otopetrin 1 is required for otolith formation in the zebrafish danio rerio

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Developmental Biology

Otopetrin 1 is required for otolith formation in the zebrafish Danio rerio

Inna Hughesa,1, Brian Blasioleb,1, David Hussc, Mark E. Warchold, Nigam P. Rathe, Belen Hurlef,

Elena Ignatovad, J. David Dickmanc, Ruediger Thalmannd, Robert Levensonb, David M. Ornitza,*

aDepartment of Molecular Biology and Pharmacology, Washington University Medical School, St. Louis, MO 63110, United StatesbDepartment of Pharmacology, Pennsylvania State University College of Medicine, Hershey, PA 17033, United States

cDepartment of Anatomy and Neurobiology, Washington University Medical School, St. Louis, MO 63110, United StatesdDepartment of Otolaryngology, Washington University Medical School, St. Louis, MO 63110, United States

eDepartment of Chemistry and Biochemistry, University of Missouri, St. Louis, MO 63110, United StatesfNational Institutes of Health, National Human Genome Research Institute, Bethesda, MD 20892-2152, United States

Received for publication 9 July 2004, revised 30 August 2004, accepted 2 September 2004

Available online 25 September 2004

Abstract

Orientation with respect to gravity is essential for the survival of complex organisms. The gravity receptor is one of the phylogenetically

oldest sensory systems, and special adaptations that enhance sensitivity to gravity are highly conserved. The fish inner ear contains three large

extracellular biomineral particles, otoliths, which have evolved to transduce the force of gravity into neuronal signals. Mammalian ears

contain thousands of small particles called otoconia that serve a similar function. Loss or displacement of these structures can be lethal for

fish and is responsible for benign paroxysmal positional vertigo (BPPV) in humans. The distinct morphologies of otoconial particles and

otoliths suggest divergent developmental mechanisms. Mutations in a novel gene Otopetrin 1 (Otop1), encoding multi-transmembrane

domain protein, result in nonsyndromic otoconial agenesis and a severe balance disorder in mice. Here we show that the zebrafish, Danio

rerio, contains a highly conserved gene, otop1, that is essential for otolith formation. Morpholino-mediated knockdown of zebrafish Otop1

leads to otolith agenesis without affecting the sensory epithelium or other structures within the inner ear. Despite lack of otoliths in early

development, otolith formation partially recovers in some fish after 2 days. However, the otoliths are malformed, misplaced, lack an organic

matrix, and often consist of inorganic calcite crystals. These studies demonstrate that Otop1 has an essential and conserved role in the timing

of formation and the size and shape of the developing otolith.

D 2004 Elsevier Inc. All rights reserved.

Keywords: Otolith; Otoconia; Otopetrin 1 (Otop1); Biomineralization; Vestibular systems

Introduction

Otoconia are small (approximately 10 Am) extracellular

biomineral particles found in the vestibular portion of the

vertebrate inner ear. Otoconia are composed of specific

polymorphs of calcium carbonate (CaCO3) crystals precipi-

tated around an organic core of extracellular matrix proteins.

0012-1606/$ - see front matter D 2004 Elsevier Inc. All rights reserved.

doi:10.1016/j.ydbio.2004.09.001

* Corresponding author. Department of Molecular Biology and

Pharmacology, Washington University School of Medicine, 660 South

Euclid Avenue, PO Box 8103, St. Louis, MO 63110. Fax: +1 314 362 7058.

E-mail address: [email protected] (D.M. Ornitz).1 These authors contributed equally to this work.

These particles are required for normal sensation of linear

acceleration and gravity in mammals (Bergstrom et al., 1998;

Lim, 1980; Ornitz et al., 1998). In teleost fish, complete loss

of the orthologous structure, the otolith, is lethal (Riley and

Moorman, 2000). In contrast to the thousands of small

otoconial particles in mammals, only three large otoliths

form in fish. Few molecules governing the development of

otoconia and otoliths have been described and those that

have seem to be specific to either structure. It has been

proposed that the polymorph of CaCO3 found in otoconia or

the otolith is determined by the major matrix proteins that

make up the organic core: Otoconin 90 is at the core of

calcitic CaCO3 otoconia of birds and mammals; Otoconin 22

276 (2004) 391–402

I. Hughes et al. / Developmental Biology 276 (2004) 391–402392

is the primary matrix component of aragonitic CaCO3

otoconia in amphibians; Otoconin 54 is the primary matrix

constituent of the vateritic CaCO3 otoconia utilized by early

jawed fish, such as the garfish (Pote and Ross, 1991); and

otolith matrix protein (omp) is the primary matrix protein of

the aragonitic fish otolith (Murayama et al., 2000). These

major matrix proteins share the ability to bind calcium or

other ions and the otoconins share one or two rigid

phospholipase A2 structural domains that may mediate

their ability to guide the formation of specific CaCO3 crystal

polymorphs (Pote et al., 1993; Wang et al., 1998).

Starmaker, an ortholog of the mammalian dentin sialopro-

tein (DSP), is an acidic phosphoprotein recently shown to

be required for normal otolith formation in the zebrafish

(Sollner et al., 2003). While expression of DSP has been

observed in the mouse inner ear, DSP knockout mice do not

appear to have significant vestibular dysfunction based on

swim testing (T. Sreenath, personal communication),

suggesting that the function of starmaker may be specific

to the fish otolith.

