harmonin (ush1c) is required in zebrafish müller …ion.uoregon.edu/sites/default/files/westerfield...

INTRODUCTION

Usher syndrome (USH) is characterized by recessively inheriteddeaf-blindness. Previous estimates of the worldwide prevalence ofUSH 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 hard-of-hearing children suggested a prevalence of 1/6000 (Kimberlinget al., 2010). USH1 is the most severe of the three clinical subtypes,with profound congenital deafness and onset of vision loss usuallybefore age 10. Vestibular dysfunction is also typical in USH1 cases.USH2, the most common clinical subtype worldwide, ischaracterized by moderate to severe congenital hearing impairmentand the onset of visual defects usually in the second decade of life.No balance problems are associated with USH2. USH3 is rare inmost populations, and distinct from the other clinical subtypes inthat both hearing and vision impairments are progressive, andvestibular dysfunction is variable (for reviews, see Saihan et al., 2009;Yan and Liu, 2010). Cochlear implants are indicated in cases of

profound deafness seen in USH1, and hearing aids can compensatefor some of the hearing loss in USH2 and USH3, but there iscurrently no treatment to counter the vision loss resulting fromretinitis pigmentosa. Early diagnosis is important for geneticcounseling and preparation for late-onset blindness.

At least 11 different loci are implicated in USH. The nineidentified USH genes encode structurally and functionallyheterogeneous proteins (Keats and Savas, 2004; Reiners et al., 2006;Ebermann et al., 2007), but most possess protein interactiondomains that suggest the ability to form molecular complexes withone another. Biochemical studies have confirmed these bindingpropensities (Reiners et al., 2005; Pan et al., 2009). Colocalizationof the USH proteins in subcellular domains of auditory hair cellsand retinal cells, along with evidence of mislocalization of USHproteins in various USH mutant backgrounds, suggest that at leastsome USH proteins directly interact with one another (Adato etal., 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 theirpotential functions in the region of the connecting cilium of thephotoreceptor (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 theear, several studies have provided evidence for colocalization of USHproteins in the tip and ankle-linking regions of the stereocilia and athair 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 tocolocalize in the retina at the connecting cilium and/or at thephotoreceptor synapse (Liu et al., 1997; Reiners et al., 2006; Liu etal., 2007; Overlack et al., 2008; Yang et al., 2010). In addition to theseregions of colocalization, many of the USH proteins are foundindividually at discrete subcellular locations (Hasson et al., 1995;

Disease Models & Mechanisms 1

Disease Models & Mechanisms 4, 000-000 (2011) doi:10.1242/dmm.006429

1Institute of Neuroscience, University of Oregon, Eugene, OR 97403-1254, USA2Department of Genetics, LSU Health Sciences Center, New Orleans, LA 70112, USA3Neuroscience Center, LSU Health Sciences Center, New Orleans, LA 70112, USA4Department of Pharmacology and Neuroscience, UCSD, La Jolla, CA 92093-0636,USA5Jules 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 ofTechnology, Muang, Nakhon Ratchasima 30000, Thailand§Present address: Research School of Biology, Australian National University,Canberra, ACT 2601, Australia¶Author for correspondence ([email protected])

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.

SUMMARY

Usher syndrome is the most prevalent cause of hereditary deaf-blindness, characterized by congenital sensorineural hearing impairment andprogressive photoreceptor degeneration beginning in childhood or adolescence. Diagnosis and management of this disease are complex, and themolecular changes underlying sensory cell impairment remain poorly understood. Here we characterize two zebrafish models for a severe form ofUsher syndrome, Usher syndrome type 1C (USH1C): one model is a mutant with a newly identified ush1c nonsense mutation, and the other is amorpholino knockdown of ush1c. Both have defects in hearing, balance and visual function from the first week of life. Histological analyses revealspecific defects in sensory cell structure that are consistent with these behavioral phenotypes and could implicate Müller glia in the retinal pathologyof 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.

Harmonin (Ush1c) is required in zebrafish Müller glialcells for photoreceptor synaptic development andfunctionJennifer 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,¶

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Reiners et al., 2006; Overlack et al., 2008; Williams et al., 2009),suggesting that at least some have functions independent of theproposed complex. In hair cells, USH proteins are thought tostabilize the growth and orientation of stereocilia during development(for reviews, see Brown et al., 2008; Yan and Liu, 2010), whereas USHproteins in the region of the connecting cilium are proposed tocontribute to the trafficking of molecular cargo from thephotoreceptor inner segment to the outer segment (Maerker et al.,2008; Liu et al., 2007). USH proteins have also been immunolocalizedat the ribbon synapses of these sensory cells, but their potentialfunctions in this region are unknown.

The USH1C gene encodes the PDZ-domain-containing proteinharmonin (Verpy et al., 2000), which has been proposed to functionas a key scaffolding molecule to which most other USH proteinscan bind (Reiners et al., 2005). Patients with mutations in USH1Cexhibit the classic USH1 pathology, with profound congenitalhearing impairment, vestibular dysfunction, and clinicallyappreciable retinal degeneration in childhood or early adolescence.USH1C mutations resulting in non-syndromic deafness have alsobeen reported (Ouyang et al., 2002). The deaf circler mouse(Johnson et al., 2003), harboring an in-frame deletion in themurine Ush1c gene, has provided a valuable model for themechanosensory hair cell defects that cause deafness in USH1Cpatients, but does not exhibit a retinal phenotype (Williams et al.,2009). Similarly, the Ush1c knockout mouse exhibits profounddeafness 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>Amutation (plus surrounding sequence) cloned from an AcadianUSH1C patient (Lentz et al., 2007), shows both the well-characterized auditory defects of the other USH1C mouse models,as well as visual defects comparable to the human disease. Inaddition to the disorganized stereocilia and hair cell degenerationobserved in the cochleae of USH1C c.216G>A mice, reduced visualfunction was noted after 1 month of age and progressive loss ofrod photoreceptors was observable after 6.5 months of age (Lentzet al., 2010). These intriguing differences in retinal pathologybetween the Ush1c knockout and the USH1C c.216G>A knock-incould indicate differences between the functions of endogenousmouse and human harmonin in the retina.

In vitro assays have demonstrated that many of the known USHproteins bind preferentially to one or more of the three PDZ domainsin harmonin, hence its depiction as a central scaffold upon whichan USH protein complex can form (Adato et al., 2005). Although thesubcellular localization of harmonin and other USH proteins in haircells is compatible with this hypothesis, there have been conflictingreports of localization of other USH proteins at the photoreceptorsynapse, where harmonin localization is undisputed (Reiners et al.,2005; Liu et al., 2007; Yang et al., 2010). Moreover, harmonin isnotably absent from the connecting cilium region (Reiners et al., 2005;Williams et al., 2009), where the presence of most other USH proteinshas 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, andthe early, pre-degenerative manifestations of the retinal pathologyin USH1C remain mysterious.

