Disease Models & Mechanisms 4, 000-000 (2011) doi:10.1242/dmm.006429
RESEARCH ARTICLE
Harmonin (Ush1c) is required in zebrafish Müller glial
cells for photoreceptor synaptic development and
function
Jennifer B. Phillips1,*, Bernardo Blanco-Sanchez1,*, Jennifer J. Lentz2,3, Alexandra Tallafuss1, Kornnika Khanobdee4,‡,
Srirangan Sampath2, Zachary G. Jacobs1, Philip F. Han1, Monalisa Mishra4, David S. Williams4,5, Bronya J. Keats2,§,
Philip Washbourne1 and Monte Westerfield1,¶
Disease Models & Mechanisms DMM
SUMMARY
Usher syndrome is the most prevalent cause of hereditary deaf-blindness, characterized by congenital sensorineural hearing impairment and
progressive photoreceptor degeneration beginning in childhood or adolescence. Diagnosis and management of this disease are complex, and the
molecular changes underlying sensory cell impairment remain poorly understood. Here we characterize two zebrafish models for a severe form of
Usher syndrome, Usher syndrome type 1C (USH1C): one model is a mutant with a newly identified ush1c nonsense mutation, and the other is a
morpholino knockdown of ush1c. Both have defects in hearing, balance and visual function from the first week of life. Histological analyses reveal
specific defects in sensory cell structure that are consistent with these behavioral phenotypes and could implicate Müller glia in the retinal pathology
of Usher syndrome. This study shows that visual defects associated with loss of ush1c function in zebrafish can be detected from the onset of vision,
and thus could be applicable to early diagnosis for USH1C patients.
INTRODUCTION
Usher syndrome (USH) is characterized by recessively inherited
deaf-blindness. Previous estimates of the worldwide prevalence of
USH ranged from 1/17,000 to 1/25,000 (Boughman et al., 1983;
Rosenberg et al., 1997). However, a r-ecent study of deaf and hardof-hearing children suggested a prevalence of 1/6000 (Kimberling
et al., 2010). USH1 is the most severe of the three clinical subtypes,
with profound congenital deafness and onset of vision loss usually
before age 10. Vestibular dysfunction is also typical in USH1 cases.
USH2, the most common clinical subtype worldwide, is
characterized by moderate to severe congenital hearing impairment
and the onset of visual defects usually in the second decade of life.
No balance problems are associated with USH2. USH3 is rare in
most populations, and distinct from the other clinical subtypes in
that both hearing and vision impairments are progressive, and
vestibular dysfunction is variable (for reviews, see Saihan et al., 2009;
Yan and Liu, 2010). Cochlear implants are indicated in cases of
1
Institute of Neuroscience, University of Oregon, Eugene, OR 97403-1254, USA
Department of Genetics, LSU Health Sciences Center, New Orleans, LA 70112, USA
3
Neuroscience Center, LSU Health Sciences Center, New Orleans, LA 70112, USA
4
Department of Pharmacology and Neuroscience, UCSD, La Jolla, CA 92093-0636,
USA
5
Jules Stein Eye Institute, UCLA, Los Angeles, CA 90095-7000, USA
*These authors contributed equally to this work
‡
Present address: School of Biology, Institute of Science, Suranaree University of
Technology, Muang, Nakhon Ratchasima 30000, Thailand
§
Present address: Research School of Biology, Australian National University,
Canberra, ACT 2601, Australia
¶
Author for correspondence ([email protected])
2
Received 30 July 2010; Accepted 23 May 2011
© 2011. Published by The Company of Biologists Ltd
This is an Open Access article distributed under the terms of the Creative Commons Attribution
Non-Commercial Share Alike License (http://creativecommons.org/licenses/by-nc-sa/3.0), which
permits unrestricted non-commercial use, distribution and reproduction in any medium provided
that the original work is properly cited and all further distributions of the work or adaptation are
subject to the same Creative Commons License terms.
Disease Models & Mechanisms
profound deafness seen in USH1, and hearing aids can compensate
for some of the hearing loss in USH2 and USH3, but there is
currently no treatment to counter the vision loss resulting from
retinitis pigmentosa. Early diagnosis is important for genetic
counseling and preparation for late-onset blindness.
At least 11 different loci are implicated in USH. The nine
identified USH genes encode structurally and functionally
heterogeneous proteins (Keats and Savas, 2004; Reiners et al., 2006;
Ebermann et al., 2007), but most possess protein interaction
domains that suggest the ability to form molecular complexes with
one another. Biochemical studies have confirmed these binding
propensities (Reiners et al., 2005; Pan et al., 2009). Colocalization
of the USH proteins in subcellular domains of auditory hair cells
and retinal cells, along with evidence of mislocalization of USH
proteins in various USH mutant backgrounds, suggest that at least
some USH proteins directly interact with one another (Adato et
al., 2005; Reiners et al., 2005; Lefevre et al., 2008; Bahloul et al.,
2010; Yang et al., 2010; Caberlotto et al., 2011).
Previous studies of USH proteins have concentrated on their
potential functions in the region of the connecting cilium of the
photoreceptor (Liu et al., 2007; van Wijk et al., 2006; Maerker et al.,
2008; Yang et al., 2010) and the stereocilia of hair cells (Adato et al.,
2005; Delprat et al., 2005; Seiler et al., 2005; Söllner et al., 2004;
Kikkawa et al., 2005; Prosser et al., 2008; Lefevre et al., 2008). In the
ear, several studies have provided evidence for colocalization of USH
proteins in the tip and ankle-linking regions of the stereocilia and at
hair cell ribbon synapses (Mburu et al., 2003; Siemens et al., 2004;
Reiners et al., 2005; Michalski et al., 2007; Kazmierczak et al., 2007;
Lefevre et al., 2008). Most of the USH proteins have been shown to
colocalize in the retina at the connecting cilium and/or at the
photoreceptor synapse (Liu et al., 1997; Reiners et al., 2006; Liu et
al., 2007; Overlack et al., 2008; Yang et al., 2010). In addition to these
regions of colocalization, many of the USH proteins are found
individually at discrete subcellular locations (Hasson et al., 1995;
1
DMM
RESEARCH ARTICLE
Reiners et al., 2006; Overlack et al., 2008; Williams et al., 2009),
suggesting that at least some have functions independent of the
proposed complex. In hair cells, USH proteins are thought to
stabilize the growth and orientation of stereocilia during development
(for reviews, see Brown et al., 2008; Yan and Liu, 2010), whereas USH
proteins in the region of the connecting cilium are proposed to
contribute to the trafficking of molecular cargo from the
photoreceptor inner segment to the outer segment (Maerker et al.,
2008; Liu et al., 2007). USH proteins have also been immunolocalized
at the ribbon synapses of these sensory cells, but their potential
functions in this region are unknown.
The USH1C gene encodes the PDZ-domain-containing protein
harmonin (Verpy et al., 2000), which has been proposed to function
as a key scaffolding molecule to which most other USH proteins
can bind (Reiners et al., 2005). Patients with mutations in USH1C
exhibit the classic USH1 pathology, with profound congenital
hearing impairment, vestibular dysfunction, and clinically
appreciable retinal degeneration in childhood or early adolescence.
USH1C mutations resulting in non-syndromic deafness have also
been reported (Ouyang et al., 2002). The deaf circler mouse
(Johnson et al., 2003), harboring an in-frame deletion in the
murine Ush1c gene, has provided a valuable model for the
mechanosensory hair cell defects that cause deafness in USH1C
patients, but does not exhibit a retinal phenotype (Williams et al.,
2009). Similarly, the Ush1c knockout mouse exhibits profound
deafness without notable retinal defects (Tian et al., 2010).
Interestingly, the c.216G>A mouse model of USH1C
(Ush1c216AA), a knock-in of the human exon 3 USH1C c.216G>A
mutation (plus surrounding sequence) cloned from an Acadian
USH1C patient (Lentz et al., 2007), shows both the wellcharacterized auditory defects of the other USH1C mouse models,
as well as visual defects comparable to the human disease. In
addition to the disorganized stereocilia and hair cell degeneration
observed in the cochleae of USH1C c.216G>A mice, reduced visual
function was noted after 1 month of age and progressive loss of
rod photoreceptors was observable after 6.5 months of age (Lentz
et al., 2010). These intriguing differences in retinal pathology
between the Ush1c knockout and the USH1C c.216G>A knock-in
could indicate differences between the functions of endogenous
mouse and human harmonin in the retina.