While there must be important differences in the proteins

and pathways required to generate small otoconia particles

versus a large otolith, some similarities must also exist. The

essential requirement for the formation of both otoliths and

otoconia is the availability of Ca2+ and CO32� ions. The

presence of carbonate ions depends on the activity of

carbonic anhydrase. The source of calcium in the endolymph

that contributes to the otoconia and the otolith is poorly

understood. Organic substances, including acidic proteins,

glycosaminoglycans (GAGs) and proteoglycans, are also

essential to regulate crystal growth (Addadi et al., 1989;

Khan, 1997) and have been identified in both otoliths

(Borelli et al., 2003) and otoconia (Tachibana and Morioka,

1992). While it is believed that these proteins and extrac-

ellular matrix molecules are required for locally increasing

Ca2+ and CO32� concentrations and as structural components

of the developing otolith or otoconia, the only protein with a

known enzymatic function required for otoconial formation

is NADPH Oxidase 3 (NOX3). NoX3 is mutated in head-tilt

mice that have nonsyndromic otoconial agenesis and may be

required for the aggregation of Otoconin 90 proteins in the

mouse ear (Paffenholz et al., 2004).

Orchestration of extracellular calcification requires bring-

ing together ionic and proteinaceous components in time

and space. The organic matrix components of otoconia are

expressed in different regions of the vestibular epithelium;

however, all matrix components must associate with an

extracellular gelatinous structure called the otolithic mem-

brane in order to localize otoconial development above the

sensory epithelium. Otoconial matrix proteins must aggre-

gate into ordered organic cores and Ca2+ and CO32�

concentrations must be locally increased to allow crystal-

lization. Coordination of these events requires the normal

formation of the otocyst (Malicki et al., 1996) and sensory

maculae (Haddon et al., 1998), as well as tight regulation of

the endolymph ionic environment (Everett et al., 1997;

Kozel et al., 1998). This process is temporally restricted, as

expression of certain major matrix proteins is dramatically

down-regulated after early development (B. Blasiole and E.

Ignatova, unpublished data). In mammals, the process of

otoconial development is essentially complete by postnatal

day 7 (Erway et al., 1986; Lim, 1973; Veenhof, 1969), and

little evidence is available for continued otoconial formation

or repair. In fish, initial rapid growth of the otolith occurs

early in otic development, but the otolith continues to grow

throughout the life of the fish, with increments of organic

matrix and calcium carbonate added daily to the otolith

surface. Disruption of any of these processes can lead to the

formation of abnormally shaped or ectopic otoconia or

otoconial agenesis.

Ectopic otoconia in humans have been proposed to cause

human vestibular dysfunction, in particular benign parox-

ysmal positional vertigo (BPPV). BPPV is a common cause

of dizziness and is associated with dislodged otoconia

entering the semicircular canals and causing abnormal

vestibular sensation in response to head rotation. It has

been estimated that in the elderly population as much as

50% of dizziness can be attributed to BPPV (Oghalai et al.,

2000). Otoconial pathology is thus a significant etiology of

balance-related falls and accidental deaths in the elderly.

While many cases of BPPV have been associated with head

trauma, vestibular neuritis, treatment with some pharmaco-

logical agents, or age-related degeneration of otoconia, the

etiology of approximately half of the cases of BPPV in

young and elderly patients is still unknown.

Mutations in a novel protein, Otopetrin 1 (Otop1), cause

nonsyndromic otoconial agenesis and a severe balance

disorder in tilted and mergulhador mice (Hurle et al.,

2003). Mutant mice display near one hundred percent

penetrance of the otoconial agenesis phenotype, with no

developmental changes in inner ear morphogenesis.Otop1 is

predicted to encode a multi-transmembrane domain protein

of unknown function with no known homology to any family

of receptors, transporters, or channels. Two independent

single-base pair mutations have been identified in mutant

mice; these mutations are in different regions of the molecule

but create identical phenotypes, suggesting that the normal

function of Otop1 is necessary for the development of

otoconia in the mouse. Homologous genes for Otop1 have

been identified in all vertebrate groups examined including

zebrafish and Fugu, where they are 41% and 44% identical,

respectively, to mouse Otop1 (Hurle et al., 2003), suggesting

that they may share a common mechanism of action in both

otolith and otoconial morphogenesis.

Here, we show that Otop1 has a conserved and essential

role in teleost otolith development. Zebrafish otop1 and

mouse Otop1 have similar expression patterns in the

developing inner ear. Morpholine oligonucleotide (morpho-

lino)-mediated knockdown of Otop1 expression resulted in

otolith agenesis in the majority of treated fish, without

affecting morphogenesis of the zebrafish inner ear. In a

small percentage of morphant animals, otolith development

I. Hughes et al. / Developmental Biology 276 (2004) 391–402 393

was greatly delayed and began at 40–50 h postfertilization

(hpf), with a variety of otolith phenotypes, including

formation of normal otoliths or otoliths lacking an organic

matrix and with an atypical crystal polymorph (calcite).

These data support a conserved role for Otop1 in the

localization and aggregation of otolith matrix proteins and

the regulation of the ionic environment of the otolithic

membrane during vestibular organ development.