Previously, zebrafish mutants for myo7a (ush1b), cdh23 (ush1d)and pcdh15 (ush1f) have been described as exhibiting severe

swimming and balance defects consistent with the auditory andvestibular component of the human disease (Ernest et al., 2000;Söllner et al., 2004; Seiler et al., 2005). Although expression in theretina has been reported for each of these genes, only themorpholino knockdown of pcdh15b has been reported to have aneffect on visual function (Seiler et al., 2005). Photoreceptororganization is severely affected in pcdh15b morphants, andelectroretinogram (ERG) traces are notably attenuated. Here, wedescribe two zebrafish models for USH1C that exhibit a strongUSH-like phenotype at early developmental stages. Young fish withdepleted ush1c function have penetrant and specific visual defectsas well as compromised hair cell development, resulting in hearingand balance impairment. Detailed analyses of these phenotypeshave confirmed a requirement for harmonin in zebrafish hair cellmorphogenesis, and have implicated Müller glial cells in supportingharmonin-mediated ribbon synapse stability and function. USH hastraditionally not been considered a developmental disease of vision.The results of this study, however, show that visual defects can bedetected from the onset of vision in zebrafish with depleted ush1cfunction, thus providing a rationale for examination of Müller cellfunction 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 tBLASTnsearches to identify a single ortholog of USH1C in the zebrafishgenome (Fig. 1A). We then used a radiation hybrid panel (Hukriedeet al., 1999) to position this gene on chromosome 25. Previousstudies of the mouse and human genes have indicated that themammalian genes express three splice forms, which vary slightlyin structure and subcellular localization (Reiners et al., 2003;Reiners et al., 2005; Williams et al., 2009). The most abundantzebrafish transcript found in whole larvae or adult retinascorresponded to isoform A, containing 21 exons that encode a 548amino acid protein with three PDZ domains and a coiled-coildomain (Fig. 1A). Zebrafish harmonin isoform A was found to bestructurally similar to the corresponding human isoform, with 70%amino acid identity overall and highest similarity in the PDZdomains and the N-terminal region (Pan et al., 2009). Ahydrophobicity plot of the human and zebrafish proteins revealeda high level of conservation, even where residue substitutions occur(Fig. 1A). Using reverse transcriptase (RT)-PCR and RACE primerswithin and between regions encoding the second and third PDZdomains, we identified unique transcripts that might correspondto the B or C isoforms of the gene, although no variant encodinga second coiled-coil or PST domain, as would be predicted for anisoform B transcript, was identified. All transcripts other than thatof isoform A were in low abundance such that we were unable toobtain sequence definitively identifying them as belonging to aparticular isoform class. The probes generated from these partialsequences showed weak, generalized signal in all tissues and timepoints analyzed (data not shown).

cRNA probes directed against either the unique sequence ofisoform A or the 5� region predicted to be common to all spliceforms yielded identical expression patterns in larval tissues. RNAexpression 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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hair cell expression persisted through all later larval stages examined[5 days post-fertilization (dpf) shown in Fig. 1B,C]. We alsoobserved strong expression in the neuromasts of the lateral line(Fig. 1C). This neuromast expression was consistent with previousreports of the zebrafish USH gene orthologs myo7a, cdh23 andpcdh15a (Ernest et al., 2000; Söllner et al., 2004; Seiler et al., 2005).Retinal expression was present beginning at 48 hpf, when retinallamination is complete and cell specification is underway. By 72hpf, strong expression was observed in a subset of cells in the innernuclear layer, which persisted through all larval stages examined(5 dpf, shown in Fig. 1D). In situ hybridization of adult retinal tissueusing all described probes showed persistent, strong expression inthe inner nuclear layer as well as expression in the outer nuclearlayer (Fig. 1E). We also observed strong expression of ush1c in thegut and swim bladder, and weaker expression in the brain and trunkmuscles (Fig. 1B), consistent with reported expression in humantissues (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 ZebrafishTILLING Consortium by screening for mutations in the first PDZ-domain-encoding region of the gene (PDZ1). We recovered a line,ush1cfh293, that harbors a nonsense mutation in the fifth exon, whichencodes the C-terminal portion of the PDZ1 domain (redarrowhead, Fig. 1A; supplementary material Fig. S1).

We also designed morpholino oligonucleotides to knock downthe function of ush1c: one to block splicing at the exon-2–intron-

2 boundary (MO1; green arrowhead, Fig. 1A) and one thatrecognizes the translation initiation site (MO2). We obtainedidentical phenotypes with both morpholinos, but found that theexon-2-targeting morpholino (MO1) provided more consistentresults; thus, MO1 was used for all morpholino experiments andall uses of ‘morpholino’, ‘MO’ or ‘morphant’ refer to MO1. UsingRT-PCR analysis of MO-injected embryos and larvae, we identifiedan aberrant splice form in which exon 2 is skipped, producing out-of-frame splicing of exons 1 and 3 that results in an earlytermination codon (supplementary material Fig. S1). This alteredsplice form predominates for the first 4 dpf, and persists through6 dpf, although recovery of the normal splice form is evident by 5dpf (Fig. 2A).

We obtained a polyclonal rabbit antibody raised against the first100 amino acids of zebrafish harmonin (red bar, Fig. 1A), a regioncommon to all predicted isoforms, and probed protein extracts ona western blot to test antibody specificity and to determine theextent of protein depletion in morphant and mutant larvae (Fig.2B). In wild-type controls, we observed a single band in the 3 dpfsample, which was slightly smaller than the corresponding bandsin the 4 dpf and 5 dpf samples. In addition to this band, which, atapproximately 60 kDa, corresponded to the predicted size ofzebrafish harmonin isoform A, we noted progressively increasingnumbers of smaller bands in the 4 dpf and 5 dpf controls. Consistentwith the time course of morpholino efficacy revealed by RT-PCR(Fig. 2A), strong knockdown of harmonin protein levels wasobserved at 3-4 dpf in ush1c morphants. Moreover, although theupper band was restored nearly to wild-type levels in the 5 dpf


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 exonsthat produce a 1.6 kb mRNA. A hydrophobicity plot of zebrafish (red) and human (blue) isoform A protein sequence shows conserved residue polarity. % aminoacid identity with human protein is indicated below the plot. The positions of the ush1cfh293 nonsense mutation (red arrowhead), the MO target (greenarrowhead) 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, lateraland posterior cristae, respectively; am: anterior macula) and an adjacent head neuromast (n). (D) ush1c transcripts are restricted to cells in the inner nuclear layerin 5 dpf sectioned retina. (E) Central section of an adult zebrafish retina showing continued strong expression in the inner retina; expression is detected in theouter nuclear layer as well. onl, outer nuclear layer; inl, inner nuclear layer; gcl, ganglion cell layer. Scale bars: 100 m (B); 20 m (C); 50 m (D,E).

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sample, the size was slightly reduced, consistent with the controlband from the 3 dpf time point (Fig. 2B). These temporal changesin size and abundance of the major band, recapitulated by therecovering morphant protein extracts, might reflect a period ofpost-translational modification.

We performed RT-PCR experiments on zebrafish cDNA frommultiple developmental stages and discovered five distinctexpressed sequence tags (ESTs), but no complete sequences of splicevariants other than isoform A. Thus, we cannot definitively identifythe smaller bands on the western blots, but postulate that theymight be additional isoforms or products of protein degradationof isoform A, the largest running band. Similar, smaller bands weredetected by western analysis of mouse harmonin (Williams et al.,2009). We did not observe any bands larger than ~60 kDa at anylarval time point examined. This, combined with the RT-PCR andRACE results described above, suggests that isoform B splicevariants are not active through 5 dpf.

No bands were detected in protein samples from ush1c mutantlarvae at any time point examined (5 dpf shown alongside wild-type sibling in Fig. 2B). However, the ush1c nonsense mutationin the fifth exon could potentially produce a truncated proteinthat would be recognized by the antibody. We investigated RNAlevels in ush1c mutants using a riboprobe against the first six

exons of ush1c. We observed reduced probe signal in all ush1c-expressing tissues examined (ear and gut tissue shown insupplementary material Fig. S2), suggesting that this mutanttranscript is unstable.

