1 Department of Biology, Emory University, Atlanta, GA 30322, USA
2 Laboratory of Molecular Genetics, National Institute of Child Health and Human
Development, National Institutes of Health, Bethesda, MD 20892, USA
Present address: Department of Anatomy and Developmental Biology, University
College London, Gower Street, London WC1E 6BT, UK
Author for correspondence (e-mail:
afrtiz{at}biology.emory.edu)
Accepted 18 November 2002
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SUMMARY |
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Key words: Foxi1, Forkhead, Zebrafish, Otic Placode, Ear, Jaw
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INTRODUCTION |
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Recent work in chick and zebrafish has identified several of the signaling
molecules that function in the earliest inductive steps. Chick FGF19
and FGF3 are expressed adjacent to the otic anlagen in the paraxial
cephalic mesoderm and later in the hindbrain, and FGF3 is
additionally expressed in the otic placode and vesicle
(Ladher et al., 2000;
Mahmood et al., 1995
).
FGF19 induces Wnt-8c expression in the neural tissue, and
together these two signals can induce expression of otic markers
(Ladher et al., 2000
). In
addition, misexpression of FGF3 induces ectopic otic vesicle
formation (Vendrell et al.,
2000
), and depletion of this molecule in explants of otic
precursor tissue results in failure to form otic vesicles
(Represa et al., 1991
). During
gastrulation, zebrafish fgf3 and fgf8 are expressed in
mesendoderm and the presumptive hindbrain
(Phillips et al., 2001
). Loss
of function of these two partially redundant genes leads to a severe reduction
in otic vesicle size and a loss of pax8 expression in the otic
anlagen (Phillips et al.,
2001
); pax8 is the earliest known marker of otic
precursor cells in both fish and mammals
(Pfeffer et al., 1998
). Thus,
these and other experiments (Kozlowski et
al., 1997
; Woo and Fraser,
1997
) show that induction of the otic placode begins as early as
mid-gastrulation and that Fgf-signaling molecules provide some of the earliest
inductive signals in this process.
Less is known about early acting genes that function within the otic
precursor cells to respond to inductive signals and elicit placode formation.
No otic defect has been reported for a knockout of mouse Pax8
(Mansouri et al., 1998), and
otic function of zebrafish pax8 has not been characterized. Zebrafish
dlx3b and dlx4b (formerly dlx3 and dlx7,
respectively), two members of the Distal-less family of transcription
factors, are expressed shortly after the onset of pax8 expression in
presumptive otic tissue (Akimenko et al.,
1994
; Ekker et al.,
1992
; Ellies et al.,
1997
). Embryos homozygous for a deletion of these genes lack otic
placodes, and morpholino-mediated knockdown of one or both genes demonstrates
their partially redundant function in otic placode development
(Solomon and Fritz, 2002
). In
mouse, targeted disruption of the Drosophila eyes absent homologue
Eya1 leads to a severe defect, as development arrests at the otic
vesicle stage and the otic vesicle regresses by apoptosis; however, early
placode induction occurs normally in these embryos
(Xu et al., 1999
).
We have identified a mutation in zebrafish, hearsay
(hsy), which disrupts the foxi1 gene and results in a
reduction or loss of the otic placode and vesicle. Foxi1 is a member of the
forkhead family of winged-helix transcription factors that share a highly
conserved, 110 amino acid DNA-binding domain
(Kaufmann and Knochel, 1996).
Founding forkhead family members, Drosophilia Forkhead
(Weigel et al., 1989
) and
mammalian HNF-3
(Lai et al.,
1990
), have been shown to function in the development of terminal
structures in the embryo and in regulation of liver gene expression,
respectively. Currently, more than 100 forkhead genes have been identified in
species ranging from yeast to humans (see the Winged Helix Proteins site;
http://www.biology.pomona.edu/fox.html).
These genes have been implicated in a broad range of biological functions,
including early patterning and morphogenesis
(Pogoda et al., 2000
), cell
specification (Miller et al.,
1993
), gene regulation in differentiated tissues
(Clevidence et al., 1994
), cell
proliferation (Dou et al.,
1999
) and tumorigenesis (Kops
and Burgering, 1999
). So far, few forkhead genes have been
implicated specifically in otic development. Of the more than a dozen forkhead
genes identified in zebrafish (Biggs and
Cavenee, 2001
; Boggetti et al.,
2000
; Dirksen and Jamrich,
1995
; Odenthal and
Nusslein-Volhard, 1998
;
Strahle et al., 1993
;
Topczewska et al., 2001
), only
one, foxg1 (formerly zBF-1), is expressed in the otic
vesicle (Toresson et al.,
1998
), and no otic defect is reported in mice that are homozygous
null for the BF-1 gene (Xuan et
al., 1995
). In mouse, Foxf2 is expressed in the otic
vesicle, as well as other tissues (Aitola
et al., 2000
), but no functional analysis has been reported for
this gene. Foxi1 is expressed in the otic vesicle, and mice
homozygous for a targeted inactivation of this gene have severely malformed
inner ears and exhibit both cochlear and vestibular dysfunction
(Hulander et al., 1998
).
Here we report the functional characterization of the hsy/foxi1 gene in zebrafish. Embryos homozygous for the hsy mutation display a severe reduction or loss of the otic placode, providing the first example of a single gene mutation that can lead to a complete loss of otic placodes. In addition, hsy mutant embryos also exhibit a malformation of the jaw. Expression analysis of foxi1 suggests a direct role for this gene in the development of the otic placode and branchial arch derivatives. We demonstrate that otic expression of pax8 is absent in hsy mutants, and that misexpression of foxi1 can induce ectopic pax8 expression. Together, these data suggest that foxi1 function is required within otic precursor cells for the earliest stages of otic placode induction.
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MATERIALS AND METHODS |
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Cloning and construction of expression plasmid
A partial zebrafish foxi1 clone was identified in an expression
pattern screen using high-throughput in situ hybridization (clone 3015)
(Kudoh et al., 2001). The
5' end of foxi1 was subsequently generated by 5'-RACE
using the SMART RACE Kit (Clontech) with a gene-specific primer
(5'-cgtctctgcttcggaccgcgtttcc-3'), and sequenced. Full-length cDNA
was then amplified and subcloned into the expression vector pCS2+.
Whole-mount in situ hybridization
Whole-mount in situ hybridization was performed as described
(Thisse and Thisse, 1998), and
double labeling as outlined (Itoh et al.,
2002
; Jowett,
2001
). Embryos were prepared for sectioning as described
(Westerfield, 2000
), and 5
µm plastic sections or 15 µm frozen sections were generated. The
zebrafish dlx3b (Ekker et al.,
1992
), dlx4b (Ellies
et al., 1997
), dlx5a
(Akimenko et al., 1994
),
dlx2a (Akimenko et al.,
1994
), pax8 (Pfeffer
et al., 1998
), pax2a
(Krauss et al., 1991
),
krox20 (Oxtoby and Jowett,
1993
), crestin (Luo
et al., 2001
; Rubinstein et
al., 2000
) and otx2
(Li et al., 1994
) probes were
previously described.
