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Otolith matrix proteins OMP-1 and Otolin-1 are necessary for normal
otolith growth and their correct anchoring onto the sensory maculae
Emi Murayamaa,b,*, Philippe Herbomelb, Atsushi Kawakamic,
Hiroyuki Takedac, Hiromichi Nagasawaa
aDepartment of Applied Biological Chemistry, Graduate School of Agricultural and Life Sciences,
The University of Tokyo, 1-1-1 Yayoi, Bunkyo, 113-8657 Tokyo, JapanbUnite Macrophages et Developpement de l’Immunite, Departement de Biologie du Developpement,
URA2578 du CNRS, Institut Pasteur, 25 rue du Dr Roux, 75724 Paris cedex 15, FrancecDepartment of Biological Science, Graduate School of Science, The University of Tokyo, 7-3-1 Hongo, Bunkyo, 113-0033 Tokyo, Japan
Received 19 October 2004; received in revised form 2 March 2005; accepted 21 March 2005
Available online 11 May 2005
Abstract
Fish otoliths are highly calcified concretions deposited in the inner ear and serve as a part of the hearing and balance systems. They consist
mainly of calcium carbonate and a small amount of organic matrix. The latter component is considered to play important roles in otolith
formation. Previously, we identified two major otolith matrix proteins, OMP-1 (otolith matrix protein-1) and Otolin-1, from salmonid
species. To assess the function of these proteins in otolith formation, we performed antisense morpholino oligonucleotide (MO)-mediated
knockdown of omp-1 and otolin-1 in zebrafish embryos. We first identified zebrafish cDNA homologs of omp-1 (zomp-1) and otolin-1
(zotolin-1). Whole-mount in situ hybridization then revealed that the expression of both zomp-1 and zotolin-1 mRNAs is restricted to the otic
vesicles. zomp-1 mRNA was expressed from the 14-somite stage in the otic placode, but the zOMP-1 protein was detected only from 26-
somite stage onwards. In contrast, zotolin-1 mRNA expression became clear around 72 hpf. MOs designed to inhibit zomp-1 and zotolin-1
mRNA translation, respectively, were injected into 1–2 cell stage embryos. zomp-1 MO caused a reduction in otolith size and an absence of
zOtolin-1 deposition, while zotolin-1 MO caused a fusion of the two otoliths, and an increased instability of otoliths after fixation. We
conclude that zOMP-1 is required for normal otolith growth and deposition of zOtolin-1 in the otolith, while zOtolin-1, a collagenous protein,
is involved in the correct arrangement of the otoliths onto the sensory epithelium of the inner ear and probably in stabilization of the otolith
matrix.
q 2005 Elsevier Ireland Ltd. All rights reserved.
Keywords: Biomineralization; Matrix protein; Morpholino; Otolith; Zebrafish
1. Introduction
The hair cell-containing epithelia of the vertebrate inner
ear are the sensory endorgans of the vestibular and auditory
systems. Otoliths are dense crystals composed of calcium
carbonate and an organic matrix that are primarily involved
in gravity sensing by the vestibular hair cells (Lowenstein,
1971). In teleosts, three otoliths, sagitta, lapillus and
0925-4773/$ - see front matter q 2005 Elsevier Ireland Ltd. All rights reserved.
doi:10.1016/j.mod.2005.03.002
* Corresponding author. Address: Institut Pasteur, 25 rue du Dr Roux,
75724 Paris cedex 15, France. Tel.: C33 1 45 68 86 21; fax: C33 1 45 68
89 21.
E-mail address: [email protected] (E. Murayama).
asteriscus, are contained in each of three inner ear sacs,
sacculus, utriculus and lagena, respectively. Each otolith is
anchored to a corresponding sensory epithelium, or macula,
via a gelatinous layer called otolithic membrane. By virtue
of its inertial mass, the displacement of the otoliths relative
to the sensory macula is able to generate the hair cell
depolarization by deflecting the sensory hair bundles
(Manning, 1924).
In zebrafish, the otic vesicle is formed by cavitation of
the otic placode around the 18-somite stage (18 h post-
fertilization—hpf). Right from this stage, two otoliths, the
future lapillus and sagitta, start to form at the anterior and
posterior poles of the otic vesicle by localized accretion of
Mechanisms of Development 122 (2005) 791–803
www.elsevier.com/locate/modo
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803792
precursor particles on the tips of kinocilia of specialized hair
cell precursors, hence called ‘tether cells’ (Riley et al.,
1997). This initial process of ‘otolith seeding’ occurs during
a period from 18 to 22 hpf (18- to 26-somite stage). Then,
precursor particles gradually disappear from the endo-
lymph, and otolith formation enters a phase of ‘otolith
growth’, from solute material. The first sensory hair cells of
the future maculae appear by 24 hpf close to the two
growing otoliths. By 72 hpf, the anterior otolith (lapillus)
exhibits an oval shape, whereas the posterior otolith
(sagitta) shows a disk-like structure. The third otolith, the
asteriscus, appears not before 9–17 days post-fertilization
(dpf), in an additional chamber of pars inferior, the lagena,
which individualizes from the sacculus at 15 dpf (Riley and
Moorman, 2000; Bever and Fekete, 2002). All chambers of
pars inferior connect until late stages, then the sacculus
becomes separated from the utriculus around 20 dpf
(Haddon and Lewis, 1996).