Reduced harmonin function produces defects in vestibular,

auditory and visual function

At 5 dpf, wild-type fish exhibited rapid evasive swimming behaviorwhen startled by a tap on the Petri dish. We tested the startle reflexof 5 dpf offspring from natural crosses between ush1c–/+

heterozygous carriers (n 952 larvae) and found that 25% (n 236)of these larvae had abnormal responses (Fig. 2C, top). Within thisgroup, 83% did not exhibit any evasive swimming behavior, andthose that did respond swam in circles. Subsequent genotypingrevealed that 100% of the larvae exhibiting these defects werehomozygous ush1cfh293 mutants.

Free-swimming 5 dpf ush1c morphant larvae exhibited vestibulardefects identical to ush1c mutants, with only 30% of the larvaeresponding to tapping on the Petri dish, and all responders swimmingin circles (n 400). These defects are similar to those reported forother zebrafish USH gene mutations or knockdowns (Ernest et al.,2000; Söllner et al., 2004; Seiler et al., 2005) and indicate reducedfunction of both the auditory and vestibular systems.

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Fig. 2. Mutation and morpholino knockdown of ush1c produce behavioral defects. (A) RT-PCR of control and splice-blocking morpholino-injected embryosfrom 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) Westernblots 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 kDaband 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 inthe 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 aslightly smaller weight, and there is a reduced number of smaller bands in this sample. In extracts from 5 dpf ush1cfh293 homozygous mutants, no bands areobserved. Extracts from wild-type siblings from the same clutch show the characteristic banding pattern of the 5 dpf uninjected controls, allowing for the slightlylonger running time for this gel. A pan-actin antibody was used as a loading control. (C) Behavioral assays of control, mutant and morpholino-injected larvaeshow (top) impaired startle response and balance, and (bottom) reduced optokinetic response, tracking an average of 50% or fewer of the dark stripes (averagetargets 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; n 30 for each group).

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To assess visual function in mutant and morphant animals, weused the optokinetic response (OKR) assay. At 5 dpf, ush1c mutantsand 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 defectsin harmonin-deficient larvae, we used immunofluorescence toexamine 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 ush1cmutants (Fig. 3B) or morphants (Fig. 3C), confirming the strongknockdown or depletion indicated by the western blot results. Usingphalloidin to visualize stereocilia, we examined individual sensorypatches. At 5 dpf, wild-type hair bundles had a characteristic shapeand orientation, projecting more or less orthogonally from the planeof each cuticular plate and appearing tightly tapered at their tips(Fig. 3D-F). All otic sensory patches in the mutant larvae displayedreduced numbers of hair bundles, as well as defects in stereociliaorganization at 5 dpf (Fig. 3G-I). We observed 40% fewer hairbundles than in wild-type siblings in the anterior and posteriormaculae, and existing stereocilia were splayed and bent, with anotable lack of tapering of the hair bundles. In morpholino-

injected animals, we found a reduced number of hair bundles, with48% fewer bundles in the anterior maculae and 33% fewer hairbundles in the posterior maculae of 5 dpf morphants comparedwith controls. Stereocilia were also disorganized, although to alesser extent than we observed in the mutant. Bundles appearedbent and disoriented relative to the plane of the cuticular plate, butthe splaying or absence of tapering at the tips was less pronouncedthan in ush1c mutants (Fig. 3J-L).

We next investigated the subcellular localization of harmoninin the anterior maculae of 5 dpf larvae (Fig. 4). In wild-type haircells, harmonin localized to both stereocilia and kinocilia as wellas within cell bodies and at synapses (Fig. 4A-G). In ush1cmutants (Fig. 4H-J), we detected reduced levels of harmonin instereocilia and kinocilia. Protein was also detected within cellbodies, but the distribution appeared abnormal compared withcontrols. Harmonin levels in hair cells of morphant larvae (Fig.4K-M) were markedly diminished at 5 dpf, although some signalcould be detected apical to the hair bundle in a position consistentwith kinocilia localization. When we injected embryos fromush1c+/– heterozygous parents with the ush1c morpholino, raisedthe larvae to 5 dpf and analyzed harmonin staining in sensorypatches, residual staining was undetectable in injectedgenotypically ush1c–/– animals, suggesting that a truncated formof harmonin is produced in the mutants. This truncated proteinmight be too low in abundance or too unstable to be detected in



Fig. 3. Harmonin is required in otic sensory patches for normal bundle development. (A-C) Phalloidin-stained F-actin (green) and anti-zebrafish harmoninantibody 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 sensorypatches and in neuromasts in ush1c mutants (B) and ush1c-MO-injected larvae (C). (D-L) High-magnification view of phalloidin-stained hair bundles in theanterior and posterior maculae of 5 dpf larvae. Hair bundle number is reduced and organization of the existing bundles is disrupted in mutant (G-I) andmorphant (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). Close-up 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 wild-type average 54.6±3.5 bundles; n 8; ush1c mutant average 33±3.5; n 7; P<0.0001. Posterior macula wild-type average 68.2±2.6 bundles; n 9; ush1c mutantaverage 40.5±4.2; n 10; P<0.0001. For the morpholino experiment, anterior macula control average 41.8±4.8 bundles; MO average 21.8±3.8; n 5 for eachgroup; P<0.0001. Posterior macula control average 71.8±4.6 bundles; MO average 48.2±6.9; n 5 for each; P<0.0001, 5 dpf. Scale bars: 50 m (A-C); 10 m (D-L).

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a western blot, but was detectable in fixed tissue using enhancedsignal detection protocols.

To investigate cell degeneration or reduced cell proliferation aspossible causes for the observed reduction in hair bundle number,we stained larvae at stages between 1 and 5 dpf with TUNEL tolabel dying cells or BrdU to label dividing cells. The results in bothcases were indistinguishable between morphants and controls (notshown), and were consistent with previously reported normal celldeath levels in the ear (Bever and Fekete, 1999). Mutant cellproliferation was also unaffected at these stages, as were cell deathlevels through 4 dpf. In 5 dpf mutants, we noted a small butsignificant increase in TUNEL-labeled cells for both the anteriormacula (0.1±0.32 for wild type vs 1.2±1.03 for mutant; n 10 foreach group; P 0.008) and posterior macula (0.1±0.32 for wild typevs 1.3±0.95 for mutant; n 10 for each; P 0.003). Althoughstatistically significant, the numbers of dead cells were insufficientto account for the extent of the reduction in bundles. To confirmthat hair cells were being specified properly in mutant larvae, weused a probe to the hair-cell-specification factor atoh1b and notedno differences in labeling of sensory patches in wild type comparedwith mutant (data not shown). Thus, we conclude that the defectsin hair bundle formation occur post-mitotically, consistent withpreviously reported studies in mammals (Lefevre et al., 2008), andthat increased cell death, reduced cell proliferation, or impaired

cell fate specification cannot be the primary causes of bundle loss.Together, these results confirm a requirement for harmonin in hairbundle development. Additionally, the milder stereociliadisorganization phenotype in ush1c morphants observed at 5 dpf,in conjunction with the partial recovery of harmonin localizationat 5 dpf observed in the region of the kinocilia, suggests thatharmonin 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 andmorphants compared with wild type (Fig. 5A,F,K; supplementarymaterial Fig. S3); however, because visual function defects wereobserved, we also examined harmonin protein localization inretinas. We observed abundant protein levels specifically in the cellbodies and processes of Müller glia at all larval stages examined(Fig. 5B-E; supplementary material Fig. S4), consistent with therestricted inner retinal expression noted by in situ hybridizationwith our ush1c riboprobes at 5 dpf (Fig. 1D). We did not observespecific photoreceptor labeling with this antibody between 5 and8 dpf.

