transition in cultured cells. EMBO J. 7, 29–35.
63. Lemaire, P., Revelant, O., Bravo, R., and Charnay, P. (1988) Two mouse genes encoding potential
transcription factors with identical DNA-binding domains are activated by growth factors in cultured cells.
Proc. Natl. Acad. Sci. U. S. A. 85, 4691–4695.
64. Nardelli, J., Gibson, T., Vesque, C., and Charnay, P. (1991) Base sequence discrimination by zinc-finger
DNA-binding domains. Nature 349, 175–178.
65. Deol, M.S. (1964) The origin of the abnormalities of the inner ear in dreher mice. J. Embryol. Exp. Morphol.
12, 727–733.
66. Cordes, S.P. and Barsh, G.S. (1994) The mouse segmentation gene kr encodes a novel basic domain-leucine
zipper transcription factor. Cell 79, 1025–1034.
67. Conlon, R.A. and Rossant, J. (1992) Exogenous retinoic acid rapidly induces anterior ectopic expression of
murine Hox-2 genes in vivo. Development 116, 357–368.
68. Conlon, R.A. (1995) Retinoic acid and pattern formation in vertebrates. TIG 11, 314–319.
69. Morriss, G.M. and Thorogood, P.V. (1978) An approach to cranial neural crest migration and differentiation
in mammalian embryos. In Development in Mammals. Johnson, M.H., Ed. Elsevier/North-Holland,
Amsterdam. pp. 363–411.
70. Lammer, E., Chen, D., Hoar, R., Agnish, A., Benke, P., Braun, J., Curry, C., Fernhoff, P., Grix, A., Lott, I.,
et al. (1985) Retinoic acid embryopathy. N. Engl. J. Med. 313, 837–841.
71. Durston, A., Timmermans, J., Hage, W., Hendriks, H., de Vries, N., Heideveld, M., and Nieuwkoop, P.
(1989) Retinoic acid causes an anteroposterior transformation in the developing central nervous system.
Nature 340, 140–144.
72. Marshall, H., Nonchev, S., Sham, M.H., Muchamore, I., Lumsden, A., and Krumlauf, R. (1992) Retinoic
acid alters hindbrain Hox code and induces transformation of rhombomeres 2/3 into a 4/5 identity. Nature
360, 737–741.
73. Mangelsdorf, D.J., Thummel, C., Beato, M., Herrlich, P., Schütz, G., Umesono, K., Blumberg, B., Kastner, P., Mark,
M., Chambon, P., et al. (1995) The nuclear receptor superfamily: the second decade. Cell 83, 835–839.
74. Dencker, L., Annerwall, E., Busch, C., and Eriksson, U. (1990) Localization of specific retinoid-binding sites and
expression of cellular retinoic-acid-binding protein (CRABP) in the early mouse embryo. Development 110, 343–352.
75. Maden, M., Horton, C., Graham, A., Leonard, L., Pizzey, J., Siegenthaler, G., Lumsden, A., and Eriksson, U. (1992)
Domains of cellular retinoic acid-binding protein I (CRABP I) expression in the hindbrain and neural crest of the
mouse embryo. Mech. Dev. 37, 13–23.
259

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
76. Ruberte, E., Friederich, V., Morriss-Kay, G., and Chambon, P. (1992) Differential distribution patterns of
CRABP-I and CRABP-II transcripts during mouse embryogenesis. Development 115, 973–989.
77. Lyn, S. and Giguere, V. (1994) Localisation of CRABP-I and CRABP-II mRNA in the early mouse embryo
by whole-mount in situ hybridisation: implications for teratogenesis and neural development. Dev. Dynam.
199, 280–291.
78. Mangelsdorf, D.J., Borgmeyer, U., Heyman, R.A., Yang Zhou, J., Ong, E.S., Oro, A.E., Kakizuka, A., and
Evans, R.M. (1992) Characterization of three RXR genes that mediate the action of 9-cis retinoic acid. Genes
Dev. 6, 329–344.
