Mutation of the atrophin2 gene in the zebrafish
disrupts signaling by fibroblast growth factor
during development of the inner ear
Yukako Asai, Dylan K. Chan*, Catherine J. Starr†, James A. Kappler, Richard Kollmar‡, and A. J. Hudspeth§
Howard Hughes Medical Institute and Laboratory of Sensory Neuroscience, The Rockefeller University, 1230 York Avenue, New York, NY 10021-6399
Contributed by A. J. Hudspeth, April 27, 2006
The development of the vertebrate inner ear depends on the
precise expression of fibroblast growth factors. In a mutagenesis
screen for zebrafish with abnormalities of inner-ear development
and behavior, we isolated a mutant line, ru622, whose phenotypic
characteristics resembled those of null mutants for the gene
encoding fibroblast growth factor 8 (Fgf8): an inconsistent startle
response, circular swimming, fused otoliths, and abnormal semi-
circular canals. Positional cloning disclosed that the mutant gene
encodes the transcriptional corepressor Atrophin2. Both the Fgf8
protein and zebrafish ‘‘similar expression to fgf genes’’ protein
(Sef), an antagonist of fibroblast growth factors induced by Fgf8
itself, were found to be overexpressed in ru622 mutants. We
therefore hypothesized that an excess of Sef eliminates Fgf8
signals and produces an fgf8 null phenotype in ru622 mutants. In
support of this idea, we could rescue larvae whose atrophin2
expression had been diminished with morpholinos by reducing the
expression of Sef as well. We propose that Atrophin2 plays a role
in the feedback regulation of Fgf8 signaling. When mutation of the
atrophin2 gene results in the overexpression of both Fgf8 and Sef,
the excessive Sef inhibits Fgf8 signaling. The resultant imbalance
of Fgf8 and Sef signals then underlies the abnormal aural devel-
opment observed in ru622.
auditory system ? hearing ? vestibular system
contains several distinct sensory epithelia that are variously
dedicated to the detection of linear acceleration, angular accel-
eration, and sound. Each sensory epithelium includes mech-
anosensory hair cells that are separated by nonsensory support-
ing cells and innervated by afferent terminals from the eighth
cranial nerve (reviewed in ref. 1).
The ear’s development involves dramatic morphological
changes, first from a simple epithelial sheet to a closed otic
vesicle, and subsequently to the more complex topology of the
adult labyrinth (2). Classical genetics offers one fruitful avenue
to the identification of genes whose products play important
roles in this progression. The zebrafish is particularly useful in
this context, for the availability of large numbers of progeny
facilitates genetic mapping. This species is also valuable in
developmental studies owing to its rapid external development
and the optical transparency of its larvae. Finally, because the
inner ear of the zebrafish resembles those of other vertebrates,
mechanisms elucidated in this organism are likely to prove of
The development of the zebrafish’s inner ear commences
during somitogenesis with the formation of the otic placode, an
area of thickened ectoderm adjacent to the hindbrain that soon
forms the hollow otic vesicle (3, 4). Two otoliths originate at the
21-somite stage as tiny, glistening granules of calcium carbonate
at the anterior and posterior ends of the otic vesicle’s lumen. By
3 days postfertilization (dpf), the anterior otolith has become a
prolate ellipsoid and the larger, posterior otolith an oblate
ellipsoid (Fig. 1). At the same time, the otic vesicle’s structure
he complex structure of the inner ear is largely conserved
across vertebrate species. In an adult animal, the inner ear
changes dramatically during the formation of the semicircular
canals (5). Three finger-like protrusions first extend from the
anterior, posterior, and lateral walls of the otic vesicle. After
these protrusions have fused together, another extends from the
ventral wall and at 3 dpf fuses with the others at the center of the
otocyst. By 5 dpf, sensory cristae that detect angular acceleration
have developed in each of the three semicircular canals.
To identify proteins involved in the development and opera-
tion of the inner ear, we conducted a mutagenesis screen in
zebrafish and isolated 16 mutant lines. In the present study, we
report the phenotype of one line, ru622, identify the affected
gene, and test hypotheses about the function of the gene’s
Phenotype of ru622 Mutants. The zebrafish mutant line ru622 was
isolated in a mutagenesis screen on the basis of an inconsistent
acoustic startle reflex at 5 dpf. Most WT larvae responded to
acoustic stimuli, such as gentle taps on the edge of the dish that
contained them, with a stereotyped escape reaction (6). Ho-
mozygous ru622 mutant larvae, however, did not react consis-
tently to acoustic stimulation. Furthermore, the mutant animals
usually remained on their sides at the bottom of their dish or
swam upside-down or in circles, all phenotypes that are associ-
ated with abnormalities of vestibular function (7).