Morpholino injections
Morpholino antisense oligonucleotides (Gene Tools, LLC, Corvallis, OR) were
as follows (complementary bases to the predicted start codon are indicated in
italics): foxi1,
5'-TAATCCGCTCTCCCTCCAGAAACAT-3'; Gene Tools, LLC standard
control oligo
(http://www.gene-tools.com/).
Alcian Green staining
Embryos were fixed overnight in 4% paraformaldehyde, followed by several
washes in PBT (phosphate-buffered solution +0.1% Tween 20) the next day. The
embryos were then placed in 30% hydrogen peroxide for approximately 20 minutes
to bleach pigment cells, washed in PBT several more times, then stained in
0.37% HCl/70% ethanol solution/0.1% Alcian Green for 1.5 hours. This was
followed by several brief washes in 1%HCl/70% ethanol. The tissues were
cleared in 50% glycerol/0.5% KOH and the cartilages were photographed.
Genotyping
DNA was prepared after in situ hybridization from embryos sorted by
phenotype, and a 607 bp PCR fragment was amplified with the following primers:
F, 5'-AAACCCCTCAGAGACGAGCACACTCA-3'; R,
5'-CTGCCAGCCGGCTTTACTTTTCTTGT-3'. The amplification products were
digested with TaqI, which only cuts the mutant sequence, and analyzed
on a 1.5% agarose gel.
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RESULTS |
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The hearsay mutation disrupts the foxi1 gene
To identify the gene disrupted by the hsy mutation, DNA was
prepared from individual mutant and wild-type embryos and tested for linkage
to polymorphic microsatellite markers
(http://zebrafish.mgh.harvard.edu)
(Shimoda et al., 1999). The
mutation was localized to LG12, between markers Z14982 (two recombinants/265
chromosomes tested) and Z8344 (two recombinants/253 chromosomes tested, data
not shown).
One candidate gene in this region, 3015, was identified as part of
a screen for expression patterns of embryonic cDNAs
(Kudoh et al., 2001). Clone
3015 was localized by radiation hybrid (RH) mapping (LN54 panel)
(Hukriede et al., 2001
;
Hukriede et al., 1999
) on LG12
in the vicinity of the hsy map position; we will provide evidence
below that the hsy locus encodes the 3015 sequence. A BLAST
search of the NCBI database with the 3015 sequence identified it as
encoding a forkhead domain-containing protein. This sequence was classified in
the Fox I class and consequently named foxi1 (see the Winged Helix
Proteins site;
http://www.biology.pomona.edu/fox.html)
(Fig. 2A). The zebrafish Foxi1
protein shares 52% identity with Xenopus FoxI1c and 40% with human
FOXI1; the forkhead domains are 95% and 94% identical, respectively.
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To test whether foxi1 is disrupted by the hsy mutation, we amplified and sequenced fragments representing the gene from individual haploid mutant and wild-type embryos. Mutant embryo DNA contained a two base-pair deletion close to the amino-terminus of the protein, creating a shift in the reading frame in codon 41 (Fig. 2B). The shifted sequence continues for an additional 117 amino acids with no similarity to known proteins. Thus, the mutation eliminates most of the wild-type Foxi1 protein including the forkhead domain, and most likely represents a null mutation. No additional differences were found between mutant and wild-type sequences. The two basepair deletion in hsy creates a TaqI restriction site, and we used this polymorphism to test for linkage of the mutant phenotype to this sequence change. Genomic DNA was isolated from individual embryos generated from crosses of hsy heterozygous parents, amplified and tested for the TaqI polymorphism: no recombinants between the hsy mutation and the TaqI polymorphism were found in more than 450 meioses tested. Conversely, no homozygotes for the TaqI polymorphism were found in 700 wild-type siblings. Thus, the hsy mutation is tightly linked to the two-base deletion, providing direct genetic evidence for the conclusion that hsy disrupts foxi1.
To test whether inhibition of foxi1 translation could phenocopy
the hsy mutation, we injected wild-type embryos with a morpholino
antisense oligonucleotide (Nasevicius and
Ekker, 2000) against foxi1 mRNA. Injection of this
morpholino produced a phenotype almost identical to that of hsy; otic
vesicles of injected embryos were very small or absent
(Fig. 1D), and jaws were
reduced in size (Fig. 1H). These injected embryos displayed variability in severity of the otic defect,
similar to the variability seen in the mutants. None of these phenotypes were
observed in embryos injected with a control morpholino
(Fig. 1C,G). This observation
confirms the conclusion, based on our linkage and sequencing data, that the
hsy phenotype is caused by disruption of the foxi1 gene.
foxi1 expression
The expression profile of foxi1 is shown in
Fig. 3. At 50% epiboly, the
earliest stage examined, foxi1 is expressed in a portion of the
ectoderm (Fig. 3A), and this
domain of expression becomes broader and stronger by 75% epiboly
(Fig. 3B). To determine whether
foxi1 is expressed in neural or nonneural ectoderm, double in situ
labeling was performed with otx2, a marker of neural ectoderm
(Li et al., 1994).
foxi1 expression does not overlap with otx2, indicating that
it is expressed in non-neural ectoderm
(Fig. 3C).
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By the beginning of somitogenesis, foxi1 expression has become confined to two domains that lie laterally to the neural plate in the dorsal ectoderm (Fig. 3D-F), whereas expression is no longer detectable in ventral ectoderm. Based on double in situ labeling with otx2, the anterior-most region of foxi1 expression lies just posterior to the midbrain-hindbrain boundary (Fig. 3D,F). At the three-somite stage, the two domains of foxi1 expression have become more compact, but are still located in approximately the same position lateral to the hindbrain (Fig. 3G,H). A transverse section of an embryo at this stage shows that the more lateral region of foxi1 expression is confined to the upper, ectodermal layer of cells, whereas more medially, ectodermal and mesendodermal expression can be seen (Fig. 3I).
To determine whether foxi1 is expressed in otic precursor cells, double labeling was performed with both pax8 and dlx3b at the one-somite stage stage. The foxi1 expression domain overlaps pax8 otic expression at this stage, sharing approximately the same posterior and medial boundaries, but extending further than pax8 in the anterior and lateral directions (Fig. 3J,K). foxi1 expression also overlaps with dlx3b at the one-somite stage, although dlx3b expression is more extensive along the a-p axis. Within the region of overlap, foxi1 and dlx3b share the same medial border, but foxi1 extends further laterally (Fig. 3L). These results indicate that foxi1 is expressed in otic precursor cells at late gastrula and early somitogenesis stages. At mid- and late-somitogenesis stages, foxi1 expression is no longer detectable in the otic placode and vesicle, respectively (Fig. 4A-C and data not shown).