79. Mangelsdorf, D.J. and Evans, R.M. (1995) The RXR heterodimers and ophan receptors. Cell 83, 841–850.
80. Mendelsohn, C., Larkin, S., Mark, M., LeMeur, M., Clifford, J., Zelent, A., and Chambon, P. (1994)
RAR·isoforms: distinct transcriptional control by retinoic acid and specific spatial patterns of promoter
activity during mouse embryonic development. Mech. Dev. 45, 227–241.
81. Mendelsohn, C., Ruberte, E., Le Meur, M., Morriss-Kay, G., and Chambon, P. (1991) Developmental
analysis of the retinoic acid-inducible RAR-beta 2 promoter in transgenic animals. Development 113, 723–
734.
82. Dollé, P., Ruberte, E., Leroy, P., Morriss-Kay, G., and Chambon, P. (1990) Retinoic acid receptors and
cellular retinoid binding proteins. I. A systematic study of their differential pattern of transcription during
mouse organogenesis. Development 110, 1133–1151.
83. Dollé, P., Fraulob, V., Kastner, P., and Chambon, P. (1994) Developmental expression of murine retinoid X
receptor (RXR) genes. Mech. Dev. 45, 91–104.
84. Simeone, A., Acampora, D., Arcioni, L., Andrews, P.W., Boncinelli, E., and Mavilio, F. (1990) Sequential
activation of HOX2 homeobox genes by retinoic acid in human embryonal carcinoma cells. Nature 346,
763–766.
85. Simeone, A., Acampora, D., Nigro, V., Faiella, A., D'Esposito, M., Stornaiuolo, A., Mavilio, F., and
Boncinelli, E. (1991) Differential regulation by retinoic acid of the homeobox genes of the four HOX loci in
human embryonal carcinoma cells. Mech. Dev. 33, 215–227.
86. Papalopulu, N., Hunt, P., Wilkinson, D., Graham, A., and Krumlauf, R. (1990) Hox-2 homeobox genes and
retinoic acid: potential roles in patterning the vertebrate nervous system. In Advances in Neural Regeneration
Research. Seil, F.J., Ed. Wiley-Liss, New York. pp. 291–307.
87. Papalopulu, N., Clarke, J., Bradley, L., Wilkinson, D., Krumlauf, R., and Holder, N. (1991) Retinoic acid
causes abnormal development and segmental patterning of the anterior hindbrain in Xenopus embryos.
Development 113, 1145–1159.
88. Moroni, M., Vigano, M., and Mavilio, F. (1993) Regulation of the human HOXD4 gene by retinoids. Mech.
Dev. 44, 139–154.
89. Hill, J., Clarke, J.D.W., Vargesson, N., Jowett, T., and Holder, N. (1995) Exogenous retinoic acid causes
specific alterations in the development of the midbrain and hindbrain of the zebrafish embryo including
positional respecification of the Mauthner neuron. Mech. Dev. 50, 3–16.
90. Papalopulu, N. and Kintner, C. (1996) A posteriorising factor, retinoic acid, reveals that anteroposterior
patterning controls the timing of neuronal differentiation in Xenopus neuroectoderm. Development 122,
3409–3418.
91. Morrison, A., Chaudhuri, C., Ariza-McNaughton, L., Muchamore, I., Kuroiwa, A., and Krumlauf, R. (1995)
Comparative analysis of chicken Hoxb-4 regulation in transgenic mice. Mech. Dev. 53, 47–59.
92. Morrison, A., Moroni, M., Ariza-McNaughton, L., Krumlauf, R., and Mavilio, F. (1996) In vitro and
transgenic analysis of a human HOXD4 retinoid-responsive enhancer. Development 122, 1895–1907.
93. Kolm, P. and Sive, H. (1995) Regulation of the Xenopus labial homeodomain genes, HoxA1 and
HoxD1: activation by retinoids and peptide growth factors. Dev. Biol. 167, 34–49.
94. Kolm, P., Apekin, V., and Sive, H. (1997) Xenopus hindbrain patterning requires retinoid signaling. Dev.
Biol. 192, 1–16.
95. Blumberg, B., Bolado, J., Moreno, T., Kintner, C., Evans, R., and Papalopulu, N. (1997) An essential role for
retinoid signaling in anteroposterior neural patterning. Development 124, 373–379.
260

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
96. Sharpe, C. and Goldstone, K. (1997) Retinoid receptors promote primary neurogenesis in Xenopus.
Development 124, 515–523.