Homozygous mutant animals appeared grossly normal and
were responsive to tactile stimulation. Perhaps because their
abnormal swimming prevented effective feeding, however, they
generally died on the seventh day of development. The structure
of their inner ears was almost normal save that in 8 of 28 ru622
mutants each ear contained only one, often misshapen otolith
(Fig. 1 A and B). This phenotype may have resulted from the
fusion of the two otoliths that occur normally; cellular labeling
disclosed that the anterior and posterior maculae often came
into close contact in mutant animals (data not shown). More-
over, 20 of 65 mutants had abnormal semicircular canals (Fig. 1
D and E). The epithelial protrusions that define the canals were
sometimes absent; when present, they failed to elongate or fuse.
The two phenotypes occurred independently; some larvae dis-
Conflict of interest statement: No conflicts declared.
Freely available online through the PNAS open access option.
Abbreviations: dpf, days postfertilization; hpf, hours postfertilization; Fgf8, zebrafish
fibroblast growth factor 8; Sef, zebrafish ‘‘similar expression to fgf genes’’ protein.
*Present address: Medical Scientist Training Program, Weill Cornell Medical College,
1300 York Avenue, New York, NY 10021.
†Present address: American Museum of Natural History, Central Park West at 79th Street,
New York, NY 10024.
‡Present address: Department of Molecular and Integrative Physiology and Beckman
Institute for Advanced Science and Technology, University of Illinois at Urbana–Cham-
paign, Urbana, IL 61801.
§To whom correspondence should be addressed. E-mail: email@example.com.
© 2006 by The National Academy of Sciences of the USA
June 13, 2006 ?
vol. 103 ?
no. 24 ?
played only the otolithic phenotype, some showed only the
semicircular-canal phenotype, and some had both.
Close examination of mutant larvae disclosed that their head
structure was also abnormal. In particular, the palatoquadrate
cartilages were shorter and the ceratohyal cartilages were ori-
ented at a greater angle than in WT animals (Fig. 1 G and H).
Like ru622 mutants, some larvae bearing the acerebellar
mutation that impairs zebrafish fibroblast growth factor 8 (Fgf8)
in the semicircular canals (8). Injection of morpholinos against
fgf8 yields a similar phenotype (Fig. 1 C and F). Like acerebellar
against fgf8 show defects of cartilage formation in the first and
second pharyngeal arches (Fig. 1I).
Hair-Cell Morphology and Function. The abnormalities of ru622
mutants suggested that the development or functioning of
sensory hair cells was affected. To determine whether the
structure of hair bundles was disturbed by the mutation, we
labeled the stereocilia of ru622 mutants with phalloidin and their
kinocilia with antibodies against acetylated tubulin. At the
light-microscopic level, the hair bundles of mutants appeared to
be normal (Fig. 2 A and B).
The functioning of hair cells in the ears of ru622 mutants was
examined by recording the microphonic potentials evoked by
signals were present in mutants, they were consistently dimin-
ished in comparison with those of WT fish.
The lateral-line system, which underlies a fish’s sensitivity to
predators, prey, and obstacles, also has hair cells whose hair
bundles are precisely oriented with respect to the body axes (10,
11). Labeling of the hair bundles in ru622 mutants disclosed that
the vectorial orientation of these cells was normal (Fig. 2 D and
E). The functioning of lateral-line hair cells was assessed by
observing the uptake of 4-(4-(diethylamino)styryl)-N-
methylpyridinium iodide, a fluorophore that enters through
normal transduction channels. The number of neuromasts la-
beled on each side of a mutant, 23 ? 2.0 (mean ? SD, n ? 64)
was slightly but significantly smaller than the value for a WT
animal contains two otoliths, a smaller, anterior one lying in the horizontal
plane and a larger, posterior one oriented vertically. (B and C) By contrast, an
ru622 mutant (B) or a larva treated with a morpholino against fgf8 (C)
begin to be delineated at this stage by three pillars extending from the
anterior, posterior, and ventral walls of the otic vesicle and by a dorsolateral
F) In an ru622 mutant (E) or an fgf8 knockdown larva (F), the formation of
semicircular canals is abnormal. (G) Alcian blue staining reveals the pattern of
cartilage formation in the rostral region of a WT larva. P, palatoquadrate
cartilage; C, ceratohyal cartilage; M, Meckel’s cartilage. (H and I) An ru622
mutant (H) or an fgf8 knockdown larva (I) is distinguished by shorter palato-
quadrate cartilages and the orientation of the ceratohyal cartilages at an
increased angle. (Scale bars: 20 ?m for A–F; 100 ?m for G–I.)