Materials and methods

In situ hybridization

The clone fb76b02.y1 containing the 3V UTR of otop1

identified from the zebrafish EST project (Washington

University) was sequenced to identify orientation and

linearized with Not1. A digoxigenin-labeled RNA probe

was generated using the Sp6 labeling kit (Roche), following

the manufacturer’s instructions.

Timed matings of zebrafish were established and embryos

isolated at specific time points based on time since fertiliza-

tion and on developmental milestones (zfin.org). Embryos

were dechorionated and fixed overnight in 4% paraformal-

dehyde in phosphate-buffered saline (PBS) at 48C. Embryos

were hybridized overnight with a digoxigenin-labeled probe.

Hybridization was detected with anti-digoxigenin–alkaline

phosphatase-conjugated IgG and visualized with BM purple

AP substrate (Roche) in the presence of 2% levamisole.

For sectioning of in situ hybridized embryos, overstained

3 dpf embryos were embedded in JB-4 plastic (Chan et al.,

2001) and sectioned at 4 Am. Similarly prepared embryos

were sectioned and stained with Richardson’s stain for

comparison of histologic structures.

Morpholine oligonucleotide-mediated knockdown of

zebrafish otop1

Two independent morpholine oligonucleotides were

designed to base pair with the 5V UTR of otop1. MO-1

covers the region from �91 to �65 bp from the ATG start

codon (TTACACCTTCAGGACCCGTTAGTTT) and MO-

2 begins at �20 bp and spans the ATG, ending at the +5

nucleotide position (ACCATGCTCGATCGCTGTCGGTA-

AA). Both morpholinos were purchased from Gene Tools,

LLC (Philomath, Oregon) and were 5V labeled with FITC.

Before injection, morpholino oligonucleotides were diluted

with Phenol Red tracer and 1� Danieau’s buffer (58 mM

NaCl, 0.7 mM KCl, 0.4 mM MgSO4, 0.6 mM Ca(NO3)2,

5 mM HEPES, pH 7.6). Timed matings were established.

Embryos were collected at the 1-and 2-cell stage and

injected with morpholino concentrations from 0.25 to 12 ng.

MO-1 was used for all of the following experiments. Dorsal

ventral axis defects and pericardial edema were both noted

to occur in morpholino-injected fish at high doses (8–12 ng

morpholino).

Morphant embryos were maintained at 288C in charcoal

filtered water and were examined and counted under a

dissection microscope. To count and photograph otoliths at

older stages, embryos were anesthetized with 1% tricaine

and immobilized in 3% methyl cellulose.

Histologic preparation

The 4 and 7 dpf wild-type and morphant fish were

collected and fixed overnight in cold 2.5% glutaraldehyde in

0.1 M sodium cacodylate buffer, pH 7.5. Animals were

washed in cold 0.1 M cacodylate buffer and dehydrated by

graded series of 25–100% ethanol. Samples were main-

tained at �208C until they could be further processed. To

prepare samples for thin sections, embryos were cleared in

100% propylene oxide and incubated with an increasing

ratio of propylene oxide to unaccelerated Durcupan at room

temperature. Fish were then incubated with 100% unaccel-

erated Durcupan overnight and transferred to accelerated

Durcupan, oriented and hardened at 608C for 48–64 h.

Sections were cut with a glass Ralph knife on a rotary

microtome at a thickness of 4 or 6 Am. Sections were stained

with Richardson stain and coverslipped.

For immunohistochemical labeling, 28 hpf morpholino-

treated and control fish were fixed overnight in 4%

paraformaldehyde. Following thorough rinsing in PBS,

specimens were cryoprotected in 30% sucrose (in PBS),

embedded in OCT compound, frozen, and sectioned at

10 Am. Sections were mounted on gelatin/CrK(SO4)2�-

coated slides, treated for 2 h in blocking solution (2%NHS,

2% NGS, 1% BSA, in PBS), and incubated overnight in

primary antibody against acetylated tubulin (Sigma T6793,

1:100). The next day, sections were incubated for 2 h in Cy3-

conjugated anti-mouse secondary antibody and cell nuclei

were labeled with bisbenzimide (Sigma).

Scanning electron microscopy

Fixed tissues were dehydrated using a series of graded

acetones. Final dehydration was performed by placing the

specimens in tetramethylsilane that was sublimated in a dry

608C oven. The specimens were then mounted on studs and

palladium coated.

Single crystal X-ray diffraction

The 7 dpf wild-type and morphant fish from the same

clutches were anesthetized and fixed in 100% ethanol.

Otoliths were removed by dissection in 100% ethanol and

were maintained dry on a covered glass slide until examina-

tion. The otoliths were mounted with grease in a random

orientation. Preliminary examination and data collectionwere

performedwith aBruker SMART1KChargeCoupledDevice

(CCD) Detector single crystal X-ray diffractometer equipped

with a sealed tube X-ray source using graphite monochro-

mated Mo Ka radiation (k = 0.71073 2) at �1238C. Typical


preliminary unit cell constant determination with a set of 45

narrow frame 0.38E scans failed due to insufficient harvested

reflections. Therefore, a data set was collected with a frame

width of 0.38 inE and counting time of 60 s/frame at a crystal

to detector distance of 4.835 cm (1301 frames at�278 2h and

230 frames at 278 2h). The double pass method of scanning

was used to exclude any noise. Thresholding the collected

frames resulted in 122 reflections. Indexing of the unit cellwas

carried out with CELL_NOW (Sheldrick, 2002) and the cell

was refined using the SMART software package (Bruker

Analytical X-ray).