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At the 18-somite stage, foxi1 is expressed in throughout the
pharyngeal arches (Fig. 4A,B),
and expression in this region remains detectable at 28 hpf
(Fig. 4C), the latest stage
examined. Sections through the pharyngeal arches of 28 hpf embryos reveal that
foxi1 expression appears to extend into the pouches that separate the
individual arches (Fig. 4D,E).
dlx2a (formerly dlx2) is expressed in cranial neural crest
cells that migrate to reside within the pharyngeal arch primordia by 16 hpf,
and expression in the arches persists through the beginning of the second day
of development (Akimenko et al.,
1994). Sections through 26 hpf embryos double-labeled for
expression of dlx2a (red) and foxi1 (purple) show that most
of the dlx2a-expressing presumptive neural crest cells do not express
foxi1 (Fig. 4F,G).
Rather, foxi1 expression partially surrounds these cells, consistent
with the position of the pouches and/or clefts that separate the neural
crest-containing arches. It is unclear whether any of the
foxi1-expressing cells also express dlx2a, as the purple
substrate masks the expression of the red. Thus, foxi1 is expressed
predominantly in cells other than neural crest in the pharyngeal arches.
foxi1 function in otic placode specification
To study the timing of foxi1 function in otic development, we
examined the expression of several early otic markers in hsy mutant
embryos. pax8 expression is detectable in the presumptive otic domain
of wild-type embryos as early as 90% epiboly, whereas pax2a is
expressed in otic cell precursors beginning in early somitogenesis, several
hours before otic placodes become visible
(Krauss et al., 1991;
Pfeffer et al., 1998
). We find
that both pax8 and pax2a expression is completely absent in
the otic domain of approximately one quarter of three- to five-somite stage
embryos from a cross between hsy heterozygotes
(Fig. 5A-D); these embryos are
presumed to represent hsy homozygotes. In the embryos in which otic
expression was absent, pax8 and pax2a expression in the
pronephros and pax2a expression in the midbrain-hindbrain boundary
was comparable to wild type, providing a control for the in situ reaction.
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dlx3b expression was also examined in three-somite stage embryos
from a cross between hsy heterozygotes. At this stage, dlx3b
is expressed in a continuous ectodermal stripe around the edge of the neural
plate, and expression is upregulated in the otic and olfactory primordia
(Akimenko et al., 1994;
Ekker et al., 1992
). We found
that one-quarter of the embryos from this cross lack this upregulation in the
otic region (Fig. 5E,F). To
confirm that these embryos were mutants, we sorted them based on this
phenotype and individually genotyped them for the hsy mutation
(Fig. 5G). One-hundred per cent
(9/9) of the embryos that lacked dlx3b otic upregulation were
homozygous for the hsy mutation, and 100% (21/21) of those that had
been judged as having normal expression were heterozygous or homozygous for
the wild-type allele.
foxi1 induces ectopic pax8 expression
To learn more about the function of foxi1, we injected an
expression construct containing the full-length coding sequence under the
control of the cyto megalo virus (CMV) promoter into one-cell stage wild-type
embryos. pax8 expression was examined in injected embryos fixed at
three- to five-somite stage. We found, in addition to the normal domains of
otic and pronephric expression, pax8 was also expressed in many
ectopic locations in the injected embryos
(Fig. 6A). This ectopic
expression was observed in 15/18 embryos injected with the expression
construct, and in 16/18 embryos injected with in vitro transcribed
foxi1 RNA (not shown). Double in situ labeling with foxi1
reveals that the ectopic domains of pax8 expression coincide with
ectopic foxi1 expression (Fig.
6A,B). This co-localization of ectopic expression was observed in
11/16 embryos injected with the foxi1 expression construct. Ectopic
expression of dlx3b, dlx5a (formerly dlx4) and
dlx4b, but not pax2a, was also observed in these embryos
(data not shown).
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Expression of late otic markers in hearsay mutants
We examined the effect of the hsy mutation on later stages of otic
development by in situ hybridization with dlx3b, dlx4b and
dlx5a. These three genes are expressed in several developing cranial
structures during mid-late somitogenesis, including the otic vesicle and the
visceral arches (Akimenko et al.,
1994; Ekker et al.,
1992
; Ellies et al.,
1997
). At 16-somite stage, dlx4b and dlx3b are
expressed in the olfactory placode and the otic vesicle in wild-type embryos
(Fig. 7A; dlx3b not
shown), and this otic expression was reduced or absent in all hsy
mutants examined (Fig. 7B;
dlx3b not shown). In 24 hpf wild-type embryos, dlx3b, dlx5a
and dlx4b are expressed in the olfactory placode, visceral arches and
otic vesicle, and dlx5a is additionally expressed in several
structures of the forebrain (Fig.
7C,E; dlx4b not shown). hsy mutants display a
variable reduction or absence of otic staining for all of these genes at this
stage (Fig. 7D,F,G;
dlx4b not shown). Although all mutant embryos examined had reduced
otic expression of these genes, the degree of reduction was variable, ranging
from no otic expression to a reduction in size of the otic expression domain.
This observation is in concordance with the variability in the otic phenotype
observed by morphological inspection, as described above. In addition, we
observe a loss of expression in the branchial arches in some of the mutant
embryos, although this phenotype is not as penetrant as the reduction in otic
expression (Fig. 7D for
dlx3b, F and G for dlx5a; dlx4b not shown).
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Effects of hearsay in jaw formation
Alcian Green staining reveals that all of the major cartilaginous elements
of the jaw are present in hsy mutants at 5 d
(Fig. 8A,B). However, these
elements are spaced more compactly in the a-p dimension, and this compression
presumably results in the reduced jaw phenotype observed in the mutant.
Specifically, the first or mandibular arch derivatives, which include the
mandibular and palatoquadrate elements, appear normal in hsy mutants.
The second or hyoid arch derivatives appear to be differentially affected:
whereas the basihyal element seems to form normally, the ceratohyal (ch)
element is often in a more perpendicular position with respect to the ap axis.
In addition, the ch element is shorter and thicker than in wild-type larvae,
and this thickening may in part be caused by a fusion with the first gill
arch. The posterior or gill arch elements are derived from the third through
seventh pharyngeal arches. Like the ch, the gill arches of hsy
mutants often lie in a more perpendicular orientation with respect to the a-p
axis, and they are shorter and more closely spaced.
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The cartilage elements of the jaw are largely derived from cranial neural
crest cells that migrate into the area of the branchial arches. foxi1
is expressed in the region of the branchial arches
(Fig. 4), providing a basis for
its potential function in the formation of this tissue. In addition to the
jaw, cranial neural crest contributes to pigmentation, the developing heart,
the enteric nervous system (ENS) and the sensory portion of cranial ganglia.
We have not been able to detect any defects in pigmentation or heart
morphology, and anti-Hu staining revealed no ENS defects (data not shown).
Cranial ganglia were not examined. To address the possible effect of the
hsy mutation on the migration and differentiation of neural crest
within the branchial arches, we examined expression of crestin, a
marker for neural crest cells (Luo et al.,
2001; Rubinstein et al.,
2000
). All of the major domains of crestin expression
were present in hsy mutants, including the mandibular, hyoid and
vagal cranial crest streams (Fig.