Morphological features of WT (A, D, and G), ru622 mutant (B, E, and
image of the crista in the lateral semicircular canal of a larva at 5 dpf, Alexa
Fluor 568-phalloidin (red) labels the stereocilia and an antiserum against
acetylated tubulin (green) labels the kinocilia and cellular cytoskeletons. (B)
The crista of a mutant larva displays similar features. (C) Extracellular micro-
phonic recordings at 5 dpf indicate that the transduction current of a WT otic
of the apical surface of a WT neuromast stained with Alexa Fluor 568-
phalloidin, the axis of each hair bundle’s polarity is denoted by a notch in the
actin-rich cuticular plate corresponding to the position of the kinocilium. The
arrows indicate the polarities observed. (E) The hair bundles of an ru622
mutant display normal polarization. (D) Fluorescence imaging of a WT larva
exposed to 4-(4-(diethylamino)styryl)-N-methylpyridinium iodide reveals a
stereotyped pattern of labeled neuromasts. (F) The labeling pattern is similar
in an ru622 mutant, but the average number of labeled neuromasts is smaller
than in a WT larva. (Scale bars: 10 ?m for A, B, D, and E; 500 ?m for F and G.)
Phenotypic characteristics of the ru622 mutation. (A) In a confocal
www.pnas.org?cgi?doi?10.1073?pnas.0603453103Asai et al.
animal, 28 ? 1.5 (n ? 50; P ? 0.001 by Student’s one-tailed t
test). The intensity of 4-(4-(diethylamino)styryl)-N-methylpyri-
dinium iodide labeling was also diminished in mutant neuro-
masts, possibly as a result of a reduction in the number of hair
cells (Fig. 2 F and G).
Identification of the Mutant Gene. We used positional cloning to
identify the mutation responsible for the ru622 phenotype. By
low- and intermediate-resolution genetic mapping, we linked the
mutation to the simple sequence-length polymorphic markers
z15422 and z42693 on chromosome 23 (Fig. 3A). Through the
genotyping of 2,247 mutant animals, which yielded 13 recombi-
to be separated by ?0.3 cM. Although this region includes
another marker, z23370, no recombinations occurred among the
genotyped larvae between this marker and the mutation. A
bacterial artificial chromosome contig from the zebrafish-
genome project, ctg10165, was found to include both z23370 and
z42693. The 13 recombinants for z15422 were tested for recom-
bination with single-nucleotide polymorphisms identified at the
end of this contig nearer z23370. Because one larva displayed a
recombination event between these polymorphisms and the
mutation, ctg10165 contained the complete candidate region
bearing the mutation.
Of the five expressed sequence tags localized to the ?280-kb
candidate region, one was identified by homology with the
corresponding genes of other species as the atrophin2 gene.
Reverse transcription and PCRs were performed on WT and
mutant larvae to examine this gene’s coding region. In contrast
to the single band amplified from WT animals, mutants yielded
two bands (Fig. 3B); this result raised the possibility of a
mutation at an exon–intron boundary. Sequencing disclosed that
one product, ?1,600 bp in length, included the entirety of the
?1-kb intron 12. In the other, smaller products amplified from
mutants, splicing was found to have occurred at two GT dinucle-
otides downstream from the ordinary boundary between exons
12 and 13. As a result, the shorter transcripts from ru622 mutants
included either a 4-bp or a 16-bp insertion between exons 12 and
13. Sequence analysis of the exon–intron boundaries of the
mutant atrophin2 gene demonstrated a single-base change (G to
A), the putative mutation, at the 5? splicing site of intron 12.
Known as RERE in humans, Atrophin2 was originally iden-
tified as a homolog of Atrophin1, the protein affected in the
neurodegenerative disease dentatorubral-pallidoluysian atrophy
(12). The coding region of the zebrafish’s atrophin2 gene com-
prises 21 exons spanning ?110 kb of the genome (Fig. 3C). The
human RERE gene includes an additional exon following the
first; the transcript lacking this exon represents a minor splicing
variant. The zebrafish genome appears to lack the corresponding
exon, however, for it could be found neither by our cDNA
cloning procedure nor by searching genomic databases.