We examined harmonin antibody labeling in wild-type, mutantand morphant retinas at 5 dpf. Harmonin protein levels in theretinas of ush1c–/– homozygous mutants were markedly reduced,although Müller cells were present and appeared morphologically

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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 hairbundles (green), kinocilia (arrowheads), cell bodies and at synapticboutons (arrow in C and D). Incubation conditions favoring hairbundle visualization (E-P) show strong colocalization of harmoninwithin the bundle structure as well as in the cuticular plate in 5 dpfwild types (E-G). Kinocilia labeling can be seen above the bundleregion (asterisks). Enhanced signal detection in mutant hair cells (H-J) shows localization persisting in the kinocilia and in existingbundle structures. Kinocilia are more visible in these panels owing tothe reduced bundle architecture in the mutant. Protein distributionseems globular and disorganized in the cell bodies (arrowheads).Hair cells of ush1c MO larvae (K-M) show strong knockdown ofharmonin, with localization persisting only in kinocilia at this stage,and no signal observed within hair bundles. Injecting themorpholino into the ush1c–/– mutants reduces subcellularlocalization of harmonin to undetectable levels in hair cells (N-P).Scale bars: 10 m.

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normal (Fig. 5G-J). Similarly, after morpholino injection, weobserved a strong reduction of harmonin labeling in Müller cellsthrough 5 dpf (Fig. 5L-O). We noted that the duration of proteindepletion was prolonged in the eye, compared with our results frommorphant hair cells, in which we observed rebounding harmoninlocalization in some kinocilia by 5 dpf (Fig. 3F). We thereforeexamined harmonin labeling in 6 dpf morphants, at which timewe observed increased harmonin labeling in Müller cells of theperipheral retina, adjacent to the ciliary marginal zone(supplementary material Fig. S4).

Owing to the apparent presence of ush1c transcript in adultphotoreceptors (Fig. 1E), we evaluated the subcellular localizationof harmonin in 6 month adult retinas (Fig. 6A-F). We continuedto see strong harmonin localization within Müller cells in the adultretina, extending past the outer limiting membrane (OLM) and intothe glial processes in the subapical region between photoreceptorcell bodies (Fig. 6, asterisks). In some adult retinas we also observedwhat might be a low level of protein localization at thephotoreceptor synapses (Fig. 6A, arrow). Occasionally, signal wasdetected in photoreceptor inner or outer segments at a lower leveland with inconsistent distribution compared with the robuststaining observed in Müller glia (Fig. 6D, arrow), suggesting thatthis signal was due to nonspecific labeling or autofluorescence,although we were unable to confirm this with our currentimmunofluorescence methods.

Because Müller cell localization of harmonin has not beenreported previously, we used antibodies against murine (Fig. 6G-I) and human (Fig. 6J-L) harmonin to label retinas from postnatalday 10-11 mouse pups. In addition to the previously describedlabeling in photoreceptors, we also observed harmoninimmunoreactivity in Müller glial cell bodies and processesextending throughout the retina. The anti-human harmoninantibody (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 Müllercells differs from published work (Reiners et al., 2005), furtherinvestigation is needed, but our results are consistent with the

possibility of a phylogenetically conserved role for harmonin inretinal glia.

Harmonin was recently detected in the postsynaptic processesof murine bipolar cells and horizontal cells, using immunoelectronmicroscopy (Williams et al., 2009). With our immunofluorescenceprotocols, we found no evidence for harmonin localization withincell bodies of bipolar or horizontal cells at any time points examinedin zebrafish (Fig. 5; Fig. 6A,C). Owing to the strong harmoninlabeling in Müller glial processes, we cannot determine whethersynaptic processes of horizontal or bipolar cells might also bepositive 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 andmorphants at 5 and 8 dpf, using an anti-active caspase-3 antibody.Death rates by retinal cell layer were as follows (n 10 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 comparedwith 0.31±0.60 in morphant retinas (P 0.5) and 3.09±1.3 in mutants(P<0.0001); in the inner nuclear layer, no labeled cells were observedin 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±0.45in morphants (P 0.16) and 0.09±0.30 in mutants (P 0.79). Notingthe small but significant elevation of inner nuclear layer (INL) celldeath in 5 dpf mutants and morphants, we co-labeled sectioned 5dpf retinas with caspase-3 and glutamine synthetase to determinewhat proportion of the dying cells were Müller glia (n 10 for eachgroup). In ush1c–/– animals, exhibiting an average of 1.88 labeledcells per eye, 67% of the cells labeled with caspase-3 were alsopositive for glutamine synthetase. Similarly, 61% of caspase-labeledcells were co-labeled with glutamine synthetase in ush1c morphantretinas, in which an average of 1.85 cells in the INL were labeled.In both cases, the remaining labeled cells could not be positivelyidentified as Müller cells, based on either their position within thenuclear layer or by labeling with the Müller cell marker. Althoughour current experiments do not rule out the possibility that a smallsubset (<0.7 cells per eye) of caspase-positive cells in the INL might



Fig. 5. Harmonin localization in the larval retina is

restricted to Müller glia. Semi-thin plastic sections stainedwith toluidine blue show consistent size, lamination andorganization of 5 dpf wild-type, mutant and morphantretinas (A,F,K). 16- m larval retina sections (B,G,L) showcolocalization of the Müller cell marker glutaminesynthetase (GS; green) and anti-zebrafish harmonin (red) inwild type (B). Both antibodies label Müller cell processesthroughout the retina. Anti-zebrafish harmonin labeling isreduced in mutant (G) and morphant (L) Müller cells. Müllercell number and morphology is unaffected. Nonspecificsignal associated with photoreceptor outer segments isobserved in all panels at this magnification. Seesupplementary material Fig. S4 for comparison. High-magnification images of anti-zebrafish harmonin and GSlabeling in Müller cell processes at the OPL (large arrow) andOLM (small arrow) of the 5 dpf retina are shown (wild type,C-E; mutant, H-J; morphant, M-O). Scale bars: 50 m(A,B,F,G,K,L); 20 m (C-E,H-J,M-O).

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be second order neurons, it is also possible that these cells wereapoptotic Müller glia that no longer produced glutamine synthetase.

At 8 dpf, we noted increased photoreceptor cell death in theush1c mutant retinas, compared with wild-type animals (wild type:0 labeled photoreceptors; mutant: 5.33±1.32 labeledphotoreceptors; P<0.0001; see supplementary material Fig. S4). Nostatistically significant differences were observed between mutantand wild-type siblings in either the inner nuclear or ganglion celllayers at this stage, nor were there significant differences seen inthe labeling of morpholino and control animals in any cell layersat 8 dpf.

Harmonin is required for photoreceptor synaptic organization and

function

The specific depletion of harmonin from Müller cells in larvalmutant and morphant retinas provided an opportunity to studythe role of harmonin in this cell type. Because the morphantspresented no significant photoreceptor cell death at 5 dpf despitestrong protein knockdown, we used morphants for the followinganalyses. We began by examining synaptic integrity in morphantlarvae at 5 dpf, using antibodies against the pre- and postsynapticproteins SV2 and GluR2/3, respectively. In control retinas, thesemarkers exhibited an organized, parallel and largely colocalizedalignment within the OPL (Fig. 7A). In ush1c morphants, bothproteins were present in the OPL, but they were remarkablymisaligned and less colocalized, indicating a disruption in synapticorganization (Fig. 7B).