8C,D). However, double in situ labeling with krox20,
which marks rhombomeres 3 and 5 (r3 and r5), reveals that the hyoid and vagal
crestin domains are more closely spaced in mutant embryos
(Fig. 8C,D). In particular, the
vagal crest domain extends past the anterior boundary of the krox20
r5 stripe in hsy mutants, ending in a region adjacent to r4
(Fig. 8D), whereas in wild-type
embryos, the anterior boundary of this expression domain lies laterally to the
middle of the krox20 r5 stripe
(Fig. 8C). This observation may
provide an explanation for the fusion of the ch element (hyoid crest
derivative) and the third gill arch (vagal crest derivative) seen in
hsy mutant larvae.
dlx2a expression in the cranial neural crest also appears to be
affected in hsy mutant embryos at the 18-somite stage
(Akimenko et al., 1994;
Ellies et al., 1997
)
(Fig. 8E,F). First, similar to
crestin, the three domains of expression in the cranial crest are
more closely spaced. Second, we consistently observe an increase in intensity
of dlx2a staining in the mutants. Likewise, the expression of
foxi1 itself is more intense in hsy mutant embryos
(Fig. 8G,H). This conclusion
was supported by genotype analysis. All the embryos derived from a cross
between hsy heterozygotes were hybridized in situ with dlx2a
or foxi1, sorted by `increased' or `normal' expression of the
examined marker, and genotyped for the hsy mutation using the
TaqI restriction polymorphism (see
Fig. 2B). For foxi1
expression at the 18-somite stage, genotyping revealed that 100% (11/11) of
increased expression embryos were homozygous mutant, and 100% (32/32) of
normal expression embryos were heterozygous or homozygous wild type
(Fig. 8I). For dlx2a
at the 18-somite stage, 86% (12/14) of increased expression embryos were
mutant and 100% (32/32) of normal expression embryos were heterozygous or wild
type. Using 24 hpf embryos we found that, for foxi1, 81% (13/16) of
increased embryos were mutant and 91% (51/56) of normal embryos were wild
type, and for dlx2a, 100% (19/19) increased embryos were mutant and
100% (33/33) of normal embryos were wild type. Thus, increased expression of
dlx2a and foxi1 in the branchial arches is a consequence of
the hsy mutation. No increase in expression of foxi1 was
detectable during gastrulation or early somitogenesis (data not shown). This
increased expression of foxi1 in hsy mutant embryos is
suggestive of a function for foxi1 as a negative transcriptional
regulator. However, it remains unclear whether this autoregulation is direct
or indirect and whether the increased expression of foxi1 or
dlx2a is caused by an increase in the number of cells expressing
these genes or elevated expression levels.
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DISCUSSION |
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In this work we have identified a mutation, hearsay, which leads
to a severe reduction or loss of the otic placode in zebrafish. Further, we
show that the hsy phenotype is caused by a disruption of
foxi1, which was identified in a screen for genes that are
differentially expressed during zebrafish embryogenesis
(Kudoh et al., 2001). Double
in situ labeling with known otic markers shows that foxi1 is
expressed very early in otic precursor cells, as early or earlier than
pax8. Loss of pax8, dlx3b and pax2a otic expression
in hsy mutant embryos demonstrates a defect in the initiation of otic
placode induction and suggests that foxi1 is the earliest-acting gene
characterized thus far within the responding otic precursor domain.
Forkhead genes are known to function as transcriptional regulators (for a
review, see Kaufmann and Knochel,
1996). Thus, it is possible that foxi1 regulates the
transcription of genes necessary for development of the otic placode in
response to signals from surrounding tissues. However, we have shown that
foxi1 is also expressed in some mesendodermal cells underlying the
presumptive otic anlagen of the ectodermal layer. Therefore, it is also
possible that foxi1 functions in a non-cell-autonomous manner to
regulate otic development. In support of the first view we have demonstrated
that misexpression of foxi1 can induce ectopic pax8
expression within the same cells. This observation, together with the loss of
pax8 otic expression in hsy, suggests a simple model in
which foxi1 would activate transcription of pax8. However,
although it appears that foxi1 is necessary, it is not sufficient for
pax8 otic expression, because not every cell that expresses
foxi1 also expresses pax8
(Fig. 3). We also find that
pax8 is not expressed in every location where foxi1 is
ectopically expressed (Fig. 6). Therefore it appears that expression of pax8 requires the function of
regulatory factors in addition to foxi1.
Variability of the hearsay phenotype: multiple steps for
otic placode induction
The loss of pax8 expression observed in hsy embryos is a
robust phenotype; expression of this gene was always absent in the presumptive
otic domains of hsy mutant embryos. However, many hsy
embryos develop an otic vesicle, albeit a small and malformed one. Thus, it
appears that there is no absolute requirement for foxi1 or
pax8 to form at least some aspects of an otic vesicle, nor is ectopic
expression of these two genes sufficient to produce one.
Why this variability in otic placode and vesicle formation? One possibility
is that another foxi1-like gene is present in the zebrafish genome.
It has been suggested that ray-finned fish have undergone an additional round
of genome duplication after diverging from the tetrapod lineage, and many gene
families (Hox, Dlx, Msx, Eng) contain additional genes in the zebrafish genome
(Postlethwait et al., 1998).
If a second foxi1 gene exists, it may be able to partially overcome
the loss of the foxi1/hsy gene.
Another possibility arises from observations that embryonic induction of
the otic placode occurs in multiple, overlapping steps
(Baker and Bronner-Fraser,
2001; Torres and Giraldez,
1998
). Recently, Groves and Bronner-Fraser have elegantly combined
transplantation experiments with molecular analysis of otic placode induction
(Groves and Bronner-Fraser,
2000
). In chick, Pax-2 is expressed in the presumptive
otic placode at the four- to five-somite stage, followed by Sox-3 at
the six-somite stage, BMP-7 at the seven-somite stage, and
Notch at the nine- to ten-somite stage, a few hours before otic
placodes become morphologically visible. When anterior, non-otic epiblast was
grafted to the presumptive otic region of progressively older (11- to
21-somite stage) hosts, the grafted tissue formed an epithelial (otic) vesicle
and often expressed Sox-3 and BMP-7. However, older hosts
rarely induced Pax-2 expression, the earliest otic marker used, in
the graft. Similarly, we observe later otic expression of markers such as
dlx3b, dlx5a and dlx4b in the subset of hsy embryos
that form a small otic vesicle, indicating that foxi1 and/or
pax8 are not absolutely required for the expression of these genes.
It has also been shown that signals for otic placode induction emanate from
multiple tissues. For example, in zebrafish, heterotopic transplants of
mesendoderm from early gastrula embryos can induce ectopic otic vesicles
(Woo and Fraser, 1997
).