The predicted Atrophin2 protein of zebrafish is 1,521 aa in
length. Its amino-terminal portion includes four domains that
resemble those of metastasis-associated protein 2 (Mta2): the
homology 2 (ELM2) domain; the SWI3, ADA2, N-CoR, and
TFIIIB (SANT) domain; and the zinc-finger (GATA) domain
(Fig. 3C). The carboxyl-terminal half of Atophin2 resembles
Atrophin1 and shares arginine-glutamic acid (RE) dipeptide
The mutant transcripts all include stop codons shortly after
exon 12 and should therefore produce truncated proteins (Fig.
3B). To test the supposition that deficiency of Atrophin2 un-
derlies the ru622 mutant phenotype, we injected WT eggs with
showed the same behavioral and morphological phenotypes as
ru622 mutants (data not shown). Moreover, morpholino injec-
tion reduced the number of hair cells in the anterior macula at
5 dpf from 57 ? 3 (n ? 4) to 47 ? 7 hair cells (n ? 9; P ? 0.01
by Student’s one-tailed t test).
Expression Pattern of Zebrafish atrophin2. We used in situ hybrid-
ization to examine the expression pattern of mRNA encoding
Atrophin2 in zebrafish. At all of the developmental stages tested,
including the late gastrula and 24 or 36 h postfertilization (hpf),
essentially every cell expressed atrophin2, although the expres-
sion in the trunk was weak at the last stage (Fig. 4). The epithelia
of the otic vesicle were labeled strongly. The labeling pattern was
identical when an antisense probe to the 5? portion of the mRNA
genomic region surrounding the ru622 locus on zebrafish chromosome 23
displays the number of recombinations with respect to five simple sequence-
length polymorphisms (blue and green) and a single-nucleotide polymor-
phism near the end of the bacterial artificial chromosome contig ctg10165
(purple). A total of 2,247 larvae were examined for each marker save z6142,
atrophin2 gene and the red line and asterisk denote the ru622 mutation. (B)
When WT RNA is used as a template, RT-PCR amplification of a cDNA region
another ?1,600 bp long. The ru622 mutation occurred at the boundary
(Right). The smaller RT-PCR band from the ru622 reaction includes two prod-
ucts. In one, splicing occurs 4 bp downstream from the mutated splice site
The large RT-PCR product contains the entirety of intron 12. (C) The genomic
arrow indicates the site of the ru622 mutation; the blue bars denote the
targets of morpholinos directed against the translation start site and the
splicing site between intron 10 and exon 11. Atrophin2 includes a bromo-
adjacent homology domain (BAH), an EGL27 and Mta1 homology 2 domain
(ELM2), a SWI3, ADA2, N-CoR and TFIIIB domain (SANT), and a zinc finger
domain (GATA). The two arrowheads indicate RE repeats.
Identification of the basis of the ru622 mutation. (A) A map of the
Asai et al.PNAS ?
June 13, 2006 ?
vol. 103 ?
no. 24 ?
was used. The corresponding sense probes, however, produced
no signal at any stage (data not shown).
In Situ Hybridization with Probes for fgf8 and sef. Because murine
atrophin2 mutants display misregulation of fgf8 expression (13),
we examined the expression pattern of zebrafish fgf8 by in situ
hybridization. Consistent with the demonstrated function of
Atrophin2 as a transcriptional corepressor (13–15), the expres-
sion of fgf8 mRNA was broader in fish injected with the
morpholino directed against atrophin2 splicing than in control
animals (Fig. 5 A and B). As estimated by quantitative PCR,
animals exposed to the atrophin2 morpholino expressed an
increased concentration of fgf8 transcripts in comparison with
control fish (Fig. 5E). Because the expression pattern of fgf8
mRNA in the ru622 mutant was opposite that expected from the
phenotype, and Fgf8 is known to activate antagonists of fibro-
blast growth factor signaling (16–18), we next examined the
possibility that the overexpression of fgf8 in the ru622 mutant
induced overproduction of an antagonist. As we expected, the
zebrafish ‘‘similar expression to fgf genes’’ protein (Sef), a known
antagonist of fibroblast growth factor, was expressed with a
similar pattern but more robustly in fish injected with the
morpholino against atrophin2 splicing than in control animals
(Fig. 5 C and D). In situ hybridization indicated that the
expression of the transcription factor pax2.1, which is expressed
at the midbrain-hindbrain boundary and in the otic vesicle at the
same developmental stage, was not affected by overexpression of
fgf8 (data not shown). Quantitative PCR confirmed that expo-
sure to the atrophin2 morpholino significantly increased the
expression of transcripts for fgf8, sef, and sprouty4, another
antagonist of Fgf signaling (Fig. 5E).