97. Langston, A.W. and Gudas, L.J. (1992) Identification of a retinoic acid responsive enhancer 3' of the murine
homeobox gene Hox-1.6. Mech. Dev. 38, 217–228.
98. Pöpperl, H. and Featherstone, M. (1993) Identification of a retinoic acid repsonse element upstream of the
murine Hox-4.2 gene. Mol. Cell. Biol. 13, 257–265.
99. Marshall, H., Studer, M., Pöpperl, H., Aparicio, S., Kuroiwa, A., and Brenner, S., Krumlauf, R. (1994) A
conserved retinoic acid response element required for early expression of the homeobox gene Hoxb-1.
Nature 370, 567–571.
100. Studer, M., Pöpperl, H., Marshall, H., Kuroiwa, A., and Krumlauf, R. (1994) Role of a conserved retinoic
acid response element in rhombomere restriction of Hoxb-1. Science 265, 1728–1732.
101. Gould, A., Itasaki, N., and Krumlauf, R. (1998) Initiation of rhombomeric Hoxb4 expression requires
induction by somites and a retinoid pathway. Neuron 21, 39–51.
102. Huang, D., Chen, S., langston, A., and Gudas, L. (1998) A conserved retinoic acid responsive element in the
murine Hoxb-1 gene is required for expression in the developing gut. Development 125, 3235–3246.
103. Packer, A., Crotty, D., Elwell, V., and Wolgemuth, D. (1998) Expression of the murine Hoxa4 gene requires
both autoregulation and a conserved retinoic acid response element. Development 125, 1991–1998.
104. Moallem, S.A. and Hales, B.F. (1995) Induction of apoptosis and cathepsin D in limbs exposed in vitro to an
activated analog of cyclophosphamide. Teratology 52, 3–14.
105. Pöpperl, H., Bienz, M., Studer, M., Chan, S., Aparicio, S., Brenner, S., Mann, R., and Krumlauf, R. (1995)
Segmental expression of Hoxb1 is controlled by a highly conserved autoregulatory loop dependent upon
exd/Pbx. Cell 81, 1031–1042.
106. Gould, A., Morrison, A., Sproat, G., White, R., and Krumlauf, R. (1997) Positive cross-regulation and
enhancer sharing: two mechanisms for specifying overlapping Hox expression patterns. Genes Dev 11, 900–
913.
107. Zhang, M., Kim, H.-J., Marshall, H., Gendron-Maguire, M., Lucas, A.D., Baron, A., Gudas, L.J., Gridley, T.,
Krumlauf, R., and Grippo, J.F. (1994) Ectopic Hoxa-1 induces rhombomere transformation in mouse
hindbrain. Development 120, 2431–2442.
108. Alexandre, D., Clarke, J., Oxtoby, E., Yan, Y.-L., Jowett, T., and Holder, N. (1996) Ectopic expression of
Hoxa-1 in the zebrafish alters the fate of the mandibular arch neural crest and phenocopies a retinoic acid -
induced phenotype. Development 122, 735–746.
109. Bell, E., Wingate, R., and Lumsden, A. (1999) Homeotic transformation of rhombomere identity after
localized Hoxb1 misexpression. Science 284, 2168–2171.
110. Gavalas, A., Studer, M., Lumsden, A., Rijli, F., Krumlauf, R., and Chambon, P. (1998) Hoxa1 and Hoxb1
synergize in patterning the hindbrain, cranial nerves and second pharyngeal arch. Development 125, 1123–
1136.
111. Studer, M., Gavalas, A., Marshall, H., Ariza-McNaughton, L., Rijli, F., Chambon, P., and Krumlauf, R. (1998)
Genetic interaction between Hoxa1 and Hoxb1 reveal new roles in regulation of early hindbrain patterning.
Development 125, 1025–1036.
112. Prince, V. and Lumsden, A. (1994) Hoxa-2 expression in normal and transposed rhombomeres: independent
regulation in the neural tube and neural crest. Development 120, 911–923.
113. Rijli, F.M., Mark, M., Lakkaraju, S., Dierich, A., Dolle, P., and 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,
1333–1349.