Mendonsa and Riley have used one eyed pinhead (oep) mutant
and notail (ntl)/oep double mutant zebrafish
embryos to show that genetic removal of mesendodermal cells leads to a severe
delay in otic placode induction (Mendonsa
and Riley, 1999
). This delay is concomitant with a loss of
pax8 induction in the otic primordia
(Phillips et al., 2001
).
Despite these defects, otic vesicles form, although they are variably smaller
than wild type and lack one or both otoliths. Together, these observations
demonstrate that otic vesicle formation can be uncoupled from the expression
of early marker genes, and that a later source of otic vesicle-inducing
signals can partially compensate for the loss of early mesendodermal
signaling. Thus, foxi1, like pax8, might only be transiently
required in early otic primordia, and continued inductive influences can
partially and variably bypass this requirement.
Induction of the otic placode
In zebrafish, fgf3 and fgf8 are expressed in both early
mesendoderm and in the developing hindbrain during the stage when otic placode
induction occurs, but not within the otic precursor tissues. These two genes
play a redundant role in otic induction; inactivation of both fgf3
and fgf8 at the same time blocks otic placode induction and
expression of pax8 in the otic primordia
(Phillips et al., 2001).
Interestingly, some of these embryos still form a small otic vesicle,
reminiscent of hsy.
Evidence from zebrafish and chick suggests that there are other factors in
addition to fgf3/8 involved in otic placode induction. In chick,
FGF19 is co-expressed with FGF3 in paraxial cephalic
mesoderm around the time of otic induction
(Ladher et al., 2000), and
transplants of mesoderm expressing FGF19 to anterior ectoderm can
induce expression of otic markers. Furthermore, FGF19 requires the
synergistic action of Wnt8c, which is induced by FGF19 in
overlying neural tissue. Although fgf19 has not been reported in
zebrafish, fgf3 is expressed early in gastrulation in cephalic
mesoderm, subjacent to the presumptive otic primordia
(Phillips et al., 2001
).
Genetic removal of mesendodermal cells in oep mutant embryos
(Schier et al., 1997
)
abolishes this fgf3 domain and leads to a delay in otic placode
induction (Mendonsa and Riley,
1999
), but inactivation of fgf3 in wild-type embryos does
not produce a similar delay (Phillips et
al., 2001
). This suggests that mesendodermal signals in addition
to fgf3 are disrupted in oep.
foxi1 and jaw formation
In addition to otic abnormalities, hsy embryos also display a
defect in jaw formation. Although all the cartilaginous elements of the jaw
are present, the spacing is more compact, and this compression presumably
leads to a reduction in size of the jaw. The cartilaginous elements arise from
the branchial arches, which are formed through complex interactions between
endoderm, mesoderm and neural crest cells that migrate to the arches from the
hindbrain. In zebrafish, several mutations have been identified that lead to
defects in inner ear and jaw formation
(Malicki et al., 1996;
Piotrowski et al., 1996
;
Whitfield et al., 1996
), but
this association is not well understood. Both defects may be the consequence
of inappropriate hindbrain patterning, an example of which is seen in the
valentino mutant, which disrupts a bZip transcription factor
expressed in the hindbrain (Moens et al.,
1998
). However, morphological analysis and expression of hindbrain
markers krox20 (Fig.
8) and fgf3/8 (not shown) show no indication of hindbrain
defects in hsy, and cranial crest derivatives outside the jaw appear
unaffected.
In an alternative model, the otic placode may provide positional cues for
neural crest cell migration (Malicki et
al., 1996). In particular, the preotic hyoid neural crest and the
postotic vagal crest migrate in close apposition to the otic placode
(Schilling and Kimmel, 1994
),
and misguided migration could be the result of otic placode defects. In
support of this model, we find that the hyoid and postotic neural crest
domains are positioned more closely to each other in hsy mutant
embryos (Fig. 7C-F). However,
we have previously shown that loss of dlx3b and dlx4b
function blocks otic placode formation but has no detectable effect on the
formation of the jaws (Solomon and Fritz,
2002
).
In a third model, foxi1 may play a direct role in the development of the branchial arches. We have shown that foxi1 is expressed within the branchial arches, apparently within the pouches that separate the individual arches. It is possible that foxi1 may regulate the expression of factors that control neural crest migration within the ectoderm. Another possibility is that foxi1 may be required for the proper development of the pouches and that loss of foxi1 function in hsy leads to a failure of the neural crest populations within the individual arches to become fully separated. In support of this, we have shown that expression domains of neural crest markers dlx2a and crestin are more closely spaced in hsy (Fig. 8). Incomplete separation of neural crest among the arches could lead to the fusion and closer spacing of the neural crest-derived jaw cartilages observed in the older mutant embryos (Fig. 8).
In the light of these results, it appears more probable that foxi1
is required for branchial arch formation. In this context, hsy is
reminiscent of the van gogh (vgo) mutation, which disrupts
the segmentation of the pharyngeal pouches, leading to malformation of the
jaw, and displays an inner ear defect, but does not affect hindbrain
patterning (Piotrowski and
Nusslein-Volhard, 2000;
Whitfield et al., 1996
).
Foxi I class genes
Foxi I class genes have been described in humans
(Larsson et al., 1995;
Pierrou et al., 1994
), mouse
(Hulander et al., 1998
;
Overdier et al., 1997
), rat
(Clevidence et al., 1993
) and
Xenopus (Lef et al.,
1994
; Lef et al.,
1996
). However, it is unclear whether zebrafish foxi1 is
orthologous to any one of these genes. The Xenopus FoxI1c
(Lef et al., 1996
),
FoxI1a and FoxI1b genes
(Lef et al., 1994
) share the
highest degree of sequence conservation with the zebrafish gene. The
expression pattern of the two Xenopus pseudoallelic variants
FoxI1a/b (XFD-2, XFD-2') does not suggest functional
similarity to zebrafish foxi1. Of the three Xenopus FoxI
genes, FoxI1c (XFD-10) is most similar to foxi1 in
sequence. However, Xenopus FoxI1c was reported to be expressed in the
neuroectoderm and somites but not in the otic placode, unlike the pattern for
foxi1 we report (Lef et al.,
1996
). A very recent report provides a more detailed description
of Xenopus FoxI1c (Pohl et al.,
2002
), which suggests that this gene is expressed in preplacodal
tissue and the branchial arches, similar to our observations for
foxi1. Thus, it now appears probable that Xenopus FoxI1c
represents the ortholog of zebrafish foxi1.
Functional analysis for a foxi paralogue has only been reported in
mouse (Hulander et al., 1998).