To investigate this phenotype further, we examined theultrastructure of cone pedicles in morphant larvae via transmissionelectron microscopy. We examined cone pedicles from retinas oftwo uninjected controls at 6 dpf in a quantitative comparison withcone pedicle ultrastructure from retinas of four stage-matchedush1c MO larvae. We found that cone pedicles in uninjectedcontrols (Fig. 7C,E) contained an average of 1.667±0.299 ribbonsper pedicle (n 36 pedicles) and a large number of mature-appearingribbon synapses, with 55% of the synaptic ribbon profiles anchoredby an arciform density (ribbons with arciform density perpedicle 1.08±0.238); 85% of these membrane-anchored ribbonswere associated with postsynaptic processes from horizontal andbipolar cells in a triad (triads per pedicle 0.97±0.128). In morphantretinas (Fig. 7D,F), the average number of ribbons per pedicle wasconsistent with controls (1.659±0.607; n 44 pedicles), but only 38%of the ribbon profiles appeared anchored by arciform densities(ribbons with arciform densities per pedicle 0.636±0.647;P 0.0002). Synaptic ribbons unanchored by an arciform densityseemed to ‘float’ in the pedicle, and postsynaptic processes, whenpresent, frequently appeared distant from these floating ribbons(Fig. 7D,F). As in the controls, most (83%) anchored ribbons inmorphants were incorporated into triads (triads perpedicle 0.523±0.412), suggesting a specific defect in ribbonassociation with the membrane. To investigate whether theobserved defects could be attributed to a developmental delay, wealso examined sections of ribbon synapses in cone pedicles of twowild-type and three morphant retinas at 8 dpf. The number ofribbons observed per pedicle (1.34±0.56; n 41) was significantlyreduced in 8 dpf morphants compared with controls (1.70±0.22;n 43; P 0.0002). As seen in the 5 dpf analysis, significantly fewerribbons in the 8 dpf morphants (52%; n 55 ribbons) compared withcontrols (82%; n 73 ribbons; P<0.0001) appeared to be anchoredto the membrane. The percentage of anchored ribbons associatedwith triads was comparable between morphants and controls (68%and 65%, respectively), again suggesting a primary defect in theformation of arciform densities. Retinal morphology was otherwiseunaffected, and significant retinal cell death was not observed in8 dpf morphants in any retinal cell layers by anti-active caspase-3labeling. Thus, we conclude that temporary depletion of harmoninfrom Müller glia during retinal development has a lasting, adverseimpact on maturation of cone ribbon synapses.

To determine the effect of the morphological defect on synaptictransmission, we recorded ERGs from 5 dpf morphant larvae (n 6)

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Fig. 6. Harmonin localizes in Müller cells through all zebrafish life stages

and is evolutionarily conserved. (A-F) Localization of anti-zebrafishharmonin (red) in Müller cells persists in adult zebrafish retinas. Some low levelsignal is sometimes observed in photoreceptor synapses (arrow in A) and inphotoreceptor outer segments (arrow in D). The strongest signal within thephotoreceptor layer is associated with Müller cell processes projecting into thesubapical region (asterisks). Nonspecific fluorescence of photoreceptor outersegments at this stage can be compared with panel H in supplementarymaterial Fig. S4. (G-L) Anti-mouse harmonin (SDI; G,I) and anti-humanharmonin (Santa Cruz; J,L) detect localization in photoreceptors and Müllercells [glutamine synthetase (GS) is labeled in green; harmonin antibodies inred] in retinas from P10-P11 mouse pups. The outer nuclear layer (onl), innernuclear layer (inl) and ganglion cell layer (gcl) are labeled in merged panels.Scale bars: 50 m (A-C,G-L); 20 m (D-F).

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and compared them with uninjected larvae (n 7). Rodphotoreceptors are detectable molecularly by 3 dpf andmorphologically by 5 dpf. However, physiological analysesdemonstrate that rods are not functional until 15 dpf (Morris andFadool, 2005). ERGs performed at 5 dpf show a cone dominantresponse in both light-adapted and dark-adapted conditions (Bilottaet al., 2001; Seeliger at el., 2002), but dark-adapted retinas at thisstage are less susceptible to bleaching and exhibit a more sensitiveON response than in light-adapted conditions (Makhankov et al.,2004). Thus, we chose to perform our experiments under scotopicconditions, in which we observed that the b-wave amplitude, anindicator of synaptic transmission, was markedly reduced in theMO-injected animals (Fig. 8A) compared with controls. The a-wave, which is primarily a read out of photoreceptor membranedepolarization as a result of phototransduction, is often quenchedby conditions conducive to generating b-wave amplitudes and istherefore not observable on the same trace. Thus, we isolated thea-wave pharmacologically with the synaptic transmission inhibitorsL-AP4 and TBOA (Trombley and Westbrook, 1992; Huang andBordey, 2004; Wong et al., 2005), whereupon we observed a slightreduction in isolated a-wave amplitude compared with controls(P 0.01), but less of an effect overall than that observed from theb-wave experiments (P 0.0007; Fig. 8B). Plotting a-wave and b-waveamplitudes against each other shows that the b-wave to a-wave ratiois significantly diminished in the morphant group (P<0.0007; Fig.8C). These data, taken together with the disorganization of synapticproteins in the OPL (Fig. 7B), indicate that postsynaptic activity isreduced relative to the level of the phototransduction response tolight stimulus (Takada et al., 2008). Although we observed a verysmall increase in the number of caspase-positive cells in the INL,the majority of which were identified as Müller glia, we would notexpect the loss of so few cells to account for such a large reduction

in the b-wave amplitude. Thus, our data suggest that loss ofharmonin in Müller cells results in decreased function of thephotoreceptor ribbon synapses.

DISCUSSION

In this study, we describe the requirement for harmonin in zebrafishmechanosensory hair cell and retinal development, providing thefirst evidence that USH proteins function in photoreceptor synaptictransmission and ribbon synapse development. Our studies alsoimplicate Müller glial cells as contributors to the retinal pathologyof USH.

The localization of harmonin in hair cells and lateral lineneuromasts indicates a phylogenetically conserved role inmechanosensory cells, and our studies of the zebrafish inner earreveal defects in hair bundle organization and development,consistent with the stereocilia elongation and organization defectsdescribed in Ush1c mutant mice (Lefevre et al., 2008). Our resultsindicate that, in zebrafish, the predominant splice variant andprotein isoform present during larval development is harmoninisoform A. The severe disruption of hair bundle development seenin ush1c mutant and morphant zebrafish provides an intriguingcontrast to the absence of hair bundle defects reported in somecolonies of dfcr mice, which harbor a deletion affecting harmoninisoform B (Grillet et al., 2009).

The presence of the USH protein harmonin in Müller glia froman early time point in retinal development and the specificdefects in synaptic structure and function in ush1c morphantsprior to detectable ush1c transcript or protein synthesis inphotoreceptors suggest a non-cell-autonomous role for harmoninin facilitating synaptic maturation in cone pedicles. Müller glialcells have established roles in the maintenance of retinal neuronsvia their regulation of glutamate neurotransmitter levels and their



Fig. 7. Synaptic maturation is impaired in ush1cmorphants. (A,B) Anti-SV2 (red) and -GluR2/3 (green)antibodies mark the pre-and postsynaptic regions of theOPL, respectively. Tightly associated localization of thesefactors is apparent in the control retina (A), and is notablydisrupted in the morphant retina (B) at 5 dpf. (C-F) Electronmicrographs of cone pedicles in 6 dpf control and morphantlarvae. In control retinas (C,E), the presence of multiple triads(boxed areas), consisting of a synaptic ribbon (r), arciformdensity (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 inpanel F), and floating ribbons (r) and distant postsynapticprocesses (h) are observed in cone pedicles. Scale bars:20 m (A,B); 1 m (C,D); 500 nm (E,F).