In vitro assays have demonstrated that many of the known USH
proteins bind preferentially to one or more of the three PDZ domains
in harmonin, hence its depiction as a central scaffold upon which
an USH protein complex can form (Adato et al., 2005). Although the
subcellular localization of harmonin and other USH proteins in hair
cells is compatible with this hypothesis, there have been conflicting
reports of localization of other USH proteins at the photoreceptor
synapse, where harmonin localization is undisputed (Reiners et al.,
2005; Liu et al., 2007; Yang et al., 2010). Moreover, harmonin is
notably absent from the connecting cilium region (Reiners et al., 2005;
Williams et al., 2009), where the presence of most other USH proteins
has been well documented (Liu et al., 2007; Maerker et al., 2008;
Yang et al., 2010). Thus, the normal cellular functions of harmonin,
the types of complexes it forms with other USH proteins in vivo, and
the early, pre-degenerative manifestations of the retinal pathology
in USH1C remain mysterious.
Previously, zebrafish mutants for myo7a (ush1b), cdh23 (ush1d)
and pcdh15 (ush1f) have been described as exhibiting severe
2
ush1c in zebrafish visual function
swimming and balance defects consistent with the auditory and
vestibular component of the human disease (Ernest et al., 2000;
Söllner et al., 2004; Seiler et al., 2005). Although expression in the
retina has been reported for each of these genes, only the
morpholino knockdown of pcdh15b has been reported to have an
effect on visual function (Seiler et al., 2005). Photoreceptor
organization is severely affected in pcdh15b morphants, and
electroretinogram (ERG) traces are notably attenuated. Here, we
describe two zebrafish models for USH1C that exhibit a strong
USH-like phenotype at early developmental stages. Young fish with
depleted ush1c function have penetrant and specific visual defects
as well as compromised hair cell development, resulting in hearing
and balance impairment. Detailed analyses of these phenotypes
have confirmed a requirement for harmonin in zebrafish hair cell
morphogenesis, and have implicated Müller glial cells in supporting
harmonin-mediated ribbon synapse stability and function. USH has
traditionally not been considered a developmental disease of vision.
The results of this study, however, show that visual defects can be
detected from the onset of vision in zebrafish with depleted ush1c
function, thus providing a rationale for examination of Müller cell
function in USH1C patients.
RESULTS
ush1c expression in zebrafish inner ear sensory patches, the lateral
line and retina
We used the human harmonin protein sequence in tBLASTn
searches to identify a single ortholog of USH1C in the zebrafish
genome (Fig. 1A). We then used a radiation hybrid panel (Hukriede
et al., 1999) to position this gene on chromosome 25. Previous
studies of the mouse and human genes have indicated that the
mammalian genes express three splice forms, which vary slightly
in structure and subcellular localization (Reiners et al., 2003;
Reiners et al., 2005; Williams et al., 2009). The most abundant
zebrafish transcript found in whole larvae or adult retinas
corresponded to isoform A, containing 21 exons that encode a 548
amino acid protein with three PDZ domains and a coiled-coil
domain (Fig. 1A). Zebrafish harmonin isoform A was found to be
structurally similar to the corresponding human isoform, with 70%
amino acid identity overall and highest similarity in the PDZ
domains and the N-terminal region (Pan et al., 2009). A
hydrophobicity plot of the human and zebrafish proteins revealed
a high level of conservation, even where residue substitutions occur
(Fig. 1A). Using reverse transcriptase (RT)-PCR and RACE primers
within and between regions encoding the second and third PDZ
domains, we identified unique transcripts that might correspond
to the B or C isoforms of the gene, although no variant encoding
a second coiled-coil or PST domain, as would be predicted for an
isoform B transcript, was identified. All transcripts other than that
of isoform A were in low abundance such that we were unable to
obtain sequence definitively identifying them as belonging to a
particular isoform class. The probes generated from these partial
sequences showed weak, generalized signal in all tissues and time
points analyzed (data not shown).
cRNA probes directed against either the unique sequence of
isoform A or the 5⬘ region predicted to be common to all splice
forms yielded identical expression patterns in larval tissues. RNA
expression was first detected in the ear at 27 hours post-fertilization
(hpf ), when the first hair cells are specified (data not shown), and
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ush1c in zebrafish visual function
RESEARCH ARTICLE
Fig. 1. Zebrafish ush1c is expressed in retina and the inner ear. (A)The predominant splice form of ush1c corresponds to mammalian isoform A, with 21 exons
that produce a 1.6 kb mRNA. A hydrophobicity plot of zebrafish (red) and human (blue) isoform A protein sequence shows conserved residue polarity. % amino
acid identity with human protein is indicated below the plot. The positions of the ush1cfh293 nonsense mutation (red arrowhead), the MO target (green
arrowhead) and the region recognized by the zebrafish harmonin antibody (red bar) are indicated. (B)At 5 dpf, ush1c transcripts are abundant in the inner retina
(black arrow), the sensory patches of the ear (white arrows), the intestinal tract (i) and the swim bladder (sb). Fainter expression in the brain (b) and trunk muscles
(m) is also observed. (C)High-magnification view of a 5 dpf whole-mount zebrafish ear showing ush1c expression in all sensory patches (ac, lc, pc: anterior, lateral
and posterior cristae, respectively; am: anterior macula) and an adjacent head neuromast (n). (D)ush1c transcripts are restricted to cells in the inner nuclear layer
in 5 dpf sectioned retina. (E)Central section of an adult zebrafish retina showing continued strong expression in the inner retina; expression is detected in the
outer nuclear layer as well. onl, outer nuclear layer; inl, inner nuclear layer; gcl, ganglion cell layer. Scale bars: 100m (B); 20m (C); 50m (D,E).
hair cell expression persisted through all later larval stages examined
[5 days post-fertilization (dpf ) shown in Fig. 1B,C]. We also
observed strong expression in the neuromasts of the lateral line
(Fig. 1C). This neuromast expression was consistent with previous
reports of the zebrafish USH gene orthologs myo7a, cdh23 and
pcdh15a (Ernest et al., 2000; Söllner et al., 2004; Seiler et al., 2005).
Retinal expression was present beginning at 48 hpf, when retinal
lamination is complete and cell specification is underway. By 72
hpf, strong expression was observed in a subset of cells in the inner
nuclear layer, which persisted through all larval stages examined
(5 dpf, shown in Fig. 1D). In situ hybridization of adult retinal tissue
using all described probes showed persistent, strong expression in
the inner nuclear layer as well as expression in the outer nuclear
layer (Fig. 1E). We also observed strong expression of ush1c in the
gut and swim bladder, and weaker expression in the brain and trunk
muscles (Fig. 1B), consistent with reported expression in human
tissues (Verpy et al., 2000; Hirai et al., 2004).
Mutation or morpholino knockdown of ush1c depletes harmonin
protein in zebrafish larvae
We obtained a nonsense mutation in ush1c from the Zebrafish
TILLING Consortium by screening for mutations in the first PDZdomain-encoding region of the gene (PDZ1). We recovered a line,
ush1cfh293, that harbors a nonsense mutation in the fifth exon, which
encodes the C-terminal portion of the PDZ1 domain (red
arrowhead, Fig. 1A; supplementary material Fig. S1).
We also designed morpholino oligonucleotides to knock down
the function of ush1c: one to block splicing at the exon-2–intronDisease Models & Mechanisms
2 boundary (MO1; green arrowhead, Fig. 1A) and one that
recognizes the translation initiation site (MO2). We obtained
identical phenotypes with both morpholinos, but found that the
exon-2-targeting morpholino (MO1) provided more consistent
results; thus, MO1 was used for all morpholino experiments and
all uses of ‘morpholino’, ‘MO’ or ‘morphant’ refer to MO1. Using
RT-PCR analysis of MO-injected embryos and larvae, we identified
an aberrant splice form in which exon 2 is skipped, producing outof-frame splicing of exons 1 and 3 that results in an early
termination codon (supplementary material Fig. S1). This altered
splice form predominates for the first 4 dpf, and persists through
6 dpf, although recovery of the normal splice form is evident by 5
dpf (Fig. 2A).
We obtained a polyclonal rabbit antibody raised against the first
100 amino acids of zebrafish harmonin (red bar, Fig. 1A), a region
common to all predicted isoforms, and probed protein extracts on
a western blot to test antibody specificity and to determine the
extent of protein depletion in morphant and mutant larvae (Fig.