114. Bailey, W.J., Kim, J., Wagner, G.P., and Ruddle, F.H. (1997) Phylogenetic reconstruction of vertebrate Hox cluster
duplications. Mol. Biol. Evol. 14, 843–853.
115. Davenne, M., Maconochie, M., Neun, R., Brunet, J.-F., Chambon, P., Krumlauf, R., and Rijli, F. (1999) Hoxa2 and
Hoxb2 control dorsoventral patterns of neuronal development in the rostral hindbrain. Neuron 22, 677–691.
116. Chisaka, O. and Capecchi, M. (1991) Regionally restricted developmental defects resulting from targeted disruption of
the mouse homeobox gene hox1.5. Nature 350, 473–479.
261

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
117. Manley, N. and Capecchi, M. (1997) Hox group 3 paralogous genes act synergistically in the formation of
somitic and neural crest-derived structures. Dev. Biol. 192, 274–288.
118. Condie, B. and Capecchi, M. (1993) Mice homozygous for a targeted disruption of Hoxd-3(Hox-4.1) exhibit anterior
transformations of the first and second cervical vertebrae, the atlas and axis. Development 119, 579–595.
119. Condie, B.G. and Capecchi, M.R. (1994) Mice with targeted disruptions in the paralogous genes Hoxa-3 and
Hoxd-3 reveal synergistic interactions. Nature 370, 304–307.
120. Manley, N. and Capecchi, M. (1998) Hox group 3 paralogs regulate the development and migration of the
thymus, thyroid and parathyroid glands. Dev. Biol. 195, 1–15.
121. Schneider-Maunoury, S., Seitanidou, T., Charnay, P., and Lumsden, A. (1997) Segmental and neuronal
architecture of the hindbrain of Krox-20 mouse mutants. Development 124, 1215–1226.
122. Frohman, M.A., Martin, G.R., Cordes, S., Halamek, L.P., and Barsh, G.S. (1993) Altered rhombomere-
specific gene expression and hyoid bone differentiation in the mouse segmentation mutant kreisler (kr).
Development 117, 925–936.
123. Theil, T., Ariza-McNaughton, L., Manzanares, M., Krumlauf, R., and Wilkinson, D. (1999) kreisler
regulates rostrocaudal identity in the hindbrain. Development, in revision.
124. Selleck, M.A. and Bronner-Fraser, M. (1995) Origins of the avian neural crest: the role of neural plate-
epidermal interactions. Development 121, 525–538.
125. Liem, K.F., Jr., Tremml, G., Roelink, H., and Jessell, T.M. (1995) Dorsal differentiation of neural plate cells
induced by BMP-mediated signals from epidermal ectoderm. Cell 82, 969–979.
126. Le Douarin, N.M., Ziller, C., and Couly, G.F. (1993) Patterning of neural crest derivatives in the avian
embryo: in vivo and in vitro studies. Dev. Biol. 159, 24–49.
127. Le Douarin, N. and Kalcheim, C. (1999) The Neural Crest. 2
nd
ed. Bard, J., Barlow, P., and Kirk, D., Eds.
Cambridge Univesity Press, U.K.
128. Noden, D. (1978) The control of avian cephalic neural crest cytodifferentiation. I. Skeletal and connective
tissues. Dev. Biol. 67, 296–312.
129. Noden, D. (1978) The control of avian cephalic neural crest cytodifferentiation. II. Neural tissues. Dev. Biol.
67, 313–329.
130. Noden, D. (1988) Interactions and fates of avian craniofacial mesenchyme. Development 103, 121–140.
131. Noden, D.M. (1986) Origins and patterning of craniofacial mesenchymal tissues. J. Craniofacial Genet. Dev.
Biol. (Suppl. 2), 15–31.
132. Andres, G. (1946) Uber Induction und Entwicklung von Kopforganen aus Unkenektoderm im Molch
(Epidermis, Plakoden and Derivate der Nemalleiste). Rev. Suisse Zool. 53, 502–510.
133. Andres, G. (1949) Untersuchungen an chimaren von Triton und Bombinator. Genetics 24, 387–534.
134. Wagner, G. (1949) Die Bedeutung der Neualleiste fur die Kpfgestaltung der Amphibienlarven. Rev. Suisse
Zool. 56, 519–620.