Mice homozygous for a targeted disruption of the Fkh10 locus
(Foxi1 Mouse Genome Informatics) exhibit circling behavior,
poor swimming ability and abnormal reaching response as well as profound
hearing impairment, demonstrating vestibular and cochlear dysfunction. In
addition, these mice display malformations of the inner ear in that a large,
irregular cavity replaces the cochlea and vestibulum. At 9.5 days post-coitus
(dpc), Fkh10 is expressed exclusively in the otic vesicle, the stage
when the otic vesicle first forms by invagination. Mouse Fkh10 is
also expressed in the epithelium of renal distal convoluted tubules at 16.0
dpc (Overdier et al., 1997
),
however no renal dysfunction has been observed in mutant mice
(Hulander et al., 1998
), and
we have not examined expression of zebrafish foxi1 at larval or adult
stages. These observations suggest that Foxi1 genes in fish and mammals play a
role in the development of the inner ear, but clear discrepancies exist
between the mouse and fish Foxi1 genes. Most notably, zebrafish foxi1
is expressed early in the otic primordia, but is no longer detectable in the
otic placodes or otic vesicles, whereas mouse Fkh10 is expressed at a
later stage in the otic vesicle. In addition, Fkh10 mutant mice do
not show any craniofacial abnormalities, and, unlike the fish foxi1
gene, the mouse gene is not expressed in the branchial arches at the otic
vesicle stage. As branchial arch expression is shared by the related zebrafish
foxi1 and Xenopus FoxI1c genes, it is possible that as yet
unexamined members of this subfamily may show expression in the arches during
murine development.
Individuals affected by Treacher Collins Syndrome (TCS; MIM #154500)
display a phenotype that includes inner ear defects and craniofacial
malformations reminiscent of the hsy phenotype. In most cases, this
syndrome is inherited in an autosomal dominant fashion, and the pathogenic
mutation has been shown to lie within the TCOF1 gene, which maps to
5q32 (The Treacher Collins Syndrome
Collaborative Group, 1996;
Edwards et al., 1997
;
Gladwin et al., 1996
;
Wise et al., 1997
). However,
individuals forming a subset of TCS patients appear to inherit the syndrome in
a recessive fashion and do not carry any obvious defects in the TCOF1
gene, even though haplotype analysis suggests that the genetic defect is
linked closely to the TCOF1 locus
(Edwards et al., 1997
). The
human FOXI1 (FKHL10) gene, which encodes a protein with 40%
sequence identity to zebrafish Foxi1, has been localized to chromosome 5q34
(Larsson et al., 1995
).
Northern analysis with 16 adult and five fetal tissues demonstrated
kidney-specific expression, but no otic tissues were analyzed
(Pierrou et al., 1994
). Based
on our data, the nearby FOXI1 gene is an intriguing candidate for
this recessively inherited syndrome, and this should be easily testable.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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---|
Adamska, M., Herbrand, H., Adamski, M., Kruger, M., Braun, T. and Bober, E. (2001). FGFs control the patterning of the inner ear but are not able to induce the full ear program. Mech. Dev. 109,303 -313.[CrossRef][Medline]
Aitola, M., Carlsson, P., Mahlapuu, M., Enerback, S. and Pelto-Huikko, M. (2000). Forkhead transcription factor FoxF2 is expressed in mesodermal tissues involved in epithelio-mesenchymal interactions. Dev. Dyn. 218,136 -149.[CrossRef][Medline]
Akimenko, M. A., Ekker, M., Wegner, J., Lin, W. and Westerfield, M. (1994). Combinatorial expression of three zebrafish genes related to distal-less: part of a homeobox gene code for the head. J. Neurosci. 14,3475 -3486.[Abstract]
Baker, C. V. and Bronner-Fraser, M. (2001). Vertebrate cranial placodes i. embryonic induction. Dev. Biol. 232,1 -61.[CrossRef][Medline]
Biggs, W. H., 3rd and Cavenee, W. K. (2001). Identification and characterization of members of the FKHR (FOX O) subclass of winged-helix transcription factors in the mouse. Mamm. Genome 12,416 -425.[CrossRef][Medline]
Boggetti, B., Argenton, F., Haffter, P., Bianchi, M. E., Cotelli, F. and Beltrame, M. (2000). Cloning and expression pattern of a zebrafish homolog of forkhead activin signal transducer (FAST), a transcription factor mediating Nodal-related signals. Mech. Dev. 99,187 -190.[CrossRef][Medline]
Chisaka, O., Musci, T. S. and Capecchi, M. R. (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.[CrossRef][Medline]
Clevidence, D. E., Overdier, D. G., Tao, W., Qian, X., Pani, L., Lai, E. and Costa, R. H. (1993). Identification of nine tissue-specific transcription factors of the hepatocyte nuclear factor 3/forkhead DNA-binding-domain family. Proc. Natl. Acad. Sci. USA 90,3948 -3952.[Abstract]
Clevidence, D. E., Overdier, D. G., Peterson, R. S., Porcella, A., Ye, H., Paulson, K. E. and Costa, R. H. (1994). Members of the HNF-3/forkhead family of transcription factors exhibit distinct cellular expression patterns in lung and regulate the surfactant protein B promoter. Dev. Biol. 166,195 -209.[CrossRef][Medline]
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.[Medline]
Dirksen, M. L. and Jamrich, M. (1995). Differential expression of fork head genes during early Xenopus and zebrafish development. Dev. Genet. 17,107 -116.[Medline]
Dou, C. L., Li, S. and Lai, E. (1999). Dual
role of brain factor-1 in regulating growth and patterning of the cerebral
hemispheres. Cereb. Cortex
9, 543-550.
Edwards, S. J., Gladwin, A. J. and Dixon, M. J. (1997). The mutational spectrum in Treacher Collins Syndrome reveals a predominance of mutations that create a premature-termination codon. Am. J. Hum. Genet. 60,515 -524.[Medline]
Ekker, M., Akimenko, M. A., Bremiller, R. and Westerfield, M. (1992). Regional expression of three homeobox transcripts in the inner ear of zebrafish embryos. Neuron 9, 27-35.[Medline]
Ellies, D. L., Stock, D. W., Hatch, G., Giroux, G., Weiss, K. M. and Ekker, M. (1997). Relationship between the genomic organization and the overlapping embryonic expression patterns of the zebrafish dlx genes. Genomics 45,580 -590.[CrossRef][Medline]
Epstein, D. J., Vekemans, M. and Gros, P. (1991). Splotch (Sp2H), a mutation affecting development of the mouse neural tube, shows a deletion within the paired homeodomain of Pax-3. Cell 67,767 -774.[Medline]
Gladwin, A. J., Dixon, J., Loftus, S. K., Edwards, S., Wasmuth,
J. J., Hennekam, R. C. and Dixon, M. J. (1996). Treacher
Collins Syndrome may result from insertions, deletions or splicing mutations,
which introduce a termination codon into the gene. Hum. Mol.
Genet. 5,1533
-1538.
Groves, A. K. and Bronner-Fraser, M. (2000).
Competence, specification and commitment in otic placode induction.
Development 127,3489
-3499.
Hukriede, N., Fisher, D., Epstein, J., Joly, L., Tellis, P.,
Zhou, Y., Barbazuk, B., Cox, K., Fenton-Noriega, L., Hersey, C. et al.
(2001). The LN54 radiation hybrid map of zebrafish expressed
sequences. Genome Res.
11,2127
-2132.