2B). In wild-type controls, we observed a single band in the 3 dpf
sample, which was slightly smaller than the corresponding bands
in the 4 dpf and 5 dpf samples. In addition to this band, which, at
approximately 60 kDa, corresponded to the predicted size of
zebrafish harmonin isoform A, we noted progressively increasing
numbers of smaller bands in the 4 dpf and 5 dpf controls. Consistent
with the time course of morpholino efficacy revealed by RT-PCR
(Fig. 2A), strong knockdown of harmonin protein levels was
observed at 3-4 dpf in ush1c morphants. Moreover, although the
upper band was restored nearly to wild-type levels in the 5 dpf
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RESEARCH ARTICLE
ush1c in zebrafish visual function
Fig. 2. Mutation and morpholino knockdown of ush1c produce behavioral defects. (A)RT-PCR of control and splice-blocking morpholino-injected embryos
from 1 to 5 dpf reveals a smaller splice form lacking exon 2 that predominates for the first 4 days of development and persists through the fifth day. (B)Western
blots from extracts of control + ush1c MO-injected embryos from 3 to 5 dpf (left) and ush1cfh293 homozygous mutant embryos alongside wild-type siblings (right)
confirm morpholino efficacy and antibody specificity. Samples from 3 dpf to 5 dpf control larvae probed with the zebrafish harmonin antibody show a 60 kDa
band consistent with the predicted size of isoform A. Additional smaller bands are visible faintly in the 4 dpf control and are more pronounced and abundant in
the 5 dpf control. The band corresponding to the isoform A variant is undetectable in 3 dpf and 4 dpf morphant samples. The major band reappears at 5 dpf at a
slightly smaller weight, and there is a reduced number of smaller bands in this sample. In extracts from 5 dpf ush1cfh293 homozygous mutants, no bands are
observed. Extracts from wild-type siblings from the same clutch show the characteristic banding pattern of the 5 dpf uninjected controls, allowing for the slightly
longer running time for this gel. A pan-actin antibody was used as a loading control. (C)Behavioral assays of control, mutant and morpholino-injected larvae
show (top) impaired startle response and balance, and (bottom) reduced optokinetic response, tracking an average of 50% or fewer of the dark stripes (average
targets tracked for wild type: 11.5±1.27; ush1c mutants: 6.13±2.78, P<0.0001; ush1c MO: 5.13±2.85, P<0.0001; n30 for each group).
sample, the size was slightly reduced, consistent with the control
band from the 3 dpf time point (Fig. 2B). These temporal changes
in size and abundance of the major band, recapitulated by the
recovering morphant protein extracts, might reflect a period of
post-translational modification.
We performed RT-PCR experiments on zebrafish cDNA from
multiple developmental stages and discovered five distinct
expressed sequence tags (ESTs), but no complete sequences of splice
variants other than isoform A. Thus, we cannot definitively identify
the smaller bands on the western blots, but postulate that they
might be additional isoforms or products of protein degradation
of isoform A, the largest running band. Similar, smaller bands were
detected by western analysis of mouse harmonin (Williams et al.,
2009). We did not observe any bands larger than ~60 kDa at any
larval time point examined. This, combined with the RT-PCR and
RACE results described above, suggests that isoform B splice
variants are not active through 5 dpf.
No bands were detected in protein samples from ush1c mutant
larvae at any time point examined (5 dpf shown alongside wildtype sibling in Fig. 2B). However, the ush1c nonsense mutation
in the fifth exon could potentially produce a truncated protein
that would be recognized by the antibody. We investigated RNA
levels in ush1c mutants using a riboprobe against the first six
4
exons of ush1c. We observed reduced probe signal in all ush1cexpressing tissues examined (ear and gut tissue shown in
supplementary material Fig. S2), suggesting that this mutant
transcript is unstable.
Reduced harmonin function produces defects in vestibular,
auditory and visual function
At 5 dpf, wild-type fish exhibited rapid evasive swimming behavior
when startled by a tap on the Petri dish. We tested the startle reflex
of 5 dpf offspring from natural crosses between ush1c–/+
heterozygous carriers (n952 larvae) and found that 25% (n236)
of these larvae had abnormal responses (Fig. 2C, top). Within this
group, 83% did not exhibit any evasive swimming behavior, and
those that did respond swam in circles. Subsequent genotyping
revealed that 100% of the larvae exhibiting these defects were
homozygous ush1cfh293 mutants.
Free-swimming 5 dpf ush1c morphant larvae exhibited vestibular
defects identical to ush1c mutants, with only 30% of the larvae
responding to tapping on the Petri dish, and all responders swimming
in circles (n400). These defects are similar to those reported for
other zebrafish USH gene mutations or knockdowns (Ernest et al.,
2000; Söllner et al., 2004; Seiler et al., 2005) and indicate reduced
function of both the auditory and vestibular systems.
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ush1c in zebrafish visual function
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Fig. 3. Harmonin is required in otic sensory patches for normal bundle development. (A-C)Phalloidin-stained F-actin (green) and anti-zebrafish harmonin
antibody labeling (red) in 80-hpf wild-type ears show localization of harmonin relative to the stereocilia in the five otic sensory patches (ac, anterior crista; am,
anterior macula; lc, lateral crista; pc, posterior crista; pm, posterior macula) and in the neuromasts (n). The anti-harmonin antibody signal is reduced in sensory
patches and in neuromasts in ush1c mutants (B) and ush1c-MO-injected larvae (C). (D-L)High-magnification view of phalloidin-stained hair bundles in the
anterior and posterior maculae of 5 dpf larvae. Hair bundle number is reduced and organization of the existing bundles is disrupted in mutant (G-I) and
morphant (J-L) anterior (G,J) and posterior (H,K) maculae, compared with the anterior and posterior maculae of wild-type controls (D and E, respectively). Closeup images of hair bundles of the anterior maculae (F,I,L) show bent and splayed bundles in mutants and morphants. For mutant analysis, anterior macular wildtype average54.6±3.5 bundles; n8; ush1c mutant average33±3.5; n7; P<0.0001. Posterior macula wild-type average68.2±2.6 bundles; n9; ush1c mutant
average40.5±4.2; n10; P<0.0001. For the morpholino experiment, anterior macula control average41.8±4.8 bundles; MO average21.8±3.8; n5 for each
group; P<0.0001. Posterior macula control average71.8±4.6 bundles; MO average48.2±6.9; n5 for each; P<0.0001, 5 dpf. Scale bars: 50m (A-C); 10m (D-L).
To assess visual function in mutant and morphant animals, we
used the optokinetic response (OKR) assay. At 5 dpf, ush1c mutants
and MO-treated larvae produced significantly fewer saccades,
tracking only 40-50% of the targets compared with wild type (Fig.
2C, bottom).
Loss of harmonin results in morphological defects in
mechanosensory hair cells
To gain insights into the cause of the hearing and balance defects
in harmonin-deficient larvae, we used immunofluorescence to
examine the mechanosensory hair cells of the otic sensory patches.
Harmonin localized to sensory patches and neuromasts by 80 hpf
(Fig. 3A). We detected no harmonin signal at 80 hpf in ush1c
mutants (Fig. 3B) or morphants (Fig. 3C), confirming the strong
knockdown or depletion indicated by the western blot results. Using
phalloidin to visualize stereocilia, we examined individual sensory
patches. At 5 dpf, wild-type hair bundles had a characteristic shape
and orientation, projecting more or less orthogonally from the plane
of each cuticular plate and appearing tightly tapered at their tips
(Fig. 3D-F). All otic sensory patches in the mutant larvae displayed
reduced numbers of hair bundles, as well as defects in stereocilia
organization at 5 dpf (Fig. 3G-I). We observed 40% fewer hair
bundles than in wild-type siblings in the anterior and posterior
maculae, and existing stereocilia were splayed and bent, with a
notable lack of tapering of the hair bundles. In morpholinoDisease Models & Mechanisms
injected animals, we found a reduced number of hair bundles, with
48% fewer bundles in the anterior maculae and 33% fewer hair
bundles in the posterior maculae of 5 dpf morphants compared
with controls. Stereocilia were also disorganized, although to a
lesser extent than we observed in the mutant. Bundles appeared
bent and disoriented relative to the plane of the cuticular plate, but
the splaying or absence of tapering at the tips was less pronounced
than in ush1c mutants (Fig. 3J-L).
We next investigated the subcellular localization of harmonin
in the anterior maculae of 5 dpf larvae (Fig. 4). In wild-type hair
cells, harmonin localized to both stereocilia and kinocilia as well
as within cell bodies and at synapses (Fig. 4A-G). In ush1c
mutants (Fig. 4H-J), we detected reduced levels of harmonin in
stereocilia and kinocilia. Protein was also detected within cell
bodies, but the distribution appeared abnormal compared with
controls. Harmonin levels in hair cells of morphant larvae (Fig.