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expression of neurotrophic factors (Fields and Stevens-Graham,2002; Newman, 2004; de Melo Reis et al., 2008). Although a rolefor Müller glia in retinal synaptogenesis has been suggested byseveral studies (Georges et al., 2006; Diaz et al., 2007), little isknown about how this putative function might be accomplished.In the brain, glutamate receptor function has been shown to beimportant for synaptogenesis (Han and Stevens, 2009; Kakegawaet al., 2009), so glutamate uptake by Müller cells might similarlybe important for synaptic development of photoreceptors.Interestingly, a recent zebrafish study (Williams et al., 2010) useda Müller-cell-specific line to ablate individual glia and observeeffects on synaptogenesis and the behavior of neighboring glia.Their data indicated that localized loss of Müller cells did notaffect the maturation of cone synapses at 5 dpf. These data areconsistent with our findings, because they argue against the smalland transient levels of Müller cell death being the cause of thereduced synaptic function and structural defects observed, andsuggest instead that harmonin in Müller cells has a paracrine effecton cone synapse stability, such that the sustained absence of thisprotein results in the observed dysfunction and progressivephotoreceptor cell death.

The cone synapse phenotype of ush1c morphants is similar tothat of the zebrafish nrc mutant, in which the synaptojanin geneis disrupted (Allwardt et al., 2001; Van Epps et al., 2004). The nrcmutant has a diminished b-wave and defects in photoreceptorsynapse formation, with floating ribbons and unassociatedpostsynaptic processes in cone pedicles. Synaptojanin has beenshown to be required for cytoskeletal organization, endocytosis,Ca2+ signaling and post-synaptic glutamate transport (Sakisakaet al., 1997; Schuske et al., 2003; Gong and DeCamilli, 2008).Although no link between synaptojanin and USH has beenproposed previously, nrc mutants have swimming and balancedefects in addition to blindness (Van Epps et al., 2004), similarto the USH-like phenotype we observe in ush1c mutants andmorphants. More recently, a report describing three new mutantalleles of zebrafish synaptojanin, isolated by screening forvestibular defects, illustrates synaptic transmission defects in thehair cells (Trapani et al., 2009).

Given the distinct, penetrant defects in visual function andphotoreceptor synapse formation in harmonin-depleted larvae, we

were surprised by the lack of harmonin localization in larvalphotoreceptors. It is puzzling that there would be no requirementfor harmonin function in larval zebrafish photoreceptors, especiallyin light of the abundant antibody signal detected in thephotoreceptors of young mice. An as-yet-undetected duplicateush1c gene in zebrafish could account for the apparent proteinlocalization discrepancies between mammals and zebrafish.However, our searches of the zebrafish genome reveal only onelocus for ush1c, and we find only one copy of this gene in theTetraodon, Takifugu and Threespine stickleback genomeassemblies, suggesting that only a single copy of the ush1c genehas been retained in the teleost lineage.

Another possible explanation for the absence of harmonin inlarval photoreceptors is the presence of ohnologous genes for twoother PDZ-domain-containing proteins: Cip98 and Pdzd7.Alignments with the human CIP98 (whirlin) protein sequence,causative of USH2D, uncovered two orthologs in the zebrafishgenome assembly. Both of these transcripts, cip98a and cip98b, areexpressed in photoreceptors from early larval time points (J.B.P.,unpublished); thus, the presence of an additional PDZ protein withsimilar binding affinities to harmonin (van Wijk et al., 2006) mightbe able to compensate for the absence of ush1c expression in thesecells. However, given that the N-terminal regions of harmonin andCip98 are not functionally interchangeable (Pan et al., 2009), anyfunctional substitution of harmonin by Cip98 in the photoreceptormust not rely on protein interactions in the N-terminal region.Although PDZD7 is not currently recognized as a monogenic causeof human USH, it has been shown to act as a modifier of USH genefunction and to interact physically with USH proteins. Bothzebrafish genes, pdzd7a and pdzd7b, are expressed inphotoreceptors at larval stages (Ebermann et al., 2010).

Our ERG recordings from ush1c morphant larvae reveal aspecific reduction in b-wave amplitude in the absence of significantretinal degeneration, which might provide an explanation for theearly cellular dysfunctions that precede retinitis pigmentosa inUSH1C patients. A retrospective ERG analysis of USH1 patientsas young as 17 months revealed abnormal waveforms prior toclinically detectable retinitis pigmentosa (Flores-Guevara et al.,2009), although other studies of USH1 patients in whichphotoreceptor loss was already clinically evident revealed no

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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 significantlyreduced 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 areonly slightly attenuated in morphant animals compared with controls. Light stimulus duration is 800 mseconds. (C) Comparison of the relative changes of b- anda-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 b-wave:a-wave ratio indicates a specific synaptic defect. Steeper slopes of the trend lines (dashed lines) represent reduced synaptic efficacy.

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synaptic dysfunction in the remaining viable photoreceptors(Jacobson et al., 2008; Williams et al., 2009). Notably, theUsh1c216AA knock-in mouse (Lentz et al., 2010) exhibits anattenuated ERG as early as 1 month of age, prior to any histologicallyappreciable cell loss. However, in contrast to our results with larvalzebrafish, both a- and b-waves were attenuated in Ush1c216AAmice. It could be that, in human manifestations of USH1C, the laterconsequence of diminished harmonin levels in photoreceptors isso severe that it obscures comparatively subtle early defects thatmight result from impaired synaptic activity.

Our data also indicate that sustained absence of harmonin inMüller cells, as seen in ush1c mutants, results in progressive retinaldegeneration. It is not clear whether this cell death results fromsynaptic defects or from some other impairment of Müller cellfunction, but the non-cell-autonomous nature of this degenerationis notable. Clinical analyses of the retinal pathology of USH patientsmight provide insight into the link between Müller cell functionand retinal integrity. Ophthalmic examinations of USH1B patients(who have mutations in the MYO7A gene) using high-resolutionoptical coherence tomography (Jacobson et al., 2009) showed thatmorphological changes in the OLM (the apical terminus of Müllercell processes) marked a transitional zone between regions in whichphotoreceptors were diseased or depleted and morphologicallynormal regions, suggesting that such alterations in OLM integritymight precede photoreceptor degeneration. Our understanding ofthe role that the OLM plays in maintaining photoreceptor integrityis incomplete; however, numerous junction proteins, includingmembers of the Crumbs complex, have been shown to localize tothe OLM (for reviews, see Gosens et al., 2008; Omri et al., 2010)and, in particular, to the subapical region, perhaps contributing tojunctions within a semipermeable barrier. Mutations in Crumbsorthologs in mammals cause progressive retinal degeneration andthe retinal pathology reveals diminished contacts betweenphotoreceptors and the OLM (Mehalow et al., 2003; van de Pavert,2004). USH proteins have not been reported in the OLM, or inMüller cells in general, before this study, although whirlin (USH2D)has been linked biochemically to the Crumbs pathway (Gosens etal., 2007).