Results

Similar expression of mouse and zebrafish otop1 in the ear

during embryonic development

The zebrafish homologue of Otop1 was identified and

described previously (Hurle et al., 2003). Using whole-

Fig. 1. Whole mount in situ hybridization analysis of otop1 mRNA expression. (A)

the ventral half of the developing otic vesicle. (B) Dorsal view at 24 hpf showing

expression in the sensory epithelium. (D) Four-Am plastic section of the otocyst of

(100�). Pale apical cells with round nuclei (arrow) are hair cells. Cells with elo

macular growth zone. (E) Similar 4 Am plastic section of a 3 dpf fish showing otop1

monolayer, consistent with expression in the mature and developing hair cells (10

entire length of the animal in the anterior and lateral line organs (arrowheads). S

mount in situ hybridization, otop1 mRNAwas first identified

in the otocyst at 18 h postfertilization (hpf). By 24 hpf, when

otolith seeding is nearly complete, otop1 expression was

localized in the developing sensory epithelium of the ear

(Figs. 1A and B). This pattern is similar to Otop1 expression

in the mouse utricle and saccule during otoconial develop-

ment (Hurle et al., 2003). At later stages, zebrafish otop1

mRNA was restricted to the utricular and saccular maculae

(Fig. 1C) in a pattern consistent with expression in precursor

and mature sensory hair cells (Figs. 1D and E). By 5 days

postfertilization (dpf), otop1 expression was greatly reduced

in the otolith organs but was identified in the neuromasts of

the lateral line system (Fig. 1F). Expression of otop1 was not

detectable in the inner ear at 7 dpf by in situ hybridization but

persisted in the anterior and lateral line (data not shown).

Reduction or loss of otop1 expression in the inner ear

suggests that otop1 has a specific role in the early develop-

ment and rapid growth of the otolith, but that it may not be

required for the daily incremental growth that continues

throughout the life of the fish.

Lateral view of a 24 hpf embryo showing significant expression of otop1 in

otop1 expression. (C) Lateral view of 3 dpf larva showing otop1 mRNA

a 3 dpf fish (dorsal is up, lateral to the right) stained with Richardson’s stain

ngated, densely staining nuclei (arrowheads) are precursor cells within the

mRNA localized to the luminal cells of the otocyst and adjacent cells in the

0�). (F) Lateral view of a 5 dpf larva showing otop1 expression along the

cale bars: A, 50 Am; B–C, 250 Am; D–E, 10 Am; and F, 50 Am.

Table 1

Otolith formation in otop1 morphant fish

Morpholino (ng) n Score (%)

0 1 2 3

A. Morpholino-mediated knockdown of otop1 30 hpf

0 53 100 0 0 0

0.25 15 67 0 0 33

0.5 46 20 2 0 78

1 29 7 0 0 93

2 42 0 0 0 100

4 24 0 4 0 96

8 56 0 0 0 100

10 50 0 0 0 100

B. Morpholino-mediated knockdown of otop1 7 dpf

0 23 100 0 0 0

1 33 9 36 31 24

2 17 0 0 12 88

4 23 0 0 9 91

Score: 0, 2 otoliths in each ear; 1, otoliths in both ears, but abnormal in

number, location, or shape; 2, otoliths in only one ear; 3, otoliths absent in

both ears.

Fig. 2. Absence of otolith formation in otop1 morphant fish. (A) Lateral view of an uninjected wild-type control at 30 hpf. (B) Lateral view of a 30 hpf

morphant fish injected with 10 ng MO-1. Otop1 morphant fish are morphologically normal at all stages of development but lack otolith formation. (C and D)

Lateral view of control and morphant 30 hpf otocysts. Otoliths are located at the poles of the developing otocyst of a control fish (arrows) (C) but are absent in

the 10-ng MO-1-injected animals (arrow) (D). Scale bars: A–B, 50 Am; and C–D, 250 Am.


Morpholino-mediated knockdown of otop1

To determine whether the essential function of Otop1 in

otoconial/otolith development is conserved in fish, the

expression of zebrafish Otop1 protein was knocked down

using antisense morpholine oligonucleotides (morpholi-

nos). Morpholinos targeted to 5VUTR and translation

initiation sequences block translation of the message

(Nasevicius and Ekker, 2000). Injection of a morpholino

designed against otop1 (MO-1) into one-cell stage embryos

resulted in complete agenesis of otoliths (Fig. 2). Injection

of a second morpholino (MO-2) targeted to an independent

region of the otop1 5VUTR reproduced this defect,

confirming that otolith agenesis in zebrafish was specific

to loss of Otop1 expression. At 30 hpf, more than 96% of

otop1 morphant fish failed to develop both the saccular and

utricular otoliths (Table 1), with no other obvious devel-

opmental defects (Fig. 2). Expression of pax2a and otx1 at

24 (Figs. 3A–D) and 48 hpf (data not shown) were

comparable in uninjected and otop1 morphant fish, an

indication that the early steps in otocyst formation pro-

gressed normally.