4K-M) were markedly diminished at 5 dpf, although some signal
could be detected apical to the hair bundle in a position consistent
with kinocilia localization. When we injected embryos from
ush1c+/– heterozygous parents with the ush1c morpholino, raised
the larvae to 5 dpf and analyzed harmonin staining in sensory
patches, residual staining was undetectable in injected
genotypically ush1c–/– animals, suggesting that a truncated form
of harmonin is produced in the mutants. This truncated protein
might be too low in abundance or too unstable to be detected in
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ush1c in zebrafish visual function
Disease Models & Mechanisms DMM
Fig. 4. Subcellular localization of harmonin in mechanosensory
hair cells. Co-labeling of wild-type 5 dpf anterior macula hair cells
(A-D) shows anti-zebrafish harmonin localization (red) in the hair
bundles (green), kinocilia (arrowheads), cell bodies and at synaptic
boutons (arrow in C and D). Incubation conditions favoring hair
bundle visualization (E-P) show strong colocalization of harmonin
within the bundle structure as well as in the cuticular plate in 5 dpf
wild types (E-G). Kinocilia labeling can be seen above the bundle
region (asterisks). Enhanced signal detection in mutant hair cells (HJ) shows localization persisting in the kinocilia and in existing
bundle structures. Kinocilia are more visible in these panels owing to
the reduced bundle architecture in the mutant. Protein distribution
seems globular and disorganized in the cell bodies (arrowheads).
Hair cells of ush1c MO larvae (K-M) show strong knockdown of
harmonin, with localization persisting only in kinocilia at this stage,
and no signal observed within hair bundles. Injecting the
morpholino into the ush1c–/– mutants reduces subcellular
localization of harmonin to undetectable levels in hair cells (N-P).
Scale bars: 10m.
a western blot, but was detectable in fixed tissue using enhanced
signal detection protocols.
To investigate cell degeneration or reduced cell proliferation as
possible causes for the observed reduction in hair bundle number,
we stained larvae at stages between 1 and 5 dpf with TUNEL to
label dying cells or BrdU to label dividing cells. The results in both
cases were indistinguishable between morphants and controls (not
shown), and were consistent with previously reported normal cell
death levels in the ear (Bever and Fekete, 1999). Mutant cell
proliferation was also unaffected at these stages, as were cell death
levels through 4 dpf. In 5 dpf mutants, we noted a small but
significant increase in TUNEL-labeled cells for both the anterior
macula (0.1±0.32 for wild type vs 1.2±1.03 for mutant; n10 for
each group; P0.008) and posterior macula (0.1±0.32 for wild type
vs 1.3±0.95 for mutant; n10 for each; P0.003). Although
statistically significant, the numbers of dead cells were insufficient
to account for the extent of the reduction in bundles. To confirm
that hair cells were being specified properly in mutant larvae, we
used a probe to the hair-cell-specification factor atoh1b and noted
no differences in labeling of sensory patches in wild type compared
with mutant (data not shown). Thus, we conclude that the defects
in hair bundle formation occur post-mitotically, consistent with
previously reported studies in mammals (Lefevre et al., 2008), and
that increased cell death, reduced cell proliferation, or impaired
6
cell fate specification cannot be the primary causes of bundle loss.
Together, these results confirm a requirement for harmonin in hair
bundle development. Additionally, the milder stereocilia
disorganization phenotype in ush1c morphants observed at 5 dpf,
in conjunction with the partial recovery of harmonin localization
at 5 dpf observed in the region of the kinocilia, suggests that
harmonin also plays a role in maintaining hair bundle integrity.
Harmonin localizes to Müller glial cells in the retina
Retinal architecture appeared unaffected in ush1c mutants and
morphants compared with wild type (Fig. 5A,F,K; supplementary
material Fig. S3); however, because visual function defects were
observed, we also examined harmonin protein localization in
retinas. We observed abundant protein levels specifically in the cell
bodies and processes of Müller glia at all larval stages examined
(Fig. 5B-E; supplementary material Fig. S4), consistent with the
restricted inner retinal expression noted by in situ hybridization
with our ush1c riboprobes at 5 dpf (Fig. 1D). We did not observe
specific photoreceptor labeling with this antibody between 5 and
8 dpf.
We examined harmonin antibody labeling in wild-type, mutant
and morphant retinas at 5 dpf. Harmonin protein levels in the
retinas of ush1c–/– homozygous mutants were markedly reduced,
although Müller cells were present and appeared morphologically
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ush1c in zebrafish visual function
RESEARCH ARTICLE
Disease Models & Mechanisms DMM
Fig. 5. Harmonin localization in the larval retina is
restricted to Müller glia. Semi-thin plastic sections stained
with toluidine blue show consistent size, lamination and
organization of 5 dpf wild-type, mutant and morphant
retinas (A,F,K). 16-m larval retina sections (B,G,L) show
colocalization of the Müller cell marker glutamine
synthetase (GS; green) and anti-zebrafish harmonin (red) in
wild type (B). Both antibodies label Müller cell processes
throughout the retina. Anti-zebrafish harmonin labeling is
reduced in mutant (G) and morphant (L) Müller cells. Müller
cell number and morphology is unaffected. Nonspecific
signal associated with photoreceptor outer segments is
observed in all panels at this magnification. See
supplementary material Fig. S4 for comparison. Highmagnification images of anti-zebrafish harmonin and GS
labeling in Müller cell processes at the OPL (large arrow) and
OLM (small arrow) of the 5 dpf retina are shown (wild type,
C-E; mutant, H-J; morphant, M-O). Scale bars: 50m
(A,B,F,G,K,L); 20m (C-E,H-J,M-O).
normal (Fig. 5G-J). Similarly, after morpholino injection, we
observed a strong reduction of harmonin labeling in Müller cells
through 5 dpf (Fig. 5L-O). We noted that the duration of protein
depletion was prolonged in the eye, compared with our results from
morphant hair cells, in which we observed rebounding harmonin
localization in some kinocilia by 5 dpf (Fig. 3F). We therefore
examined harmonin labeling in 6 dpf morphants, at which time
we observed increased harmonin labeling in Müller cells of the
peripheral retina, adjacent to the ciliary marginal zone
(supplementary material Fig. S4).
Owing to the apparent presence of ush1c transcript in adult
photoreceptors (Fig. 1E), we evaluated the subcellular localization
of harmonin in 6 month adult retinas (Fig. 6A-F). We continued
to see strong harmonin localization within Müller cells in the adult
retina, extending past the outer limiting membrane (OLM) and into
the glial processes in the subapical region between photoreceptor
cell bodies (Fig. 6, asterisks). In some adult retinas we also observed
what might be a low level of protein localization at the
photoreceptor synapses (Fig. 6A, arrow). Occasionally, signal was
detected in photoreceptor inner or outer segments at a lower level
and with inconsistent distribution compared with the robust
staining observed in Müller glia (Fig. 6D, arrow), suggesting that
this signal was due to nonspecific labeling or autofluorescence,
although we were unable to confirm this with our current
immunofluorescence methods.
Because Müller cell localization of harmonin has not been
reported previously, we used antibodies against murine (Fig. 6GI) and human (Fig. 6J-L) harmonin to label retinas from postnatal
day 10-11 mouse pups. In addition to the previously described
labeling in photoreceptors, we also observed harmonin
immunoreactivity in Müller glial cell bodies and processes
extending throughout the retina. The anti-human harmonin
antibody (Fig. 6J-L) also cross-reacted in 5 dpf zebrafish retinas,
labeling Müller cells, but not photoreceptors (data not shown).
Given that the observed localization of harmonin in murine Müller
cells differs from published work (Reiners et al., 2005), further
investigation is needed, but our results are consistent with the
Disease Models & Mechanisms
possibility of a phylogenetically conserved role for harmonin in
retinal glia.
Harmonin was recently detected in the postsynaptic processes
of murine bipolar cells and horizontal cells, using immunoelectron
microscopy (Williams et al., 2009). With our immunofluorescence
protocols, we found no evidence for harmonin localization within
cell bodies of bipolar or horizontal cells at any time points examined
in zebrafish (Fig. 5; Fig. 6A,C). Owing to the strong harmonin
labeling in Müller glial processes, we cannot determine whether
synaptic processes of horizontal or bipolar cells might also be
positive for harmonin in the outer plexiform layer (OPL).
Given the retinal degeneration experienced by USH1C patients,
we examined retinal cell death in zebrafish ush1c mutants and
morphants at 5 and 8 dpf, using an anti-active caspase-3 antibody.