Rescue of the ru622 Phenotype. If overproduction of Sef eliminates
Fgf8 signals, resulting in a phenocopy of the fgf8 null mutation,
then it should be possible to rescue affected larvae by reducing
the amount of Sef protein. A morpholino against sef was
At 5 dpf, seven of eight (88%) larvae injected with the atrophin2
morpholino alone showed the ru622 phenotype of abnormal
equilibrium. Coinjection reduced the fraction of abnormal an-
imals to 23 of 53 (43%) for 1 ng of the sef morpholino and to 10
of 34 (29%) for 5 ng. In a control experiment, the use of 5 ng of
a sef morpholino with five mismatches resulted in 25 of 34 (74%)
abnormal animals. These results suggest that overproduced Sef
protein eliminates Fgf8 signals, causing ru622 fish to manifest a
phenotype similar to that of fgf8 null mutants.
Originally identified in a mutagenesis screen on the basis of
defective startle responses, zebrafish of the ru622 line have been
shown to be affected by mutation of the atrophin2 gene. Con-
served throughout the metazoans, atrophins act as transcrip-
tional corepressors whose principal effects, especially in the
mouse, involve signaling by fibroblast growth factors (13–15).
in zebrafish is the similarity of its phenotype to that associated
with the acerebellar mutation, which inactivates the fgf8 gene (8,
9). Both ru622 mutants and fgf8 morphants display similar
defects in the formation of otoliths, semicircular canals, and
(B) of the late gastrula. (C) At 24 hpf, essentially all tissue except the yolk is
labeled. (D) Labeling diminishes from the caudal extreme by 36 hpf. (E)
at 36 hpf. (Scale bar: 100 ?m.)
The expression pattern of the atrophin2 gene. Whole-mount in situ
telencephalon, optic stalk, isthmus, otic vesicle, developing somites, and tail
bud. (B) fgf8 is expressed more broadly in an atrophin2 knockdown embryo.
(C) A 26-somite control embryo treated with sef probe displays prominent
shows a similar pattern of labeling but more extensive expression than in
messages for fgf8, sef, and sprouty4 in atrophin2 knockdown animals; signif-
icance values were obtained with Student’s one-tailed t test. Control animals
were injected with water. The error bars indicate standard deviations. (Scale
bar: 100 ?m.)
www.pnas.org?cgi?doi?10.1073?pnas.0603453103 Asai et al.
head cartilage (Fig. 1). Unlike ru622 mutants, however, acer-
ebellar larvae lose their midbrain-hindbrain boundary. It is
possible that maternally supplied Atrophin2 prevents such de-
fects in early development.
Knocking out the atrophin2 gene disrupts fibroblast growth
factor signaling in the mouse. In the neural plate of affected
animals, the expression of fgf8 is not appropriately limited to the
anterior neural ridge (13). Mutant animals additionally show
diminished expression of sonic hedgehog along the anterior
midline. Although sonic hedgehog plays a role in inner-ear
development (19–22), it is unlikely for two reasons that reduc-
tion of this signal accounts for the phenotype of ru622 mutants.
First, the otic vesicle’s formation is initiated by signals from the
hindbrain, where the expression of sonic hedgehog is normal in
atrophin2 knockout mice. Second, although zebrafish slow mus-
cle omitted mutants are deficient in responsiveness to all hedge-
hog ligands (21, 23), ru622 larvae clearly differ phenotypically
from these animals. Unlike the inner ears of ru622 and fgf8 null
mutants, those of slow muscle omitted animals are symmetrical
along the anteroposterior axis; the two otoliths, for example,
both resemble the normal anterior otolith (21). Because Atro-
phin2 is ubiquitously expressed, the localized defects caused by
its deficiency may reflect the specificity of interaction between
Atrophin2 and nuclear receptors. Atrophin2 binds to several
nuclear receptors of subfamily 2, but not to those of subfamily 1
(24). The effects of knocking out atrophin2 gene function on the
later development of the mouse’s ear unfortunately cannot be
investigated, for mutants succumb to heart failure shortly after
the appearance of the otic vesicles.