135. Wagner, G. (1959) Untersuchungen an Bombinator-Triton-Chimareren. Rouux Arch. Entwicklungsmech.
151, 136–158.
136. Noden, D. (1983) The role of the neural crest in patterning of avian cranial skeletal, connective, and muscle
tissues. Dev. Biol. 96, 144–165.
137. Couly, G. and LeDouarin, N. (1988) The fate map of cephalic neural primordium at the presomitic to the 3-
somtie stage in the avian embryo. Development 103(Suppl.), 101–113.
138. Saldivar, J., Krull, C., Krumlauf, R., Ariza-McNaughton, L., and Bronner-Fraser, M. (1996) Rhombomere of
origin determines autonomous versus environmentally regulated expression of Hoxa3 in the avian embryo.
Development 122, 895–904.
139. Hunt, P., Ferretti, P., Krumlauf, R., and Thorogood, P. (1995) Restoration of normal Hox code and branchial
arch morphogenesis after extensive deletion of hindbrain neural crest. Dev. Biol. 168, 584–597.
140. Hunt, P., Clarke, J.D.W., Buxton, P., Ferretti, P., and Thorogood, P. (1998) Stability and plasticity of neural
crest patterning and branchial arch Hox code after extensive cephalic crest rotation. Dev. Biol. 198, 82–104.
262

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
141. Grapin-Botton, A., Bonnin, M.-A., Ariza-McNaughton, L., Krumlauf, R., and LeDouarin, N.M. (1995)
Plasticity of transposed rhombomeres: Hox gene induction is correlated with phenotypic modifications.
Development 121, 2707–2721.
142. Couly, G., Grapin-Botton, A., Coltey, P., Ruhin, B., and 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 128, 3445–3459.
143. Trainor, P. and Krumlauf, R. (2000) Patterning the cranial neural crest: Hindbrain segmentation and Hox
gene pasticity. Nat. Rev. Neurosci. 1, 116–124.
144. Trainor, P. and Krumlauf, R. (2000) Plasticity in mouse neural crest cells reveals a new patterning role for
cranial mesoderm. Nat. Cell Biol. 2, 96–102.
145. Schilling, T. (2001) Plasticity of zebrafish Hox expression in the hindbrain and cranial neural crest
hindbrain. Dev. Biol. 231, 201–216.
146. Maconochie, M., Krishnamurthy, R., Nonchev, S., Meier, P., Manzanares, M., Mitchell, P., and Krumlauf,
R. (1999) Regulation of Hoxa2 in cranial neural crest cells involves members of the AP-2 family.
Development 126, 1483–1494.
147. Frasch, M., Chen, X., and Lufkin, T. (1995) Evolutionary-conserved enhancers direct region-specific expression of the
murine Hoxa-1 and Hoxa-2 loci in both mice and Drosophila. Development 121, 957–974.
148. Trainor, P.A. and Krumlauf, R. (2001) Hox genes, neural crest cells and branchial arch patterning. Curr. Opin. Cell
Biol. 13, 698–705.
149. Veitch, E., Begbie, J., Schilling, T.F., Smith, M.M., and Graham, A. (1999) Pharyngeal arch patterning in the absence
of neural crest. Curr. Biol. 9, 1481–1484.
150. Schaeffer, B. (1987) Deutoerstome monophyly and phylogeny. Evol. Biol. 21, 179–234.
151. Holland, P.W.H. and Garcia-Fernandez, J. (1996) Hox genes and chordate evolution. Dev. Biol. 173, 382–
395.
152. Noden, D.M. (1982) Patterns and organization of craniofacial skeletogenic and myogenic mesenchyme: a
perspective. Prog. Clin. Biol. Res. 101, 167–203.
153. Noden, D. (1987) Interactions between cephalic neural crest and mesodermal populations. In Developmental
and Evolutionary Aspects of the Neural Crest. Maderson, P.F.A., Ed. John Wiley & Sons, New York. pp.
89–119.
154. Trainor, P.A., Tan, S.S., and Tam, P.P.L. (1994) Cranial paraxial mesoderm-regionalization of cell fate and
impact on craniofacila development in mouse embryos. Development 120, 2925–2932.