Death rates by retinal cell layer were as follows (n10 for all groups;
see also supplementary material Fig. S4): in the photoreceptor layer,
wild-type animals at 5 dpf had 0.20±0.41 labeled cells compared
with 0.31±0.60 in morphant retinas (P0.5) and 3.09±1.3 in mutants
(P<0.0001); in the inner nuclear layer, no labeled cells were observed
in wild type, compared with 1.86±1.23 in morphants (P<0.0001)
and 1.88±0.73 in mutants (P<0.0001); in the ganglion cell layer,
0.062±0.25 cells were labeled in controls, compared with 0.25±0.45
in morphants (P0.16) and 0.09±0.30 in mutants (P0.79). Noting
the small but significant elevation of inner nuclear layer (INL) cell
death in 5 dpf mutants and morphants, we co-labeled sectioned 5
dpf retinas with caspase-3 and glutamine synthetase to determine
what proportion of the dying cells were Müller glia (n10 for each
group). In ush1c–/– animals, exhibiting an average of 1.88 labeled
cells per eye, 67% of the cells labeled with caspase-3 were also
positive for glutamine synthetase. Similarly, 61% of caspase-labeled
cells were co-labeled with glutamine synthetase in ush1c morphant
retinas, in which an average of 1.85 cells in the INL were labeled.
In both cases, the remaining labeled cells could not be positively
identified as Müller cells, based on either their position within the
nuclear layer or by labeling with the Müller cell marker. Although
our current experiments do not rule out the possibility that a small
subset (<0.7 cells per eye) of caspase-positive cells in the INL might
7
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RESEARCH ARTICLE
Fig. 6. Harmonin localizes in Müller cells through all zebrafish life stages
and is evolutionarily conserved. (A-F) Localization of anti-zebrafish
harmonin (red) in Müller cells persists in adult zebrafish retinas. Some low level
signal is sometimes observed in photoreceptor synapses (arrow in A) and in
photoreceptor outer segments (arrow in D). The strongest signal within the
photoreceptor layer is associated with Müller cell processes projecting into the
subapical region (asterisks). Nonspecific fluorescence of photoreceptor outer
segments at this stage can be compared with panel H in supplementary
material Fig. S4. (G-L)Anti-mouse harmonin (SDI; G,I) and anti-human
harmonin (Santa Cruz; J,L) detect localization in photoreceptors and Müller
cells [glutamine synthetase (GS) is labeled in green; harmonin antibodies in
red] in retinas from P10-P11 mouse pups. The outer nuclear layer (onl), inner
nuclear layer (inl) and ganglion cell layer (gcl) are labeled in merged panels.
Scale bars: 50m (A-C,G-L); 20m (D-F).
be second order neurons, it is also possible that these cells were
apoptotic Müller glia that no longer produced glutamine synthetase.
At 8 dpf, we noted increased photoreceptor cell death in the
ush1c mutant retinas, compared with wild-type animals (wild type:
0 labeled photoreceptors; mutant: 5.33±1.32 labeled
photoreceptors; P<0.0001; see supplementary material Fig. S4). No
statistically significant differences were observed between mutant
and wild-type siblings in either the inner nuclear or ganglion cell
layers at this stage, nor were there significant differences seen in
the labeling of morpholino and control animals in any cell layers
at 8 dpf.
8
ush1c in zebrafish visual function
Harmonin is required for photoreceptor synaptic organization and
function
The specific depletion of harmonin from Müller cells in larval
mutant and morphant retinas provided an opportunity to study
the role of harmonin in this cell type. Because the morphants
presented no significant photoreceptor cell death at 5 dpf despite
strong protein knockdown, we used morphants for the following
analyses. We began by examining synaptic integrity in morphant
larvae at 5 dpf, using antibodies against the pre- and postsynaptic
proteins SV2 and GluR2/3, respectively. In control retinas, these
markers exhibited an organized, parallel and largely colocalized
alignment within the OPL (Fig. 7A). In ush1c morphants, both
proteins were present in the OPL, but they were remarkably
misaligned and less colocalized, indicating a disruption in synaptic
organization (Fig. 7B).
To investigate this phenotype further, we examined the
ultrastructure of cone pedicles in morphant larvae via transmission
electron microscopy. We examined cone pedicles from retinas of
two uninjected controls at 6 dpf in a quantitative comparison with
cone pedicle ultrastructure from retinas of four stage-matched
ush1c MO larvae. We found that cone pedicles in uninjected
controls (Fig. 7C,E) contained an average of 1.667±0.299 ribbons
per pedicle (n36 pedicles) and a large number of mature-appearing
ribbon synapses, with 55% of the synaptic ribbon profiles anchored
by an arciform density (ribbons with arciform density per
pedicle1.08±0.238); 85% of these membrane-anchored ribbons
were associated with postsynaptic processes from horizontal and
bipolar cells in a triad (triads per pedicle0.97±0.128). In morphant
retinas (Fig. 7D,F), the average number of ribbons per pedicle was
consistent with controls (1.659±0.607; n44 pedicles), but only 38%
of the ribbon profiles appeared anchored by arciform densities
(ribbons with arciform densities per pedicle0.636±0.647;
P0.0002). Synaptic ribbons unanchored by an arciform density
seemed to ‘float’ in the pedicle, and postsynaptic processes, when
present, frequently appeared distant from these floating ribbons
(Fig. 7D,F). As in the controls, most (83%) anchored ribbons in
morphants were incorporated into triads (triads per
pedicle0.523±0.412), suggesting a specific defect in ribbon
association with the membrane. To investigate whether the
observed defects could be attributed to a developmental delay, we
also examined sections of ribbon synapses in cone pedicles of two
wild-type and three morphant retinas at 8 dpf. The number of
ribbons observed per pedicle (1.34±0.56; n41) was significantly
reduced in 8 dpf morphants compared with controls (1.70±0.22;
n43; P0.0002). As seen in the 5 dpf analysis, significantly fewer
ribbons in the 8 dpf morphants (52%; n55 ribbons) compared with
controls (82%; n73 ribbons; P<0.0001) appeared to be anchored
to the membrane. The percentage of anchored ribbons associated
with triads was comparable between morphants and controls (68%
and 65%, respectively), again suggesting a primary defect in the
formation of arciform densities. Retinal morphology was otherwise
unaffected, and significant retinal cell death was not observed in
8 dpf morphants in any retinal cell layers by anti-active caspase-3
labeling. Thus, we conclude that temporary depletion of harmonin
from Müller glia during retinal development has a lasting, adverse
impact on maturation of cone ribbon synapses.
To determine the effect of the morphological defect on synaptic
transmission, we recorded ERGs from 5 dpf morphant larvae (n6)
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DMM
Fig. 7. Synaptic maturation is impaired in ush1c
morphants. (A,B)Anti-SV2 (red) and -GluR2/3 (green)
antibodies mark the pre-and postsynaptic regions of the
OPL, respectively. Tightly associated localization of these
factors is apparent in the control retina (A), and is notably
disrupted in the morphant retina (B) at 5 dpf. (C-F)Electron
micrographs of cone pedicles in 6 dpf control and morphant
larvae. In control retinas (C,E), the presence of multiple triads
(boxed areas), consisting of a synaptic ribbon (r), arciform
density (arrow) and postsynaptic processes from bipolar (b)
and horizontal (h) cells, indicate normal synaptic maturation.
An enlargement of a triad is shown in E. In morphant retinas
(D,F), fewer triads are present (boxed areas in D; enlarged in
panel F), and floating ribbons (r) and distant postsynaptic
processes (h) are observed in cone pedicles. Scale bars:
20m (A,B); 1m (C,D); 500 nm (E,F).
and compared them with uninjected larvae (n7). Rod
photoreceptors are detectable molecularly by 3 dpf and
morphologically by 5 dpf. However, physiological analyses
demonstrate that rods are not functional until 15 dpf (Morris and
Fadool, 2005). ERGs performed at 5 dpf show a cone dominant
response in both light-adapted and dark-adapted conditions (Bilotta
et al., 2001; Seeliger at el., 2002), but dark-adapted retinas at this
stage are less susceptible to bleaching and exhibit a more sensitive
ON response than in light-adapted conditions (Makhankov et al.,
2004). Thus, we chose to perform our experiments under scotopic
conditions, in which we observed that the b-wave amplitude, an
indicator of synaptic transmission, was markedly reduced in the
MO-injected animals (Fig. 8A) compared with controls. The awave, which is primarily a read out of photoreceptor membrane
depolarization as a result of phototransduction, is often quenched
by conditions conducive to generating b-wave amplitudes and is
therefore not observable on the same trace. Thus, we isolated the
a-wave pharmacologically with the synaptic transmission inhibitors
L-AP4 and TBOA (Trombley and Westbrook, 1992; Huang and
Bordey, 2004; Wong et al., 2005), whereupon we observed a slight
reduction in isolated a-wave amplitude compared with controls
(P0.01), but less of an effect overall than that observed from the
b-wave experiments (P0.0007; Fig. 8B). Plotting a-wave and b-wave
amplitudes against each other shows that the b-wave to a-wave ratio
is significantly diminished in the morphant group (P<0.0007; Fig.