Tether cells function at the poles of the otocyst and are

required for normal otolith formation (Riley and Grunwald,

1996). Development of this cell type, examined by

acetylated tubulin immunohistochemistry of wild-type and

injected animals, showed distribution of hair cell kinocilia in

a similar pattern to uninjected, age-matched controls (Figs.

3E and F). These data suggest that otop1 knockdown does

not disrupt normal patterning or cell differentiation in the

developing inner ear, but specifically disrupts otolith

formation.

Fish with abnormal otolith development often have visible

otolith seeding particles within the otocyst by 24 hpf (Riley

and Grunwald, 1996; Riley et al., 1997; Sumanas et al., 2003)

even when actual otolith formation is delayed. No otolith

matrix material was observed in otop1 morphant fish at 24

hpf by differential interference contrast (DIC) microscopy

(Fig. 3G). To determine if the expression of genes known to

be required for formation of the otolith matrix was disrupted

in otop1 morphant fish, the expression of zebrafish otolith

matrix protein (omp) (E. Ignatova, unpublished data) and

starmaker (Sollner et al., 2003) mRNAwere assessed during

Fig. 3. Normal gene expression and otocyst morphogenesis in morphants. (A) The pax2a expression in wild-type and (B) morphant otocyst. (C) otx1

expression in a wild-type and (D) morphant otocyst. (E) Immunohistochemistry for acetylated tubulin on 10 Am frozen sections through a 28 hpf wild-type

otocyst. Tether cell kinocilia (arrow) and otolith (arrowhead) are evident in magnified insert (EV). (F) Morphant otocyst examined for acetylated tubulin. Tether

cell kinocilia are evident in magnified insert (FV) (arrow) but no otolith seeding particles were present. (G) DIC image of the otocyst of a morphant fish (100�)

at 24 hpf. No aggregated matrix was visible in several ears examined in this manner. (H) omp expression in wild-type and (I) morphant otocyst. (J) starmaker

expression in wild-type and (K) morphant otocyst. A–D and H–K are whole mount in situ hybridizations of 24 hpf embryos. All otop1 morphant fish were

injected with 4 ng MO-1 at the one-cell stage. Rostral is on the left. Scale bars: A–D, H–K, 250 Am; E–F, 10 Am; G, 25 Am.


early otic development. The expression patterns of omp and

starmaker were similar in wild-type and morphant fish at

24 hpf (Figs. 3H–K) and 48 hpf (data not shown), suggesting

that loss of Otop1 does not disrupt the normal expression of

genes encoding otolith matrix proteins.

Fig. 4. Normal formation of the lateral line system in otop1 morphant fish. (A–

neuromast. The cupula and kinocilia bundle has formed normally in the morphant a

(C) and 10 ng MO-1-injected morphant anterior neuromast showing a similar com

cells. Scale bars indicate 10 Am. C and D are stained with Richardson’s stain.

Normal formation of the lateral line system in otop1

morphants

Despite expression of otop1 in the developing neuro-

masts of the lateral line system, morphant animals showed

B) SEM image of a 4 dpf WT (A) and morphant (B) posterior lateral line

nimal. (C and D) Four-Am plastic section through a 7 dpf uninjected control

plement of hair cells (pale, apical cells with round nuclei) and supporting


no obvious defects in the morphogenesis of these structures.

Examination of the neuromasts of live morphants by DIC at

3 and 5 dpf (data not shown) and scanning electron

microscopy of 4 dpf morphants (Figs. 4A and B) revealed

normal formation of the cupula, the extracellular fibrous

matrix covering the neuromast that transduces movement to

the underlying hair cells. Plastic sections through 7 dpf

morphants stained with Richardson’s stain showed that the

distribution and number of anterior, trunk, and posterior

lateral line neuromasts was similar to that of wild-type

animals. In addition, otop1 morphant neuromasts appeared

to have a similar complement of support and hair cells (Figs.

4C and D). Neuromast formation and migration are unlikely

to be directly affected by otop1 knockdown, as these

structures develop later than the otolith. The normal

distribution of the neuromasts in 7 dpf morphant fish

Fig. 5. Morphogenesis of the otocyst and sensory epithelium. (A and B) Lateral vi

the utricular otolith is smaller and located in the rostral portion of the otocyst, the

ampullae (arrowhead) are similar in control and in 10 ng MO-1 morphant fish with

wild-type fish (100�) showing development of the hair cells (pale apical cells, arr

of a 7 dpf morphant saccular macula showing a similar distribution of cell types. (E

utricular otolithic membrane at 7 dpf. Removal of the otolith disrupted the wild-ty

absence of the otolith. The otolithic membrane is fibrous and connects the stereoci

10 Am; E–F, 10 Am.

suggests that there was no disruption of the normal

patterning or migration of the neuromast precursors. Addi-

tional examination of these sections showed no histological

difference between morphant and wild-type fish in any other

structure (data not shown).