Although the extent to which Müller cells might contribute tothe USH retinal pathology remains to be investigated, ourhistological analyses of both zebrafish and mouse retinas, and ourfunctional studies in ush1c morphants and mutants, suggest thatadditional investigation of Müller cell function is required tounderstand the complex etiology of USH. As gene replacementtherapies are developed for USH patients, it will be important toknow whether retinal cell types in addition to photoreceptorsshould be targeted for optimal rescue of the vision defects. Theability to isolate Müller-cell-associated pathology in our zebrafishUSH1C model will allow further investigation of the roles ofMüller cells as well as investigations of USH protein function atthe OLM.

In summary, we have characterized the hearing and vision defectsin a zebrafish ush1c mutant and morpholino knockdown. Ourfindings support a phylogenetically conserved developmental rolefor harmonin isoform A in hair bundle organization, implicateMüller cells as contributors to retinal dysfunction andphotoreceptor cell death, and demonstrate that visual defects canbe detected from the onset of vision in these zebrafish models.

METHODS

Animal strains and maintenance

All zebrafish studies were conducted with Oregon AB wild-typelarvae or ush1cfh293 mutants in the AB background. Animals wereraised in a 10-hour dark and 14-hour light cycle, and maintainedas described (Westerfield, 2007). Owing to the swimming andbalance defects exhibited by ush1c mutants and morphants, allexperimental and control larvae for experiments occurring up to8 dpf were raised without food so as not to give normal swimmersa nutritional advantage over their swimming-impaired siblings.Animals were staged according to the standard series (Kimmel etal., 1995), or by hpf or dpf. All animal-use protocols were IACUCapproved.

Cloning of zebrafish ush1c cDNA and sequencing

tBLASTn queries with the human harmonin protein sequence wereused to identify the zebrafish sequence, and primers based on thissequence were used to amplify the full-length open reading frameof isoform A and partial sequence from additional isoforms fromwhole larvae or adult retinal cDNA (see supplementary material TableS1 for primer sequences). PCR products were purified using gelelectrophoresis and sequenced on an ABI 3100 using the fluorescentdideoxy terminator method. The gene is cataloged in NCBI underaccession number NC_007136. All novel transcripts were submittedto the NCBI dbEST, with accession numbers HO243997-HO244001and HS587020. Gene and protein names follow the zebrafishnomenclature guidelines (https://wiki.zfin.org/display/general/ZFIN+Zebrafish+Nomenclature+Guidelines).

Genetic mapping of zebrafish ush1cThe zebrafish ush1c locus was mapped by PCR using the LN54collection of radiation hybrids (obtained from Marc Ekker,University of Ottawa), which contain the zebrafish AB9 cell DNAand mouse B78 cell DNA (Hukriede et al., 1999).

ush1c mutant discovery

Labeled primers (supplementary material Table S1) to PCR amplifya 500 bp region surrounding exon 5 of ush1c were generated andused to screen 8448 genomes in a TILLING screen as described(Moens et al., 2008). Sperm from the identified carrier sample werethawed and used to fertilize wild-type AB eggs. Adult F1 carriers ofthe mutation were identified by fin-clip genotyping, and identifiedheterozygotes were crossed to produce homozygous offspring.

Morpholino injections

Antisense morpholinos (GeneTools, Philomath, OR) were injectedinto one-cell-stage embryos as described (Nasciveus and Ekker,2000). Embryos were injected with 1.8-2.0 nM of ush1c MO1targeted to the splice donor site of exon 2 (5�-TAGAATTG -AGACTTACTGATGGTAC-3�), or 0.6-1.0 nM of ush1c MO2targeted to the translation start site (5�-TTTTCAGTCCGT -TGGTCGTCTTGCT-3�).

RNA isolation and RT-PCR

Total RNA was isolated from euthanized larvae at 24 hpf to 7 dpfusing Trizol (Invitrogen, Carlsbad, CA). Reverse transcription wasperformed according to manufacturer instructions using theSuperscript Reverse Transcriptase II kit (Invitrogen).


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Western blot

Embryos were homogenized in 10 mM Tris-HCl, pH 7.4, 150 mMNaCl, 1 mM EDTA, 0.1% Triton X-100, 0.1% SDS and 5 mg/mlprotease inhibitor cocktail (Pierce, Rockford, IL). Samples were runon an 8% acrylamide gel and transferred to nitrocellulosemembranes with Nupage reagents (Invitrogen). Membranes werewashed in PBS twice, each for 15 minutes, incubated in blockingsolution (3% NFDM in PBS) for 1 hour at room temperature, thenin 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 5minutes each in PBS. HRP was detected using the ECL plusdetection system (Pierce). To detect actin, membranes wererehydrated and incubated in donkey anti-mouse HRP for 1 hour,rinsed three times for 5 minutes each in PBS, and labeled with theECL plus detection system (Pierce).

In situ hybridization and immunohistochemistry

In situ hybridization on whole and sectioned tissue and antibodylabeling on sectioned tissue was performed as described (Jensenet al., 2001; Ebermann et al., 2010). For imaging of subcellularprotein localization in sectioned retinas, incubation with primaryantibodies and a goat anti-mouse Alexa-Fluor-488 to visualize theglutamine synthetase antibody was followed by incubation with ananti-rabbit biotinylated antibody diluted in block (10% NGS, 2%BSA in PBS-T) overnight at 4°C. Sections were then rinsed fourtimes for 5 minutes each in PBS and signal was detected with theVectastain ABC Elite kit (Vector Laboratories, Burlingame, CA)and TSA kit (Perkin-Elmer, Waltham, MA). For whole-mountimaging 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 timesfor 5 minutes each in PBS + 0.01% Triton X-100, and thenpermeabilized with 2.5% Triton X-100 in PBS at room temperatureuntil otoliths dissolved. Larvae were rinsed four times for 5 minuteseach in PBS + 0.01% Triton X-100, blocked for 2 hours at roomtemperature in 10% NGS, 10% BSA in PBS + 0.01% Triton X-100,then incubated with Alexa-Fluor-488–phalloidin diluted in blockfor 24 hours at 4°C. Following four 5-minute PBS + 0.01% TritonX-100 washes, larvae were blocked using the Avidin/Biotin blockingkit (Molecular Probes, Carlsbad, CA), and incubated in antibodysolutions overnight at 4°C. Larvae were rinsed four times for 30minutes each in PBS-Triton, followed by signal detection with theVectastain ABC Elite kit (Vector) and TSA (Perkin-Elmer) kit. Afterrinsing four times for 5 minutes each in PBS-Triton, larvae wereimmersed in Vectashield (Vector) and mounted in a lateralorientation on chambered slides. Images were captured on a ZeissLSM5 confocal or BioRad Radiance 2100 confocal microscope. Thefollowing antibodies and concentrations were used in this study:rabbit anti-harmonin (SDIX) antibody generated against aa1-100of zebrafish harmonin, 1:200 for sections, 1:1000 for whole mount;rabbit anti-harmonin (SDIX) antibody generated against aa306-363of mouse harmonin, 1:200; goat anti-harmonin (Santa CruzBiotechnology, Santa Cruz, CA) against the N-terminus of thehuman protein, 1:40; rabbit anti-harmonin (Sigma-Aldrich, StLouis, MO) against aa42-118 of human harmonin, 1:40; mouse anti-

glutamine synthetase (Millipore), 1:500; mouse zpr1 (ZebrafishInternational Resource Center, Eugene, OR), 1:200; mouse anti-SV2(DSHB), 1:2000; rabbit anti-GluR2/3 (Millipore), 1:50; rabbit anti-active caspase-3 (BD Biosciences, San Jose, CA), 1:200; mouse anti-acetylated tubulin (Sigma), 1:500. Goat anti-mouse, chicken anti-goat and goat anti-rabbit Alexa-Fluor-488, -568 and -633 secondaryantibodies (Molecular Probes) were used at 1:300 and the anti-rabbit biotinylated antibody was used at 1:500. Alexa-Fluor-488–phalloidin (Invitrogen) was diluted to a final concentration of2.5 M.