Why might the phenotype of larvae overexpressing fgf8 re-
semble that of fgf8 null mutants? We hypothesize that Atrophin2
acts in the zebrafish as a corepressor in the regulation of fgf8 and
sef expression. Fgf8 evidently down-regulates its own expression
by negative feedback: in acerebellar mutants, transcription of the
fgf8 gene is elevated (8). If Atrophin2 is involved in this pathway,
then the expression of Fgf8 may not be down-regulated in
atrophin2 mutants owing to the absence of appropriate feedback.
In addition, the binding of Fgf8 to its receptor is known to
activate the expression of antagonists to signaling by fibroblast
growth factors (16–18). It is therefore plausible that excessive
Fgf8 protein in atrophin2 mutants induces additional sef expres-
sion as well (Fig. 5B). Although Sef activity should normally
silence the Fgf pathway and inhibit Sef expression, an absence of
Atrophin2 may disable this negative feedback and permit con-
tinuing expression of Sef. The activity of Sef may then dominate
that of Fgf8, leaving larvae with an fgf8 null phenotype. This
schema is supported by three results. First, our results from
quantitative PCR and in situ hybridization indicate that injection
of a morpholino against atrophin2 enhances the expression of
both fgf8 and sef (Fig. 5). Next, the injection of sef mRNA into
embryos produces a phenotype similar to that of fgf8 deficiency,
including a smaller otic vesicle (17). Finally, reducing the ex-
pression of sef rescues larvae whose atrophin2 expression has
been lowered with morpholinos.
The ru622 mutation provides an interesting demonstration of
the developmental consequences of imbalanced signaling be-
tween a growth factor and its antagonist. A more detailed
analysis of the inner ear’s sensory organs, including the patterns
of hair-cell polarization, afferent and efferent innervation, and
neural projection, may reveal additional consequences of the
mutation. More importantly, further studies are needed to
ascertain how perturbation of fibroblast growth factor signaling
culminates in the observed structural and functional phenotypes.
Materials and Methods
Fish Husbandry and Mutagenesis. Zebrafish were maintained and
bred at 28°C by standard procedures (25). The ru622 line was
isolated in an F3screen after mutagenesis with ethylnitrosourea
Alcian Blue Staining. Larvae were fixed for 12 h at 4°C in 4%
(wt?vol) paraformaldehyde in PBS, rinsed with PBS containing
0.1% (vol?vol) Tween, and transferred for 12 h into 0.1%
(wt?vol) alcian blue in 80% (vol?vol) ethanol and 20% (vol?vol)
glacial acetic acid. The specimens were rinsed in ethanol and
rehydrated gradually in an ethanol series. After treatment for
tetraborate, larvae were bleached in 3% (vol?vol) H2O2and 1%
(wt?vol) KOH, then placed in 70% (wt?vol) glycerol with 1%
(wt?vol) KOH for microscopic observation.
Immunolabeling and Phalloidin Labeling. Larvae were fixed, per-
meabilized, and labeled in blocking solution as described (26).
(6-11B-1; Sigma) were used at a dilution of 1?1,000; the Alexa
Fluor 488-conjugated goat anti-mouse IgG secondary antiserum
(Molecular Probes) was used at 1?100. We labeled filamentous
actin with 2 units per ml of Alexa Fluor 568-conjugated phal-
loidin (Molecular Probes).
Microphonic-Potential Recording and Neuromast Labeling. Micro-
phonic potentials produced by whole maculae in response to
vibrational stimulation were measured as described (27) from
WT and ru622 mutant larvae at 5 dpf. To label neuromasts (26),
we immersed larvae for 5 min in 200 ?M 4-(4-(diethylamino)s-
tyryl)-N-methylpyridinium iodide (Molecular Probes), then
rinsed them several times with water.
Positional Cloning. In a mapping outcross, ru622 heterozygotes of
the AB strain were mated with WT WIK animals. A pair of
carriers of the ru622 mutation were identified among the off-
spring and inbred; their progeny were screened for the behav-
ioral phenotypes and fixed in methanol.