Delayed otolith formation in otop1 morphant fish

A small percentage of morphant fish that had completely

failed to develop otoliths by 30 hpf exhibited otolith

formation at 4 dpf. The late formation of otoliths in these

fish occurred as a function of the morpholino dose injected

(Table 1). Visible otolith particles were first noted between

40 and 50 hpf (data not shown). This timing is consistent

with dilution of the morpholino by growth of the fish

(Nasevicius and Ekker, 2000) and possible reexpression of

ew of a 5 dpf wild-type (A) and morphant (B) otocyst. In the wild-type fish,

saccular otolith is larger and located centrally. The semicircular canals and

otolith agenesis. (C) Plastic section through the utricular macula of a 7 dpf

ow) and supporting cells (darker basal cells, arrowhead). (D) Plastic section

and F) Scanning electron micrographs of a wild-type (E) and morphant (F)

pe otolithic membrane. Morphant fish formed an otolithic membrane in the

lia of all hair cells in the macula. O, otolith. Scale bars: A–B, 250 Am; C–D,


Otop1. The process of delayed otolith formation appeared

similar to the normal formation of otoliths earlier in

development in that multiple small seeding particles

agglomerated and attached to the sensory epithelium (Riley

et al., 1997). However, in some cases, some seeding

particles did not attach to the sensory maculae and were

found lodged in the developing semicircular canals or were

free floating in the otic cavity (Figs. 6I and J).

By 5 dpf, the development of the zebrafish inner ear is

essentially complete, with the formation of the semicircular

canals and associated sensory maculae (Whitfield et al.,

2002). When compared to wild-type fish, morphant fish

injected with 10 ng of MO-1 had normal sensory maculae

and canal formation but lacked otoliths (Figs. 5A and B). At

7 dpf, the morphant epithelium had a normal distribution of

hair and supporting cells in the saccular sensory macula, as

well as normal transitional cells and thin nonsensory

epithelium (Figs. 5C and D). Scanning electron microscopy

was used to compare the size of the sensory maculae,

distribution of hair cells, and the formation of the otolithic

Fig. 6. Delayed otolith formation and dysmorphology in otop1 morphant fish. (A–F

MO-1-injected fish (B–F). (A) Wild-type fish develop two ovoid otoliths. (B) Del

location. (C) Otoliths that are identical in location, but with an oblong shape and d

wild-type otolith and attached to the saccular macula. The irregular shape and larg

cuboidal otolith located in the saccular sensory macula. (F) Multiple otoliths with p

to a sensory macula. (G) Four-Am plastic section through a 7 dpf 1 ng MO-1-injecte

macula. Richardson’s stain identifies concentric rings of organic matrix within the

showing an angular otolith on the utricular sensory patch with no obvious organic

from an 8-ng MO-1-injected fish. At 4 dpf, numerous irregularly shaped otoliths a

SEM image of the same 4 dpf morphant otocyst. Note the presence of aggregate

additional free-floating crystals on the otocyst wall (several otolith particles were lo

and J is 25 Am.

membrane. Formation of the gravity organ sensory maculae

and the otolithic membrane appeared normal in fish injected

with MO-1. In wild-type animals, the otolith had to be

removed to examine the underlying macular epithelium; in

the instances examined, this led to tearing of this fibrous

membrane (Fig. 6E). In morphant animals that did not form

an otolith, the membrane remained intact and in contact with

each hair cell (Fig. 6F). This arrangement of the fibrous

matrix of the otolithic membrane in fish is presumably to

simultaneously transduce the motion of the otolith to all hair

cells of the macula. In mammals and birds, hair cells do not

appear to directly contact the otolithic membrane matrix.

In birds and teleost fish, two other vestibular maculae

form during later larval stages: the lagena with an

accompanying otolithic/otoconial membrane (8–12 dpf in

zebrafish, with otolith formation beginning at 9 dpf; Bever

and Fekete, 2002; Riley and Moorman, 2000), and the

macula neglecta (17–20 dpf), which lacks an otolith

(Whitfield et al., 2002). The formation of the lagenar otolith

and the sensory structures of lagena and the macula neglecta

) Lateral views of 7 dpf otocysts (rostral to left) of a wild-type (A) and 1 ng

ayed otoliths in morphant fish can appear similar to wild type in shape and

istinct straight edges. (D) Formation of a single otolith that is larger than a

e size may indicate fusion of the early otolith seeding particles. (E) A single

olyhedral structures. The posterior-most otolith did not appear to be attached

d morphant otocyst showing a wild-type-like otolith attached to the saccular

otolith structure. (H) Four-Am plastic section through the opposite otocyst

matrix within the crystal. (I) Lateral view of a 4 dpf otocyst (rostral to left)

re found throughout the otic cavity and in the semicircular canal (arrow). (J)

s of crystals attached to the utricular and saccular otolithic membranes and

st in preparation). Scale bar for A–F is 250 Am; G–H is 10 Am; I is 250 Am;


occurs too late in development to be affected by otop1

morpholino injection into fertilized eggs (Nasevicius and

Ekker, 2000).

By 7 dpf, a variety of otolith phenotypes was noted in

otop1 morphant fish with late-forming otoliths (Table 1).

Some of the most striking examples were seen in fish

exposed to relatively low concentrations of the morpholino.

In animals injected with 1 ng of MO-1, observed pheno-

types ranged from two near normal otoliths in each ear (Fig.