Hukriede, N. A., Joly, L., Tsang, M., Miles, J., Tellis, P.,
Epstein, J. A., Barbazuk, W. B., Li, F. N., Paw, B., Postlethwait, J. H. et
al. (1999). Radiation hybrid mapping of the zebrafish genome.
Proc. Natl. Acad. Sci. USA
96,9745
-9750.
Hulander, M., Wurst, W., Carlsson, P. and Enerback, S. (1998). The winged helix transcription factor Fkh10 is required for normal development of the inner ear. Nat. Genet. 20,374 -376.[CrossRef][Medline]
Itoh, M., Kudoh, T., Dedekian, M., Kim, C. H. and Chitnis, A.
B. (2002). A role for iro1 and iro7 in the establishment of
an anteroposterior compartment of the ectoderm adjacent to the
midbrain-hindbrain boundary. Development
129,2317
-2327.
Jacobson, A. G. (1963). The determination and positioning of the nose, lens and ear. III. Effects of reversing the antero-posterior axis of epidermis, neural plate and neural fold. J. Exp. Zool. 154,285 -291.
Jacobson, A. G. (1966). Inductive processes in embryonic development. Science 152, 25-34.[Medline]
Jowett, T. (2001). Double in situ hybridization techniques in zebrafish. Methods 23,345 -358.[CrossRef][Medline]
Kaufmann, E. and Knochel, W. (1996). Five years on the wings of fork head. Mech. Dev. 57, 3-20.[CrossRef][Medline]
Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B. and Schilling, T. F. (1995). Stages of embryonic development of the zebrafish. Dev. Dyn. 203,253 -310.[Medline]
Kops, G. J. and Burgering, B. M. (1999). Forkhead transcription factors: new insights into protein kinase B (c-akt) signaling. J. Mol. Med. 77,656 -665.[CrossRef][Medline]
Kozlowski, D. J., Murakami, T., Ho, R. K. and Weinberg, E. S. (1997). Regional cell movement and tissue patterning in the zebrafish embryo revealed by fate mapping with caged fluorescein. Biochem. Cell Biol. 75,551 -562.[CrossRef][Medline]
Krauss, S., Johansen, T., Korzh, V. and Fjose, A. (1991). Expression of the zebrafish paired box gene pax[zf-b] during early neurogenesis. Development 113,1193 -1206.[Abstract]
Kudoh, T., Tsang, M., Hukriede, N. A., Chen, X., Dedekian, M.,
Clarke, C. J., Kiang, A., Schultz, S., Epstein, J. A., Toyama, R. et al.
(2001). A gene expression screen in zebrafish embryogenesis.
Genome Res. 11,1979
-1987.
Ladher, R. K., Anakwe, K. U., Gurney, A. L., Schoenwolf, G. C.
and Francis-West, P. H. (2000). Identification of synergistic
signals initiating inner ear development. Science
290,1965
-1967.
Lai, E., Prezioso, V. R., Smith, E., Litvin, O., Costa, R. H. and Darnell, J. E., Jr (1990). HNF-3A, a hepatocyte-enriched transcription factor of novel structure is regulated transcriptionally. Genes Dev. 4,1427 -1436.[Abstract]
Larsson, C., Hellqvist, M., Pierrou, S., White, I., Enerback, S. and Carlsson, P. (1995). Chromosomal localization of six human forkhead genes, freac-1 (FKHL5), -3 (FKHL7), -4 (FKHL8), -5 (FKHL9), -6 (FKHL10), and -8 (FKHL12). Genomics 30,464 -469.[CrossRef][Medline]
Lef, J., Clement, J. H., Oschwald, R., Koster, M. and Knochel, W. (1994). Spatial and temporal transcription patterns of the forkhead related XFD-2/XFD-2' genes in Xenopus laevis embryos. Mech. Dev. 45,117 -126.[Medline]
Lef, J., Dege, P., Scheucher, M., Forsbach-Birk, V., Clement, J. H. and Knochel, W. (1996). A fork head related multigene family is transcribed in Xenopus laevis embryos. Int. J. Dev. Biol. 40,245 -253.[Medline]
Li, Y., Allende, M. L., Finkelstein, R. and Weinberg, E. S. (1994). Expression of two zebrafish orthodenticle-related genes in the embryonic brain. Mech. Dev. 48,229 -244.[CrossRef][Medline]
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.[Medline]
Luo, R., An, M., Arduini, B. L. and Henion, P. D. (2001). Specific panneural crest expression of zebrafish Crestin throughout embryonic development. Dev. Dyn. 220,169 -174.[CrossRef][Medline]
Mahmood, R., Kiefer, P., Guthrie, S., Dickson, C. and Mason,
I. (1995). Multiple roles for FGF-3 during cranial neural
development in the chicken. Development
121,1399
-1410.
Malicki, J., Schier, A. F., Solnica-Krezel, L., Stemple, D. L.,
Neuhauss, S. C., Stainier, D. Y., Abdelilah, S., Rangini, Z., Zwartkruis, F.
and Driever, W. (1996). Mutations affecting development of
the zebrafish ear. Development
123,275
-283.
Mansouri, A., Chowdhury, K. and Gruss, P. (1998). Follicular cells of the thyroid gland require Pax8 gene function. Nat. Genet. 19, 87-90.[CrossRef][Medline]
Mendonsa, E. S. and Riley, B. B. (1999). Genetic analysis of tissue interactions required for otic placode induction in the zebrafish. Dev. Biol. 206,100 -112.[CrossRef][Medline]
Miller, L. M., Gallegos, M. E., Morisseau, B. A. and Kim, S. K. (1993). lin-31, a Caenorhabditis elegans HNF-3/fork head transcription factor homolog, specifies three alternative cell fates in vulval development. Genes Dev. 7, 933-947.[Abstract]
Moens, C. B., Cordes, S. P., Giorgianni, M. W., Barsh, G. S. and
Kimmel, C. B. (1998). Equivalence in the genetic control of
hindbrain segmentation in fish and mouse. Development
125,381
-391.
Nasevicius, A. and Ekker, S. C. (2000). Effective targeted gene `knockdown' in zebrafish. Nat. Genet. 26,216 -220.[CrossRef][Medline]
Odenthal, J. and Nusslein-Volhard, C. (1998). fork head domain genes in zebrafish. Dev. Genes Evol. 208,245 -258.[CrossRef][Medline]
Overdier, D. G., Ye, H., Peterson, R. S., Clevidence, D. E. and
Costa, R. H. (1997). The winged helix transcriptional
activator HFH-3 is expressed in the distal tubules of embryonic and adult
mouse kidney. J. Biol. Chem.
272,13725
-13730.
Oxtoby, E. and Jowett, T. (1993). Cloning of the zebrafish krox-20 gene (krx-20) and its expression during hindbrain development. Nucleic Acids Res. 21,1087 -1095.[Abstract]
Pfeffer, P. L., Gerster, T., Lun, K., Brand, M. and Busslinger,
M. (1998). Characterization of three novel members of the
zebrafish Pax2/5/8 family: dependency of Pax5 and Pax8 expression on the
Pax2.1 (noi) function. Development
125,3063
-3074.