155. Couly, G. and Le Douarin, N. (1990) Head morphogenesis in embryonic avian chimeras: evidence for a
segmental pattern in the ectoderm corresponding to the neuromeres. Development 108, 543–558.
156. Lumsden, A.G. (1988) Spatial organization of the epithelium and the role of neural crest cells in the
initiation of the mammalian tooth germ. Development 103(Suppl.), 155–169.
157. Tucker, A.S. and Sharpe, P.T. (1999) Molecular genetics of tooth morphogenesis and patterning: the right
shape in the right place. J. Dent. Res. 78, 826–834.
158. Tucker, A.S., Yamada, G., Grigoriou, M., Pachnis, V., and Sharpe, P.T. (1999) Fgf-8 determines rostral-
caudal polarity in the first branchial arch. Development 126, 51–61.
159. Trumpp, A., Depew, M.J., Rubenstein, J.L., Bishop, J.M., and Martin, G.R. (1999) Cre-mediated gene
inactivation demonstrates that FGF8 is required for cell survival and patterning of the first branchial arch.
Genes Dev. 13, 3136–3148.
160. Baker, C.V. and Bronner-Fraser, M. (2001) Vertebrate cranial placode. I. Embryonic induction. Dev. Biol.
232, 1–-61.
161. Begbie, J., Brunet, J.F., Rubenstein, J.L., and Graham, A. (1999) Induction of the epibranchial placodes.
Development 126, 895–902.
162. Epperlein, H.H. (1974) The ectomesenchymal-endodermal interaction-system (EEIS) of Triturus alpestris in
tissue culture. I. Observations on attachment, migration and differentiation of neural crest cells.
Differentiation 2, 151–168.
263

Trainor: Craniofacial Development TheScientificWorldJOURNAL (2003) 3, 240-264
163. Piotrowski, T. and Nusslein-Volhard, C. (2000) The endoderm plays an important role in patterning the
segmented pharyngeal region in zebrafish (Danio rerio). Dev. Biol. 225, 339–356.
164. Couly, G., Creuzet, S., Bennaceur, S., Vincent, C., and 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, 1061–1073.
165. Lee, S.H., Fu, K.K., Hui, J.N., and Richman, J.M. (2001) Noggin and retinoic acid transform the identity of
avian facial prominences. Nature 414, 909–912.
166. Hu, D. and Helms, J.A. (1999) The role of sonic hedgehog in normal and abnormal craniofacial
morphogenesis. Development 126, 4873–4884.
167. Barlow, A.J. and Francis-West, P.H. (1997) Ectopic application of recombinant BMP-2 and BMP-4 can
change patterning of developing chick facial primordia. Development 124, 391–398.
168. Irving, C. and Mason, I. (2000) Signalling by fgf8 from the isthmus patterns the anterior hindbrain and
establishes the anterior limit of Hox gene expression. Development 127, 177–186.
169. Trainor, P.A., Ariza-McNaughton, L., and Krumlauf, R. (2002) Role of the isthmus and FGFs in resolving
the paradox of neural crest plasticity and prepatterning. Science 295, 1288–1291.
170. Kanzler, B., Kuschert, S.J., Liu, Y.-H., and Mallo, M. (1998) Hoxa2 restricts the chondrogenic domain and
inhibits bone formation during development of the branchial area. Development 125, 2587–2597.
171. Grammatopoulos, G.A., Bell, E., Toole, L., Lumsden, A., and Tucker, A.S. (2000) Homeotic transformation
of branchial arch identity after Hoxa2 overexpression. Development 127, 5355–5365.
172. Pasqualetti, M., Ori, M., Nardi, I., and Rijli, F.M. (2000) Ectopic Hoxa2 induction after neural crest
migration results in homeosis of jaw elements in Xenopus. Development 127, 5367–5378.
173. Gans, C. and Northcutt, R. (1983) Neural crest and the origin of vertebrates: a new head. Science 220, 268–
274.
This article should be referenced as follows:
Trainor, Paul A. (2003) Making headway: The roles of Hox genes and neural crest cells in craniofacial development.
TheScientificWorldJOURNAL 3, 240–264.
Handling Editor:
Roger Keynes, Principal Editor for Embryology — a domain of TheScientificWorldJOURNAL.
264