8C). These data, taken together with the disorganization of synaptic
proteins in the OPL (Fig. 7B), indicate that postsynaptic activity is
reduced relative to the level of the phototransduction response to
light stimulus (Takada et al., 2008). Although we observed a very
small increase in the number of caspase-positive cells in the INL,
the majority of which were identified as Müller glia, we would not
expect the loss of so few cells to account for such a large reduction
Disease Models & Mechanisms
in the b-wave amplitude. Thus, our data suggest that loss of
harmonin in Müller cells results in decreased function of the
photoreceptor ribbon synapses.
DISCUSSION
In this study, we describe the requirement for harmonin in zebrafish
mechanosensory hair cell and retinal development, providing the
first evidence that USH proteins function in photoreceptor synaptic
transmission and ribbon synapse development. Our studies also
implicate Müller glial cells as contributors to the retinal pathology
of USH.
The localization of harmonin in hair cells and lateral line
neuromasts indicates a phylogenetically conserved role in
mechanosensory cells, and our studies of the zebrafish inner ear
reveal defects in hair bundle organization and development,
consistent with the stereocilia elongation and organization defects
described in Ush1c mutant mice (Lefevre et al., 2008). Our results
indicate that, in zebrafish, the predominant splice variant and
protein isoform present during larval development is harmonin
isoform A. The severe disruption of hair bundle development seen
in ush1c mutant and morphant zebrafish provides an intriguing
contrast to the absence of hair bundle defects reported in some
colonies of dfcr mice, which harbor a deletion affecting harmonin
isoform B (Grillet et al., 2009).
The presence of the USH protein harmonin in Müller glia from
an early time point in retinal development and the specific
defects in synaptic structure and function in ush1c morphants
prior to detectable ush1c transcript or protein synthesis in
photoreceptors suggest a non-cell-autonomous role for harmonin
in facilitating synaptic maturation in cone pedicles. Müller glial
cells have established roles in the maintenance of retinal neurons
via their regulation of glutamate neurotransmitter levels and their
9
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ush1c in zebrafish visual function
DMM
Fig. 8. Deficits in synaptic function of photoreceptors in ush1c morphants. (A,B)Representative larval zebrafish ERG recordings showing b-wave amplitudes
(A) or a-wave amplitudes (B) of uninjected control (black) and MO-injected (gray) larvae at unattenuated light intensity. (A)b-wave amplitudes are significantly
reduced in morphant animals. (B)The a-wave of these same subjects was isolated by treatment with L-AP4 and TBOA to block synaptic transmission. a-waves are
only slightly attenuated in morphant animals compared with controls. Light stimulus duration is 800 mseconds. (C)Comparison of the relative changes of b- and
a-wave amplitudes in morphant and control larvae obtained at 0, –1 and –2 log units of light intensity relative to unattenuated light intensity. The decreased bwave:a-wave ratio indicates a specific synaptic defect. Steeper slopes of the trend lines (dashed lines) represent reduced synaptic efficacy.
expression of neurotrophic factors (Fields and Stevens-Graham,
2002; Newman, 2004; de Melo Reis et al., 2008). Although a role
for Müller glia in retinal synaptogenesis has been suggested by
several studies (Georges et al., 2006; Diaz et al., 2007), little is
known about how this putative function might be accomplished.
In the brain, glutamate receptor function has been shown to be
important for synaptogenesis (Han and Stevens, 2009; Kakegawa
et al., 2009), so glutamate uptake by Müller cells might similarly
be important for synaptic development of photoreceptors.
Interestingly, a recent zebrafish study (Williams et al., 2010) used
a Müller-cell-specific line to ablate individual glia and observe
effects on synaptogenesis and the behavior of neighboring glia.
Their data indicated that localized loss of Müller cells did not
affect the maturation of cone synapses at 5 dpf. These data are
consistent with our findings, because they argue against the small
and transient levels of Müller cell death being the cause of the
reduced synaptic function and structural defects observed, and
suggest instead that harmonin in Müller cells has a paracrine effect
on cone synapse stability, such that the sustained absence of this
protein results in the observed dysfunction and progressive
photoreceptor cell death.
The cone synapse phenotype of ush1c morphants is similar to
that of the zebrafish nrc mutant, in which the synaptojanin gene
is disrupted (Allwardt et al., 2001; Van Epps et al., 2004). The nrc
mutant has a diminished b-wave and defects in photoreceptor
synapse formation, with floating ribbons and unassociated
postsynaptic processes in cone pedicles. Synaptojanin has been
shown to be required for cytoskeletal organization, endocytosis,
Ca2+ signaling and post-synaptic glutamate transport (Sakisaka
et al., 1997; Schuske et al., 2003; Gong and DeCamilli, 2008).
Although no link between synaptojanin and USH has been
proposed previously, nrc mutants have swimming and balance
defects in addition to blindness (Van Epps et al., 2004), similar
to the USH-like phenotype we observe in ush1c mutants and
morphants. More recently, a report describing three new mutant
alleles of zebrafish synaptojanin, isolated by screening for
vestibular defects, illustrates synaptic transmission defects in the
hair cells (Trapani et al., 2009).
Given the distinct, penetrant defects in visual function and
photoreceptor synapse formation in harmonin-depleted larvae, we
10
were surprised by the lack of harmonin localization in larval
photoreceptors. It is puzzling that there would be no requirement
for harmonin function in larval zebrafish photoreceptors, especially
in light of the abundant antibody signal detected in the
photoreceptors of young mice. An as-yet-undetected duplicate
ush1c gene in zebrafish could account for the apparent protein
localization discrepancies between mammals and zebrafish.
However, our searches of the zebrafish genome reveal only one
locus for ush1c, and we find only one copy of this gene in the
Tetraodon, Takifugu and Threespine stickleback genome
assemblies, suggesting that only a single copy of the ush1c gene
has been retained in the teleost lineage.
Another possible explanation for the absence of harmonin in
larval photoreceptors is the presence of ohnologous genes for two
other PDZ-domain-containing proteins: Cip98 and Pdzd7.
Alignments with the human CIP98 (whirlin) protein sequence,
causative of USH2D, uncovered two orthologs in the zebrafish
genome assembly. Both of these transcripts, cip98a and cip98b, are
expressed in photoreceptors from early larval time points (J.B.P.,
unpublished); thus, the presence of an additional PDZ protein with
similar binding affinities to harmonin (van Wijk et al., 2006) might
be able to compensate for the absence of ush1c expression in these
cells. However, given that the N-terminal regions of harmonin and
Cip98 are not functionally interchangeable (Pan et al., 2009), any
functional substitution of harmonin by Cip98 in the photoreceptor
must not rely on protein interactions in the N-terminal region.
Although PDZD7 is not currently recognized as a monogenic cause
of human USH, it has been shown to act as a modifier of USH gene
function and to interact physically with USH proteins. Both
zebrafish genes, pdzd7a and pdzd7b, are expressed in
photoreceptors at larval stages (Ebermann et al., 2010).
Our ERG recordings from ush1c morphant larvae reveal a
specific reduction in b-wave amplitude in the absence of significant
retinal degeneration, which might provide an explanation for the
early cellular dysfunctions that precede retinitis pigmentosa in
USH1C patients. A retrospective ERG analysis of USH1 patients
as young as 17 months revealed abnormal waveforms prior to
clinically detectable retinitis pigmentosa (Flores-Guevara et al.,
2009), although other studies of USH1 patients in which
photoreceptor loss was already clinically evident revealed no
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Disease Models & Mechanisms DMM
ush1c in zebrafish visual function
synaptic dysfunction in the remaining viable photoreceptors
(Jacobson et al., 2008; Williams et al., 2009). Notably, the
Ush1c216AA knock-in mouse (Lentz et al., 2010) exhibits an
attenuated ERG as early as 1 month of age, prior to any histologically
appreciable cell loss. However, in contrast to our results with larval
zebrafish, both a- and b-waves were attenuated in Ush1c216AA
mice. It could be that, in human manifestations of USH1C, the later
consequence of diminished harmonin levels in photoreceptors is
so severe that it obscures comparatively subtle early defects that
might result from impaired synaptic activity.