Genomic DNA isolation and PCRs were performed as de-
scribed (28). Bulked-segregant analysis was conducted with
primers (Research Genetics, Huntsville, AL) flanking each of
215 simple sequence-length polymorphisms. After the chromo-
somal linkage had been established, additional markers were
found in databases (Tu ¨bingen Map of Zebrafish Genome,
http:??wwwmap.tuebingen.mpg.de; Massachusetts General
Hospital?Cardiovascular Research Center Zebrafish Server,
http:??zebrafish.mgh.harvard.edu; The Zebrafish Information
Genome Project Initiative, http:??188.8.131.52?zonrhmap-
per?maps.htm). To identify single-nucleotide polymorphisms
and ultimately the mutation, genomic DNA was amplified by the
PCR with primers designed with information from the Zebrafish
Genome-Sequencing Project (www.ensembl.org?danio?rerio?
index.html) and sequenced. This source also provided mapping
information on expressed sequence tags and predicted introns
and exons with the GENESCAN program.
Morpholino Injection. Morpholinos (Gene Tools, Philomath, OR)
were dissolved in Danieau solution [58 mM NaCl, 0.7 mM KCl,
0.6 mM Ca(NO3)2, 0.4 mM MgCl2, and 5 mM Hepes at pH 7.2]
and injected in 1- to 5-ng doses into one- or two-cell embryos
with 0.05% (wt?vol) phenol red as a marker. Presented in a 5?-3?
orientation, the morpholino sequences were: atrophin2, TCCT-
TGGAGGCTGTAAACACAAATT; atrophin2 mismatch con-
trol, TCgTTGcAGcCTcTAAACAgAAATT; fgf8, GAGTCT-
CATGTTTATAGCCTCAGTA; sef, CGCAAGTCTCCGTGA-
CCCAGCCATT; and sef mismatch control, CGgAAcTCTCg-
Asai et al.PNAS ?
June 13, 2006 ?
vol. 103 ?
no. 24 ?
In Situ Hybridization. Whole-mount in situ hybridization was Download full-text
performed on homozygous albino larvae (29). Digoxigenin-
labeled RNA probes were synthesized according to the instruc-
tions of the manufacturer (Ambion, Austin, TX). The atrophin2
probe included ?1,000 bp of 3? coding sequence, whereas the
fgf8 probe included the entire coding sequence. The cDNA
fragment amplified with the primers TCCTGAAAAAGTA-
GATTCAGGATTG and ATGTTAAAGGACTCTCACAT-
TCAGG served as a sef probe. Hybridization and washing were
performed at 65°C for all probes.
Quantitative PCR. After total RNA had been prepared from pools
of 20 embryos (RNeasy; Qiagen, Valencia, CA), cDNA was
synthesized from 1 ?g of total RNA (SuperScript First Strand
Synthesis; Invitrogen). Real-time quantitative PCR was con-
ducted (ABI Prism 7700; Applied Biosystems) with a melting
temperature of 60°C. Reactions were conducted in a total
volume of 15 ?l with 7.5 ?l of iTaq SYBR Green Supermix with
ROX (Bio-Rad), 0.5–50 ng of cDNA, and 100 nM each of the
forward and reverse primers. Each reaction was run in triplicate;
dilution series of cDNA (1:0, 1:10, and 1:100) were used to
generate calibration curves for each primer set. The forward and
reverse primer sequences were: ?-actin, AAGATCATTGC-
CCCACCTGAG and CCGTTTAGAAGCATTTGCGGT; fgf8,
AATCGCAGAGCACAGACCCTT and GGCTTTCGGTC-
CTCTCCTTTT; sef, GGCGCTCGTTATATGTTGCAA and
TTTATTAGGCAGCGGCGGA. The triplicate Ct values for
each sample were averaged, and the results were normalized by
the amount of product from an endogenous control message
(?-actin). The data presented represent a comparison of results
from embryos injected with the morpholino against atrophin2
with those from a control group injected with water. Each
experiment was conducted three times.
We thank Ms. P. Espitia and Mr. A. Afolalu for outstanding fish
husbandry, Dr. M. Vologodskaia and Ms. Y. Castellanos for technical
assistance, Dr. H. Lo ´pez-Schier for help with immunolabeling, and Dr.