Electron microscopy

Tissue was prepared as described previously (Schmitt and Dowling,1999). Ultrathin sections were obtained from Epon-embeddedtissue, stained with uranyl acetate and lead citrate, and examinedby conventional transmission electron microscopy.

Optokinetic response

To measure optokinetic response, we used a modification of themethod of Brokerhoff (Brokerhoff, 2006). Larvae were placed in asmall Petri dish containing 3% methylcellulose and mounted on astationary platform surrounded by a rotating drum of 8 cm indiameter. The drum interior displays ten alternating black and whitestripes, each subtending an angle of 28°. The larvae were orientedin an upright position, illuminated with fiber optic lights and viewedthrough a dissecting microscope positioned over the drum, whichrotated at 5 rpm clockwise and then counter clockwise, for 30seconds in each direction. Eye movements were recorded duringthis time period, after which the larvae were washed out of themethylcellulose and returned to embryo medium.

ERG recordings

ERG recordings (Makhankov et al., 2004) were made to examineretinal function of morphants (morpholino-injected larvae)compared with control larvae. 5 dpf larvae were incubated inRinger’s solution for at least 1 hour prior to the experiment anddark-adapted for a minimum of 30 minutes. Under dim redillumination, larvae anesthetized with tricaine (Sigma Aldrich) andparalyzed with 300 M pancuronium bromide (Sigma Aldrich)were placed on a piece of sponge so that the pupil of one eye wason axis with the stimulus light beam. A recording electrode wasplaced on the cornea to measure the change in voltage induced bystimulation. The reference electrode was a AgCl-coated silver wireinserted into the sponge. The recording electrode was connectedvia a AgCl-coated silver wire to a low noise recording amplifier(Axopatch 200A with a CV201A head stage), and data wereacquired and stored digitally (Digidata 1200 interface and pClamp7and pClamp8 software; recording hardware and software fromAxon Instruments). Signals were filtered at 2 kHz.

StimulationRecordings were performed in a Faraday cage housing a Zeiss UEMmicroscope, with a 4� objective lens used for visualizing the eyeand delivering light stimuli. An ultra bright white LED array (Atlasseries high brightness module NT-42D0-0426 from LaminaLighting, with an LED driver circuit locally designed andconstructed by the technical support group) was used to presentthe stimulus. Stimulus durations were 800 mseconds. The light



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intensity of the unattenuated beam was 1500 W/cm2. Appropriateneutral density filters were inserted into the light beam to achieveaccurate light stimulus intensities over a 5 log10 unit range. Threeto five responses per intensity level were averaged and filtered for60 Hz noise.

Isolation of the a-wave componentDirectly after recording the b-wave (in Ringer’s solution),pharmacological compounds were applied externally to the eyeusing an Eppendorf pipette. The compounds used in this study were

L-(+)-2-amino-4-phosphonobutyric acid (L-AP4; 30-300 M), DL-threo-b-Benzyloxyaspartic acid (DL-TBOA; 15 M) and Ringer’ssolution with 5% DMSO as control. After 15-30 minutes, the a-wave traces were recorded.

Data analysisThe amplitudes of the b-waves were measured from the bottom ofthe a-wave to the peak of the b-wave; the a-wave was measuredfrom the baseline to the peak of the a-wave. We used larvae thathad a consistent response to a test flash delivered (±10%) at thebeginning and the end of the session. Unhealthy or damaged wild-type larvae that showed a decreased ERG or changed waveform atthe end of the experiment compared with the beginning were notincluded in the analysis. For statistics, we used a linear mixed modelapproach and a Student’s t-test.

ACKNOWLEDGEMENTS

The authors thank David Raible, Kelly Owens and John Constable for help withexperimental design and reagents, Tom Titus and the Cecilia Moens laboratory fortheir work on the TILLING screen, and Jeremy Wegner for technical assistance.Support was provided by NIH EY07042, DC004186, DC010447 and HD22486, andby the Foundation Fighting Blindness. J.B.P. was supported by an American HeartAssociation 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 themanuscript with input from the other authors. B.B.-S. designed and performedexperiments and analyzed data. J.J.L. designed and performed experiments andcontributed experimental reagents. A.T., K.K. and M.M. performed experiments andanalyzed data. S.S., Z.G.J. and P.F.H. performed experiments. D.S.W., P.W., B.J.K. andM.W. conceived and designed the experiments.

SUPPLEMENTARY MATERIAL

Supplementary material for this article is available athttp://dmm.biologists.org/lookup/suppl/doi:10.1242/dmm.006429/-/DC1

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TRANSLATIONAL IMPACT

Clinical issueUsher syndrome (USH) is a recessively inherited disease of combined deafnessand blindness. Hearing impairment is usually apparent from birth, whereasvision loss is slow and progressive, beginning in the first or second decade oflife. Cochlear implants and hearing aids can compensate for some of thehearing deficits, but there is currently no treatment to counter the vision loss,which is a form of retinitis pigmentosa. Early diagnosis is important for geneticcounseling and preparation for late onset blindness.

There is a great deal of clinical variability among patients with USH, andnine genes that can cause this disease when mutated have been identified todate. These genes encode a wide variety of proteins that have complex anddynamic localization patterns within the cells of the inner ear and the retinathat are affected in USH. Discovering more about how these mutations causedeafness and blindness at the cellular and molecular level will enhance ourunderstanding of the etiology of this disease, and contribute to thedevelopment of effective treatments.

ResultsIn this paper, the authors describe a zebrafish model for Usher type 1C(USH1C), a severe form of USH in humans that is characterized by profoundcongenital deafness, balance problems and a relatively early onset ofprogressive vision loss. The authors disrupt the normal function of the ush1cgene by using morpholino oligonucleotides to interfere with mRNA splicing. Inaddition, they identify an ush1c zebrafish mutant by screening DNA fromindividual mutagenized fish to find a lesion in the ush1c gene. Bothmorpholino-injected animals and ush1c mutants have distinct defects inhearing, balance and vision that are apparent in the first days of life. Theauthors examine the sensory cells involved in these behaviors –mechanosensory hair cells and retinal cells – and discover defects in theirdevelopment and function. In the ear, the structural portion of themechanosensory hair cells that are important for intercepting sound wavesdoes not form properly. Surprisingly, the authors discover that, in the retina,the ush1c gene functions in Müller glia rather than in photoreceptors, the celltype that has been reported to express Ush1c in mammals. Subsequently, theyshow that this gene is also expressed in mammalian Müller glia. Importantly,although zebrafish ush1c is not active in photoreceptors, loss of ush1c functionin Müller cells leads to compromised photoreceptor function and cell death.

Implications and future directionsFurther insights into the role of ush1c in the development and function ofsensory cells are vital to providing early diagnosis and intervention to forestallthe devastating effects of this disease. This work contributes to theunderstanding of USH1C pathology by providing information about thespecific molecules involved in the formation of stereocilia in mechanosensoryhair cells, and by identifying a new retinal cell type through which ush1cfunctions to facilitate and maintain synaptic connections betweenphotoreceptor cells and interneurons of the retina, namely Müller glia. USHhas traditionally not been considered a developmental disorder of vision. Theresults of this study, however, show that visual defects in the zebrafish modelof USH1C can be detected from the onset of vision, and thus might haveclinical relevance for the early diagnosis of USH1C in humans.

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