Our data also indicate that sustained absence of harmonin in
Müller cells, as seen in ush1c mutants, results in progressive retinal
degeneration. It is not clear whether this cell death results from
synaptic defects or from some other impairment of Müller cell
function, but the non-cell-autonomous nature of this degeneration
is notable. Clinical analyses of the retinal pathology of USH patients
might provide insight into the link between Müller cell function
and retinal integrity. Ophthalmic examinations of USH1B patients
(who have mutations in the MYO7A gene) using high-resolution
optical coherence tomography (Jacobson et al., 2009) showed that
morphological changes in the OLM (the apical terminus of Müller
cell processes) marked a transitional zone between regions in which
photoreceptors were diseased or depleted and morphologically
normal regions, suggesting that such alterations in OLM integrity
might precede photoreceptor degeneration. Our understanding of
the role that the OLM plays in maintaining photoreceptor integrity
is incomplete; however, numerous junction proteins, including
members of the Crumbs complex, have been shown to localize to
the OLM (for reviews, see Gosens et al., 2008; Omri et al., 2010)
and, in particular, to the subapical region, perhaps contributing to
junctions within a semipermeable barrier. Mutations in Crumbs
orthologs in mammals cause progressive retinal degeneration and
the retinal pathology reveals diminished contacts between
photoreceptors and the OLM (Mehalow et al., 2003; van de Pavert,
2004). USH proteins have not been reported in the OLM, or in
Müller cells in general, before this study, although whirlin (USH2D)
has been linked biochemically to the Crumbs pathway (Gosens et
al., 2007).
Although the extent to which Müller cells might contribute to
the USH retinal pathology remains to be investigated, our
histological analyses of both zebrafish and mouse retinas, and our
functional studies in ush1c morphants and mutants, suggest that
additional investigation of Müller cell function is required to
understand the complex etiology of USH. As gene replacement
therapies are developed for USH patients, it will be important to
know whether retinal cell types in addition to photoreceptors
should be targeted for optimal rescue of the vision defects. The
ability to isolate Müller-cell-associated pathology in our zebrafish
USH1C model will allow further investigation of the roles of
Müller cells as well as investigations of USH protein function at
the OLM.
In summary, we have characterized the hearing and vision defects
in a zebrafish ush1c mutant and morpholino knockdown. Our
findings support a phylogenetically conserved developmental role
for harmonin isoform A in hair bundle organization, implicate
Müller cells as contributors to retinal dysfunction and
photoreceptor cell death, and demonstrate that visual defects can
be detected from the onset of vision in these zebrafish models.
Disease Models & Mechanisms
RESEARCH ARTICLE
METHODS
Animal strains and maintenance
All zebrafish studies were conducted with Oregon AB wild-type
larvae or ush1cfh293 mutants in the AB background. Animals were
raised in a 10-hour dark and 14-hour light cycle, and maintained
as described (Westerfield, 2007). Owing to the swimming and
balance defects exhibited by ush1c mutants and morphants, all
experimental and control larvae for experiments occurring up to
8 dpf were raised without food so as not to give normal swimmers
a nutritional advantage over their swimming-impaired siblings.
Animals were staged according to the standard series (Kimmel et
al., 1995), or by hpf or dpf. All animal-use protocols were IACUC
approved.
Cloning of zebrafish ush1c cDNA and sequencing
tBLASTn queries with the human harmonin protein sequence were
used to identify the zebrafish sequence, and primers based on this
sequence were used to amplify the full-length open reading frame
of isoform A and partial sequence from additional isoforms from
whole larvae or adult retinal cDNA (see supplementary material Table
S1 for primer sequences). PCR products were purified using gel
electrophoresis and sequenced on an ABI 3100 using the fluorescent
dideoxy terminator method. The gene is cataloged in NCBI under
accession number NC_007136. All novel transcripts were submitted
to the NCBI dbEST, with accession numbers HO243997-HO244001
and HS587020. Gene and protein names follow the zebrafish
nomenclature guidelines (https://wiki.zfin.org/display/general/
ZFIN+Zebrafish+Nomenclature+Guidelines).
Genetic mapping of zebrafish ush1c
The zebrafish ush1c locus was mapped by PCR using the LN54
collection of radiation hybrids (obtained from Marc Ekker,
University of Ottawa), which contain the zebrafish AB9 cell DNA
and mouse B78 cell DNA (Hukriede et al., 1999).
ush1c mutant discovery
Labeled primers (supplementary material Table S1) to PCR amplify
a 500 bp region surrounding exon 5 of ush1c were generated and
used to screen 8448 genomes in a TILLING screen as described
(Moens et al., 2008). Sperm from the identified carrier sample were
thawed and used to fertilize wild-type AB eggs. Adult F1 carriers of
the mutation were identified by fin-clip genotyping, and identified
heterozygotes were crossed to produce homozygous offspring.
Morpholino injections
Antisense morpholinos (GeneTools, Philomath, OR) were injected
into one-cell-stage embryos as described (Nasciveus and Ekker,
2000). Embryos were injected with 1.8-2.0 nM of ush1c MO1
targeted to the splice donor site of exon 2 (5⬘-TAGAATTGAGACTTACTGATGGTAC-3⬘), or 0.6-1.0 nM of ush1c MO2
targeted to the translation start site (5⬘-TTTTCAGTCCGTTGGTCGTCTTGCT-3⬘).
RNA isolation and RT-PCR
Total RNA was isolated from euthanized larvae at 24 hpf to 7 dpf
using Trizol (Invitrogen, Carlsbad, CA). Reverse transcription was
performed according to manufacturer instructions using the
Superscript Reverse Transcriptase II kit (Invitrogen).
11
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Disease Models & Mechanisms DMM
Western blot
Embryos were homogenized in 10 mM Tris-HCl, pH 7.4, 150 mM
NaCl, 1 mM EDTA, 0.1% Triton X-100, 0.1% SDS and 5 mg/ml
protease inhibitor cocktail (Pierce, Rockford, IL). Samples were run
on an 8% acrylamide gel and transferred to nitrocellulose
membranes with Nupage reagents (Invitrogen). Membranes were
washed in PBS twice, each for 15 minutes, incubated in blocking
solution (3% NFDM in PBS) for 1 hour at room temperature, then
in primary antibodies [rabbit anti-zebrafish harmonin (SDIX,
Newark, DE) 1:300; mouse monoclonal pan-actin (Millipore,
Billerica, MD) 1:5000] in blocking solution overnight at 4°C.
Membranes were rinsed three times for 5 minutes each in PBS,
incubated in donkey anti-rabbit HRP (Jackson, West Grove, PA)
for 1 hour at room temperature, then rinsed three times for 5
minutes each in PBS. HRP was detected using the ECL plus
detection system (Pierce). To detect actin, membranes were
rehydrated and incubated in donkey anti-mouse HRP for 1 hour,
rinsed three times for 5 minutes each in PBS, and labeled with the
ECL plus detection system (Pierce).
In situ hybridization and immunohistochemistry
In situ hybridization on whole and sectioned tissue and antibody
labeling on sectioned tissue was performed as described (Jensen
et al., 2001; Ebermann et al., 2010). For imaging of subcellular
protein localization in sectioned retinas, incubation with primary
antibodies and a goat anti-mouse Alexa-Fluor-488 to visualize the
glutamine synthetase antibody was followed by incubation with an
anti-rabbit biotinylated antibody diluted in block (10% NGS, 2%
BSA in PBS-T) overnight at 4°C. Sections were then rinsed four
times for 5 minutes each in PBS and signal was detected with the
Vectastain ABC Elite kit (Vector Laboratories, Burlingame, CA)
and TSA kit (Perkin-Elmer, Waltham, MA). For whole-mount
imaging of stereocilia, 5 dpf larvae were fixed in ‘BT’ fix (4% PFA,
0.15 mM CaCl2, 4% sucrose in PBS) for 48 hours, rinsed four times
for 5 minutes each in PBS + 0.01% Triton X-100, and then
permeabilized with 2.5% Triton X-100 in PBS at room temperature
until otoliths dissolved. Larvae were rinsed four times for 5 minutes
each in PBS + 0.01% Triton X-100, blocked for 2 hours at room
temperature in 10% NGS, 10% BSA in PBS + 0.01% Triton X-100,
then incubated with Alexa-Fluor-488–phalloidin diluted in block
for 24 hours at 4°C. Following four 5-minute PBS + 0.01% Triton
X-100 washes, larvae were blocked using the Avidin/Biotin blocking
kit (Molecular Probes, Carlsbad, CA), and incubated in antibody
solutions overnight at 4°C. Larvae were rinsed four times for 30
minutes each in PBS-Triton, followed by signal detection with the
Vectastain ABC Elite kit (Vector) and TSA (Perkin-Elmer) kit. After
rinsing four times for 5 minutes each in PBS-Triton, larvae were
immersed in Vectashield (Vector) and mounted in a lateral
orientation on chambered slides. Images were captured on a Zeiss
LSM5 confocal or BioRad Radiance 2100 confocal microscope. The
following antibodies and concentrations were used in this study:
rabbit anti-harmonin (SDIX) antibody generated against aa1-100
of zebrafish harmonin, 1:200 for sections, 1:1000 for whole mount;
rabbit anti-harmonin (SDIX) antibody generated against aa306-363
of mouse harmonin, 1:200; goat anti-harmonin (Santa Cruz
Biotechnology, Santa Cruz, CA) against the N-terminus of the
human protein, 1:40; rabbit anti-harmonin (Sigma-Aldrich, St
Louis, MO) against aa42-118 of human harmonin, 1:40; mouse anti12
ush1c in zebrafish visual function
glutamine synthetase (Millipore), 1:500; mouse zpr1 (Zebrafish
International Resource Center, Eugene, OR), 1:200; mouse anti-SV2
(DSHB), 1:2000; rabbit anti-GluR2/3 (Millipore), 1:50; rabbit antiactive caspase-3 (BD Biosciences, San Jose, CA), 1:200; mouse antiacetylated tubulin (Sigma), 1:500. Goat anti-mouse, chicken antigoat and goat anti-rabbit Alexa-Fluor-488, -568 and -633 secondary
antibodies (Molecular Probes) were used at 1:300 and the antirabbit biotinylated antibody was used at 1:500. Alexa-Fluor488–phalloidin (Invitrogen) was diluted to a final concentration of
2.5 M.
Electron microscopy
Tissue was prepared as described previously (Schmitt and Dowling,
1999). Ultrathin sections were obtained from Epon-embedded
tissue, stained with uranyl acetate and lead citrate, and examined
by conventional transmission electron microscopy.