Phillips, B. T., Bolding, K. and Riley, B. B. (2001). Zebrafish fgf3 and fgf8 encode redundant functions required for otic placode induction. Dev. Biol. 235,351 -365.[CrossRef][Medline]
Pierrou, S., Hellqvist, M., Samuelsson, L., Enerback, S. and Carlsson, P. (1994). Cloning and characterization of seven human forkhead proteins: binding site specificity and DNA bending. EMBO J. 13,5002 -5012.[Abstract]
Piotrowski, T. and Nusslein-Volhard, C. (2000). The endoderm plays an important role in patterning the segmented pharyngeal region in zebrafish (Danio rerio). Dev. Biol. 225,339 -356.[CrossRef][Medline]
Piotrowski, T., Schilling, T. F., Brand, M., Jiang, Y. J.,
Heisenberg, C. P., Beuchle, D., Grandel, H., van Eeden, F. J., Furutani-Seiki,
M., Granato, M. et al. (1996). Jaw and branchial arch mutants
in zebrafish II: anterior arches and cartilage differentiation.
Development 123,345
-356.
Pogoda, H. M., Solnica-Krezel, L., Driever, W. and Meyer, D. (2000). The zebrafish forkhead transcription factor FoxH1/Fast1 is a modulator of nodal signaling required for organizer formation. Curr. Biol. 10,1041 -1049.[CrossRef][Medline]
Pohl, B. S., Knochel, S., Dillinger, K. and Knochel, W. (2002). Sequence and expression of FoxB2 (XFD-5) and FoxI1c (XFD-10) in Xenopus embryogenesis. Mech. Dev. 117,283 -287.[CrossRef][Medline]
Postlethwait, J. H., Yan, Y. L., Gates, M. A., Horne, S., Amores, A., Brownlie, A., Donovan, A., Egan, E. S., Force, A., Gong, Z. et al. (1998). Vertebrate genome evolution and the zebrafish gene map. Nat. Genet. 18,345 -349.[Medline]
Represa, J., Leon, Y., Miner, C. and Giraldez, F. (1991). The int-2 proto-oncogene is responsible for induction of the inner ear. Nature 353,561 -563.[CrossRef][Medline]
Rubinstein, A. L., Lee, D., Luo, R., Henion, P. D. and Halpern, M. E. (2000). Genes dependent on zebrafish cyclops function identified by AFLP differential gene expression screen. Genesis 26,86 -97.[CrossRef][Medline]
Schier, A. F., Neuhauss, S. C., Helde, K. A., Talbot, W. S. and
Driever, W. (1997). The one-eyed pinhead gene
functions in mesoderm and endoderm formation in zebrafish and interacts with
no tail. Development
124,327
-342.
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.
Shimoda, N., Knapik, E. W., Ziniti, J., Sim, C., Yamada, E., Kaplan, S., Jackson, D., de Sauvage, F., Jacob, H. and Fishman, M. C. (1999). Zebrafish genetic map with 2000 microsatellite markers. Genomics 58,219 -232.[CrossRef][Medline]
Solomon, K. S. and Fritz, A. (2002). Concerted
action of two dlx paralogs in sensory placode formation.
Development 129,3127
-3136.
Strahle, U., Blader, P., Henrique, D. and Ingham, P. W. (1993). Axial, a zebrafish gene expressed along the developing body axis, shows altered expression in cyclops mutant embryos. Genes Dev. 7,1436 -1446.[Abstract]
The Treacher Collins Syndrome Collaborative Group (1996). Positional cloning of a gene involved in the pathogenesis of Treacher Collins Syndrome. Nat. Genet. 12,130 -136.[Medline]
Thisse, C. and Thisse, B. (1998). High resolution whole-mount in situ hybridization. The Zebrafish Science Monitor 5,8 -9.
Topczewska, J. M., Topczewski, J., Solnica-Krezel, L. and Hogan, B. L. (2001). Sequence and expression of zebrafish foxc1a and foxc1b, encoding conserved forkhead/winged helix transcription factors. Mech. Dev. 100,343 -347.[CrossRef][Medline]
Toresson, H., Martinez-Barbera, J. P., Bardsley, A., Caubit, X. and Krauss, S. (1998). Conservation of BF-1 expression in amphioxus and zebrafish suggests evolutionary ancestry of anterior cell types that contribute to the vertebrate telencephalon. Dev. Genes Evol. 208,431 -439.[CrossRef][Medline]
Torres, M. and Giraldez, F. (1998). The development of the vertebrate inner ear. Mech. Dev. 71, 5-21.[CrossRef][Medline]
Vendrell, V., Carnicero, E., Giraldez, F., Alonso, M. T. and
Schimmang, T. (2000). Induction of inner ear fate by FGF3.
Development 127,2011
-2019.
Waddington, C. H. (1937). The determination of the auditory placode in the chick. J. Exp. Biol. 14,232 -239.
Weigel, D., Jurgens, G., Kuttner, F., Seifert, E. and Jackle, H. (1989). The homeotic gene fork head encodes a nuclear protein and is expressed in the terminal regions of the Drosophila embryo. Cell 57,645 -658.[Medline]
Westerfield, M. (2000). The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio rerio). Eugene, OR: University of Oregon Press.
Whitfield, T. T., Granato, M., van Eeden, F. J., Schach, U.,
Brand, M., Furutani-Seiki, M., Haffter, P., Hammerschmidt, M., Heisenberg, C.
P., Jiang, Y. J. et al. (1996). Mutations affecting
development of the zebrafish inner ear and lateral line.
Development 123,241
-254.
Wise, C. A., Chiang, L. C., Paznekas, W. A., Sharma, M., Musy,
M. M., Ashley, J. A., Lovett, M. and Jabs, E. W. (1997).
TCOF1 gene encodes a putative nucleolar phosphoprotein that exhibits
mutations in Treacher Collins Syndrome throughout its coding region.
Proc. Natl. Acad. Sci. USA
94,3110
-3115.
Woo, K. and Fraser, S. E. (1997). Specification
of the zebrafish nervous system by nonaxial signals.
Science 277,254
-257.
Xu, P. X., Adams, J., Peters, H., Brown, M. C., Heaney, S. and Maas, R. (1999). Eya1-deficient mice lack ears and kidneys and show abnormal apoptosis of organ primordia. Nat. Genet. 23,113 -117.[CrossRef][Medline]
Xuan, S., Baptista, C. A., Balas, G., Tao, W., Soares, V. C. and Lai, E. (1995). Winged helix transcription factor BF-1 is essential for the development of the cerebral hemispheres. Neuron 14,1141 -1152.[Medline]
Yntema, C. L. (1933). Experiments on the determination of the ear ectoderm in the embryo of amblyostoma punctatum.J. Exp. Zool. 65,317 -357.
Yntema, C. L. (1950). An induction of the ear from foreign ectoderm in the salamander. J. Exp. Zool. 113,211 -244.