S. Koshida for providing the in situ hybridization probe for zebrafish fgf8.
Dr. B. B. Riley, Dr. H. Takeda, and the members of our research group
provided valuable comments on the manuscript. This investigation was
supported by National Institutes of Health Grant DC00241. Y.A. is an
Associate and A.J.H. is an Investigator of the Howard Hughes Medical
1. Hudspeth, A. J. (1998) Nature 341, 397–404.
2. Barald, K. F. & Kelley, M. W. (2004) Development (Cambridge, U.K.) 131,
3. Haddon, C. & Lewis, J. (1996) J. Comp. Neurol. 365, 113–128.
4. Whitfield, T. T., Riley, B. B., Chiang, M.-Y. & Phillips, B. (2002) Dev. Dyn. 223,
5. Haddon, C. & Lewis, J. (1991) Development (Cambridge, U.K.) 112, 541–550.
6. Kimmel, C. B., Patterson, J. & Kimmel, R. O. (1974) Dev. Physiol. 7, 47–60.
7. Nicolson, T., Ru ¨sch, A., Friedrich, R. W., Granato, M., Ruppersberg, J. P. &
Nu ¨sslein-Volhard, C. (1998) Neuron 20, 271–283.
8. Leger, S. & Brand, M. (2002) Mech. Dev. 119, 91–108.
9. Walshe, J. & Mason, I. (2003) Dev. Biol. 264, 522–536.
10. Ghysen, A. & Dambly-Chaudiere, C. (2004) Curr. Opin. Neurobiol. 14, 67–73.
11. Lo ´pez-Schier, H., Starr, C. J., Kappler, J. A., Kollmar, R. & Hudspeth, A. J.
(2004) Dev. Cell 7, 401–412.
12. Yanagisawa, H., Bundo, M., Miyashima, T., Okamura-Ohno, Y., Todokoro, K.,
Tokunaga, K. & Yamada, M. (2000) Hum. Mol. Genet. 9, 1433–1442.
13. Zoltewicz, J. S., Stewart, N. J., Leung, R. & Peterson, A. S. (2004) Development
(Cambridge, U.K.) 131, 3–14.
14. Erkner, A., Roure, A., Charroux, B., Delaage, M., Holway, N., Core, N., Vola,
C., Angelats, C., Pages, F., Fasano, L. & Kerridge, S. (2002) Development
(Cambridge, U.K.) 129, 1119–1129.
15. Zhang, S., Xu, L., Lee, J. & Xu, T. (2002) Cell 108, 45–56.
16. Furthauer, M., Reifers, F., Brand, M., Thisse, B. & Thisse, C. (2001) Devel-
opment (Cambridge, U.K.) 128, 2175–2186.
17. Furthauer, M., Lin, W., Ang, S. L., Thisse, B. & Thisse, C. (2002) Nat. Cell Biol.
18. Tsang, M., Friesel, R., Kudoh, T. & Dawid, I. B. (2002) Nat. Cell Biol. 4,
19. Riccomagno, M. M., Martinu, L., Mulheisen, M., Wu, D. K. & Epstein, D. J.
(2002) Genes Dev. 16, 2365–2378.
20. Liu, W., Li, G., Chien, J. S., Raft, S., Zhang, H., Chiang, C. & Frenz, D. A.
(2002) Dev. Biol. 248, 240–250.
21. Hammond, K. L., Loynes, H. E., Folarin, A. A., Smith, J. & Whitfield, T. T.
(2003) Development (Cambridge, U.K.) 130, 1403–1417.
22. Riccomagno, M. M., Takada, S. & Epstein, D. J. (2005) Genes Dev. 19,
23. Barresi, M. J., Stickney, H. L. & Devoto, S. H. (2000) Development (Cambridge,
U.K.) 127, 2189–2199.
24. Wang, L., Rajan, H., Pitman, J. L., McKeown, M. & Tsai, C.-C. (2006) Genes
Dev. 20, 525–530.
25. Westerfield, M. (1995) The Zebrafish Book (University of Oregon Press,
26. Kappler, J. A., Starr, C. J., Chan, D. K., Kollmar, R. & Hudspeth, A. J. (2004)
Proc. Natl. Acad. Sci. USA 101, 13056–13061.
27. Starr, C. J., Kappler, J. A., Chan, D. K., Kollmar, R. & Hudspeth, A. J. (2004)
Proc. Natl. Acad. Sci. USA 101, 2572–2577.
28. Zhang, J., Talbot, W. S. & Schier, A. F. (1998) Cell 92, 214–251.
29. Prince, V. E., Moens, C. B., Kimmel, C. B. & Ho, R. K. (1998) Development
(Cambridge, U.K.) 125, 393–406.
www.pnas.org?cgi?doi?10.1073?pnas.0603453103Asai et al.