6B) to a single large rounded otolith (Fig. 6D), to single and

multiple polyhedral forms (Figs. 6C, E, and F). In some

examples, these crystals resembled mammalian otoconia

with polyhedral shapes and sharp edges.

Interestingly, some fish displayed mixed phenotypes.

For example, in one otocyst, the morphant fish formed a

normally shaped saccular otolith (Fig. 6G) and a small

uncalcified utricular otolith that stained strongly with

Richardson’s stain (data not shown) (Richardson et al.,

1960). The organic matrix of the normally shaped otolith

stained lightly with Richardson’s stain, highlighting the

daily growth of the otolith by alternating deposition of

organic matrix and inorganic CaCO3. This demonstrates

that a normal appearing otolith can develop after a critical

window of development proposed to extend from 18 to

24 hpf (Riley et al., 1997). In the opposite ear, both the

utricular (Fig. 6H) and saccular otoliths (data not shown)

were roughly cuboidal. In these otoliths, no organic

matrix could be identified, and multiple histological

sections suggested that it was made up of a single

inorganic crystal. The structure of the large, polyhedral

otolith closely resembles the bgiantQ calcitic otoconia

described in several mouse mutants with defects in

otoconial synthesis (Erway and Grider, 1984; Lim et al.,

1978; Ornitz et al., 1998). Such a change in morphology

of the morphant otolith indicated a possible change in the

mechanisms of mineralization (see below).

In rare cases, animals injected with higher doses of

morpholino recovered otolith formation. Ectopic mineraliza-

tion was first noted in these animals at approximately 72 hpf.

In these instances, otolith particles did not aggregate well and

could be identified throughout the otic cavity, including in

the developing semicircular canals at 4 dpf (Figs. 6I and J).

Abnormal otoliths in otop1 morphants are composed of

calcite

Pure CaCO3 can form crystals with one of three

distinct crystalline polymorphs: calcite, aragonite, or

vaterite. At 7 dpf, wild-type otoliths (Fig. 7A) are

composed of thousands of aragonitic CaCO3 crystallites

arranged in multiple orientations over the surface of the

growing otolith. In contrast, otoconia contain an organic

core and a crystalline casing composed of calcite, the

most stable polymorph of CaCO3 (Carlstrom et al., 1953).

The crystalline appearance of otoliths that formed in

morphant fish at 7 dpf (Fig. 7B) suggested a possible

change in crystal polymorph from aragonite to calcite.

Single crystal X-ray diffraction of wild-type otoliths

yielded a crystalline dust diffraction pattern (Fig. 7C) that

is consistent with the disordered arrangement of aragonitic

crystallites that has been previously identified by powder

X-ray diffraction (Sollner et al., 2003). Notably, a set of

unit cell parameters consistent with published values for

aragonite (http://ruby.colorado.edu/smyth/min/aragonite.

html) was obtained by indexing harvested reflections.

Morphant otoliths, with a shape similar to wild-type

otoliths (Fig. 6G), gave a similar diffraction pattern (data

not shown). The polyhedral otoliths evident in some

morphants appeared similar in shape to mammalian

calcitic otoconia by scanning electron microscopy (SEM)

(Fig. 7B). Single crystal X-ray diffraction analysis of this

type of otolith yielded a single crystal diffraction pattern

with an identifiable unit cell at �1238C of the following:

a = 4.992 (6), b = 4.992 (1), c = 17.012(2) 2, a =

90.00(1), b = 90.01(1), c = 120.01(1), V = 366.8 (1) (Fig.

7D). These parameters match published values for calcite

(http://ruby.colorado.edu/smyth/min/calcite.html).

Discussion

Otopetrin 1 is essential for formation of both otoconia

and otoliths. Thus, Otop1 must function early in the otolith

and otoconial developmental pathway, prior to specification

of architecture and CaCO3 polymorphs of these divergent

structures. The presence of pure calcite crystals in morphant

animals that initiated otolith formation outside the critical

period of 18–24 hpf proposed by Riley et al. (1997)

suggests that the ions required for the biomineralization and

the proteins that control crystal growth are not coordinately

regulated in otop1 fish. These data also suggest that

zebrafish otop1 may regulate the ionic environment of the

otolith and that following dilution of the inhibitory effects of

the otop1 morpholino between 30 and 96 hpf, crystals form

in a purely inorganic manner. This is likely due to

temporally restricted expression of proteins that form the

initial seeding particles or the organic matrix of the otolith

during the initial rapid growth phase of the otolith during

early development. Interestingly, the crystalline patterns

observed in otop1 morphants are similar to those observed

in starmaker morphant fish (Sollner et al., 2003). This

suggests that disruption of a variety of components of the

otolith developmental pathway can trigger a default

mechanism, which leads to formation of inorganic crystals.

Under these conditions, formation of calcite, the most stable

polymorph of CaCO3, is favored.