Optokinetic response
To measure optokinetic response, we used a modification of the
method of Brokerhoff (Brokerhoff, 2006). Larvae were placed in a
small Petri dish containing 3% methylcellulose and mounted on a
stationary platform surrounded by a rotating drum of 8 cm in
diameter. The drum interior displays ten alternating black and white
stripes, each subtending an angle of 28°. The larvae were oriented
in an upright position, illuminated with fiber optic lights and viewed
through a dissecting microscope positioned over the drum, which
rotated at 5 rpm clockwise and then counter clockwise, for 30
seconds in each direction. Eye movements were recorded during
this time period, after which the larvae were washed out of the
methylcellulose and returned to embryo medium.
ERG recordings
ERG recordings (Makhankov et al., 2004) were made to examine
retinal function of morphants (morpholino-injected larvae)
compared with control larvae. 5 dpf larvae were incubated in
Ringer’s solution for at least 1 hour prior to the experiment and
dark-adapted for a minimum of 30 minutes. Under dim red
illumination, larvae anesthetized with tricaine (Sigma Aldrich) and
paralyzed with 300 M pancuronium bromide (Sigma Aldrich)
were placed on a piece of sponge so that the pupil of one eye was
on axis with the stimulus light beam. A recording electrode was
placed on the cornea to measure the change in voltage induced by
stimulation. The reference electrode was a AgCl-coated silver wire
inserted into the sponge. The recording electrode was connected
via a AgCl-coated silver wire to a low noise recording amplifier
(Axopatch 200A with a CV201A head stage), and data were
acquired and stored digitally (Digidata 1200 interface and pClamp7
and pClamp8 software; recording hardware and software from
Axon Instruments). Signals were filtered at 2 kHz.
Stimulation
Recordings were performed in a Faraday cage housing a Zeiss UEM
microscope, with a 4⫻ objective lens used for visualizing the eye
and delivering light stimuli. An ultra bright white LED array (Atlas
series high brightness module NT-42D0-0426 from Lamina
Lighting, with an LED driver circuit locally designed and
constructed by the technical support group) was used to present
the stimulus. Stimulus durations were 800 mseconds. The light
dmm.biologists.org
ush1c in zebrafish visual function
TRANSLATIONAL IMPACT
Clinical issue
Usher syndrome (USH) is a recessively inherited disease of combined deafness
and blindness. Hearing impairment is usually apparent from birth, whereas
vision loss is slow and progressive, beginning in the first or second decade of
life. Cochlear implants and hearing aids can compensate for some of the
hearing deficits, but there is currently no treatment to counter the vision loss,
which is a form of retinitis pigmentosa. Early diagnosis is important for genetic
counseling and preparation for late onset blindness.
There is a great deal of clinical variability among patients with USH, and
nine genes that can cause this disease when mutated have been identified to
date. These genes encode a wide variety of proteins that have complex and
dynamic localization patterns within the cells of the inner ear and the retina
that are affected in USH. Discovering more about how these mutations cause
deafness and blindness at the cellular and molecular level will enhance our
understanding of the etiology of this disease, and contribute to the
development of effective treatments.
DMM
Results
In this paper, the authors describe a zebrafish model for Usher type 1C
(USH1C), a severe form of USH in humans that is characterized by profound
congenital deafness, balance problems and a relatively early onset of
progressive vision loss. The authors disrupt the normal function of the ush1c
gene by using morpholino oligonucleotides to interfere with mRNA splicing. In
addition, they identify an ush1c zebrafish mutant by screening DNA from
individual mutagenized fish to find a lesion in the ush1c gene. Both
morpholino-injected animals and ush1c mutants have distinct defects in
hearing, balance and vision that are apparent in the first days of life. The
authors examine the sensory cells involved in these behaviors –
mechanosensory hair cells and retinal cells – and discover defects in their
development and function. In the ear, the structural portion of the
mechanosensory hair cells that are important for intercepting sound waves
does not form properly. Surprisingly, the authors discover that, in the retina,
the ush1c gene functions in Müller glia rather than in photoreceptors, the cell
type that has been reported to express Ush1c in mammals. Subsequently, they
show that this gene is also expressed in mammalian Müller glia. Importantly,
although zebrafish ush1c is not active in photoreceptors, loss of ush1c function
in Müller cells leads to compromised photoreceptor function and cell death.
Implications and future directions
Further insights into the role of ush1c in the development and function of
sensory cells are vital to providing early diagnosis and intervention to forestall
the devastating effects of this disease. This work contributes to the
understanding of USH1C pathology by providing information about the
specific molecules involved in the formation of stereocilia in mechanosensory
hair cells, and by identifying a new retinal cell type through which ush1c
functions to facilitate and maintain synaptic connections between
photoreceptor cells and interneurons of the retina, namely Müller glia. USH
has traditionally not been considered a developmental disorder of vision. The
results of this study, however, show that visual defects in the zebrafish model
of USH1C can be detected from the onset of vision, and thus might have
clinical relevance for the early diagnosis of USH1C in humans.
intensity of the unattenuated beam was 1500 W/cm2. Appropriate
neutral density filters were inserted into the light beam to achieve
accurate light stimulus intensities over a 5 log10 unit range. Three
to five responses per intensity level were averaged and filtered for
60 Hz noise.
Isolation of the a-wave component
Directly after recording the b-wave (in Ringer’s solution),
pharmacological compounds were applied externally to the eye
using an Eppendorf pipette. The compounds used in this study were
Disease Models & Mechanisms
RESEARCH ARTICLE
L-(+)-2-amino-4-phosphonobutyric acid (L-AP4; 30-300 M), DLthreo-b-Benzyloxyaspartic acid (DL-TBOA; 15 M) and Ringer’s
solution with 5% DMSO as control. After 15-30 minutes, the awave traces were recorded.
Data analysis
The amplitudes of the b-waves were measured from the bottom of
the a-wave to the peak of the b-wave; the a-wave was measured
from the baseline to the peak of the a-wave. We used larvae that
had a consistent response to a test flash delivered (±10%) at the
beginning and the end of the session. Unhealthy or damaged wildtype larvae that showed a decreased ERG or changed waveform at
the end of the experiment compared with the beginning were not
included in the analysis. For statistics, we used a linear mixed model
approach and a Student’s t-test.
ACKNOWLEDGEMENTS
The authors thank David Raible, Kelly Owens and John Constable for help with
experimental design and reagents, Tom Titus and the Cecilia Moens laboratory for
their work on the TILLING screen, and Jeremy Wegner for technical assistance.
Support was provided by NIH EY07042, DC004186, DC010447 and HD22486, and
by the Foundation Fighting Blindness. J.B.P. was supported by an American Heart
Association Postdoctoral Fellowship. D.S.W. is a Jules and Doris Stein RPB Professor.
COMPETING INTERESTS
The authors declare that they do not have any competing or financial interests.
AUTHOR CONTRIBUTIONS
J.B.P. designed and performed experiments, analyzed data and wrote the
manuscript with input from the other authors. B.B.-S. designed and performed
experiments and analyzed data. J.J.L. designed and performed experiments and
contributed experimental reagents. A.T., K.K. and M.M. performed experiments and
analyzed data. S.S., Z.G.J. and P.F.H. performed experiments. D.S.W., P.W., B.J.K. and
M.W. conceived and designed the experiments.
SUPPLEMENTARY MATERIAL
Supplementary material for this article is available at
http://dmm.biologists.org/lookup/suppl/doi:10.1242/dmm.006429/-/DC1
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