Otop1 is the first described molecule that has a

comparable knockdown/mutant phenotype in the develop-

ing otolith/otoconia of fish and mice. Otolith development

appears to be exquisitely sensitive to the concentration of

otop1 protein, as doses as low as 0.5 ng of morpholino were

sufficient to cause agenesis of the otolith in 78% of injected

http:\\www.ruby.colorado.edu\&sim;smyth\min\aragonite.html

http:\\www.ruby.colorado.edu\smyth\min\calcite.html

Fig. 7. Calcitic otolith formation in otop1 morphant fish. (A) Scanning electron microscopy of the otic cavity of a 7 dpf uninjected age-matched control

showing a smaller utricular (left) and larger saccular otolith. Both otoliths are rounded and cover the entire sensory maculae. (B) Eight-ng MO-1-injected

morphant otic cavity at 7 dpf. Otoliths are angular. The sensory epithelium is visible below the utricular otolith. The morphant otoliths resembled inorganic

crystals instead of organic calcification. (C) The 3608 rotation image of a single crystal X-ray diffraction of a wild-type 7 dpf otolith. Little prominent

diffraction pattern is present, indicating that calcium carbonate aragonite crystallites are arranged in a dustlike mosaic pattern across the surface of the otolith.

(D) Single crystal X-ray diffraction of a morphant otolith showing that otoliths similar to those above (B) behave as a single crystal. The unit cell derived from

this diffraction pattern was consistent with the calcitic polymorph of calcium carbonate. Scale bar indicates 50 Am.


fish (Table 1). This suggests that Otop1 regulates a critical

step in otolith formation and that protein concentration may

be tightly regulated. For example, Otop1 may regulate the

function or localization of other proteins required for otolith

development. During mouse otoconial development, Otop1

is localized to the otolithic membrane (Hurle et al., 2003),

an extracellular gelatinous superstructure made up of many

proteins that supports otoconial formation and maintenance.

Location in the extracellular space is particularly surprising,

as Otop1 is predicted to be an integral membrane protein.

This may indicate the presence of the protein on membrane

bound vesicles called globular substance (Tateda et al.,

1998), which are thought to be precursors of otoconia

(Erway et al., 1986; Preston et al., 1975; Ross, 1979). In this

location, Otop1 could function as a channel or transporter,

regulating the contents or function of exocytotic vesicles or

may act as a structural protein required for the attachment or

nucleation of otoconia.

In mouse mutants for Otop1, loss of gravity sensation

results in relatively mild behavioral deficits under normal

conditions (Hurle et al., 2003; Ornitz et al., 1998).

Animals are unable to swim when dropped in water but

are able to walk and rear normally. They do not exhibit

circling or head tossing behavior, which has been

identified in animals with other types of vestibular defects.

This could be due to compensatory mechanisms to

maintain balance, including the use of visual cues,

semicircular canals, and the proprioceptive system. Zebra-

fish with abnormal otolith formation have difficulty

orienting to gravity and are often unable to swim and

feed (Mizuno and Ijiri, 2003; Riley and Grunwald, 1996;

Riley and Moorman, 2000; Riley et al., 1997). While the

behavioral phenotypes of morphant animals were not

specifically examined, several instances were noted in

which morphant fish that had developed apparently

normal otoliths were unable to orient dorsal side up,

even when lit from above. No circling behavior was

observed, though most of the fish were raised in relatively

shallow water to allow morphant fish to inflate their swim

bladders. We propose that the delay in otolith formation in

these animals may lead to deficits in the formation of

neuronal circuitry between otolith organs and the vestib-

ular nuclei. Morphant fish that did not develop otoliths

primarily rested on their side at the bottom of the well,

even at 7 dpf. Most fish had an intact startle response

indicating normal function of the lateral line organs.

Interestingly, some morphants with a single saccular or

even semicircular canal-located otolith were able to swim

efficiently when lit from above, but would tilt or turn

upside down when resting.


Human vestibular dysfunction is an increasing clinical

problem (National Institute on Deafness and Other Com-

munication Disorders, 2002). Degeneration or displacement

of otoconia is a significant etiology of age-related balance

disorders and benign paroxysmal positional vertigo (BPPV)

(Lim, 1984; Tusa, 2001). In addition, commonly used

pharmacological agents, such as aminoglycoside antibiotics,

can also lead to disruption of otoconial structure and

function (Johnsson et al., 1980; Takumida et al., 1997).

The presence of ectopic calcified particles in late-developing

otoliths of morphant fish resembles the pathology associated

with human BPPV (Figs. 6I and J). The phenotype may

provide a useful model to elucidate the mechanism leading

to ectopic otoconia in BPPV. In addition, the studies

presented here suggest that reactivating the expression of

OTOP1 in the ear of patients with vestibular dysfunction

may enhance the mineralization of remaining otoconial

particles and reestablish otoconial function. Further under-

standing of the role of Otop1 and other proteins required for

otoconial formation may assist in formulating therapeutic

approaches aimed at improving otoconial stability over time

and possibly facilitating otoconial regeneration, in addition

to adding to our knowledge of mechanisms of calcification

in this and other systems.

Acknowledgments

The authors would like to thank Keith C. Cheng at

Pennsylvania State University College of Medicine for the

use of microscopy and imaging equipment. This work was

funded by NIH grant DC02236 (D.M.O., R.T.), DC006283

(M.E.W.), and MH068789 (R.L.). We thank I. Thalmann, I.

Boime, and K. Lavine for critically reading the manuscript

and for insightful discussion and T. Nicolson for providing

the starmaker in situ hybridization probe.

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