It is known that fish otoliths grow diurnally, and form
daily rings within their microstructure (Campana and
Neilson, 1985). Each ring is composed of an incremental
zone in which CaCO3 predominates, and a discontinuous
zone mainly composed of organic matrix (Watabe et al.,
1982). Thus, otolith growth is based on the alternate
deposition of calcium carbonate and proteins on the otolith
surface (Mugiya, 1987). Based on these findings, the
organic matrix has been considered to play important
roles in otolith formation. A number of proteins that
accumulate in otoliths or otoconia (the equivalent of otoliths
in tetrapods) have been identified from various animal
species. Otoconin-22 (Pote et al., 1993; Yaoi et al., 2003)
was identified in amphibian otoconia, while its homolog
Otoconin-90/95 was independently found in mammalian
otoconia (Wang et al., 1998; Verpy et al., 1999). Calbindin-
D28K was then identified in the otoconia of chick (Balsamo
et al., 2000) and lizard (Piscopo et al., 2003). Recently,
Sumanas et al. (2003) and Sollner et al. (2003) examined the
function of the chaperone protein GP96 and Starmaker
protein, respectively, in otolith formation in zebrafish.
Knockdown of GP96 resulted in an otolith seeding defect,
whereby the seeding particles did not adhere to the kinocilia
of the tether cells (Sumanas et al., 2003). In contrast, loss of
function of Starmaker protein led to a change in crystal
polymorph of the otoliths from aragonite to calcite, and thus
to a change in otolith morphology (Sollner et al., 2003).
We previously identified two major otolith matrix
proteins, OMP-1 (otolith matrix protein-1) and Otolin-1,
from the otoliths of two salmonid species. They are the
major components of EDTA-soluble and -insoluble matrix
protein fractions, respectively (Murayama et al., 2000,
2002). OMP-1 is a member of the transferrin family of
proteins and shows 40% similarity with the C-terminal half
of human melanotransferrin, a monomeric glycoprotein
identified in human melanoma cells and thought to play a
role in iron metabolism (Aisen and Leibman, 1972;
Woodbury et al., 1980). Otolin-1 is a collagenous protein
that belongs to the type VIII and X collagen family. So far,
these collagens were mainly reported to be present in non-
calcified tissues, such as basement membranes (type VIII;
Labermeier et al., 1983; Shuttleworth, 1997) and hyper-
trophic chondrocytes (type X; Schmid and Conrad, 1982).
We have previously shown by immunohistochemical
examination that OMP-1 and Otolin-1 proteins are co-
localized in otoliths of embryonic and adult rainbow trout
inner ears (Murayama et al., 2004). In addition, Otolin-1
was localized in fibrous material connecting otolith
primordia to the sensory epithelium, likely to be the otolith
membrane (Murayama et al., 2004). In the sacculus of the
adult inner ear, OMP-1 is synthesized by most epithelial
cells outside the sensory maculae, while Otolin-1 protein is
found in a restricted group of cylindrical cells located next
to the marginal zone of the sensory epithelium (Murayama
et al., 2004).
Here, in order to investigate the roles of OMP-1 and
Otolin-1 in otolith formation, we identified the homologous
genes from zebrafish and analyzed the knockdown pheno-
types in zebrafish embryos, using antisense morpholino
oligonucleotides (MO).
2. Results and discussion
2.1. Isolation of zebrafish cDNA homologs of omp-1
and otolin-1
In order to isolate zebrafish homologs of omp-1
(Murayama et al., 2000) and otolin-1 (Murayama et al.,
2002), we performed PCR using degenerate primers on
zebrafish inner ear cDNA (see Section 3). The PCR
produced cDNA fragments of approximately 400 bp for
zebrafish omp-1 (zomp-1) and 200 bp for zebrafish otolin-1
(zotolin-1), respectively. Sequence analyses revealed that
the predicted amino acid sequences had high degrees of
sequence identity with the rainbow trout OMP-1 (81.6%)
and the chum salmon Otolin-1 (84%), respectively. To
obtain the 5 0 end of each cDNA, we performed RACE
reactions and obtained additional sequence including a
translation initiation codon. The remaining sequences 3 0 to
the cloned cDNA fragments and the gene structures were
deduced using the zebrafish genome sequence (available at
http://www.ensembl.org/Danio_rerio/) and a GENSCAN
program (Burge and Karlin, 1997) (Fig. 1).
The deduced amino acid sequence of zebrafish OMP-1
(zOMP-1) shows 84% overall similarity with rainbow trout
OMP-1 (rtOMP-1, Fig. 1A). All the cysteine residues and N-
linked glycosylation sites are conserved between zOMP-1
and rtOMP-1. In addition, both proteins also show about
50% similarity to the C-terminal lobe (C-lobe) of human
melanotransferrin (hMTf, Fig. 1A). In particular, 12 out of
15 cysteine residues are conserved between zOMP-1 and the
C-lobe of hMTf, suggesting that the two proteins may
have similar folding and tertiary structure. It is known that
Fig. 1. Schematic representation of zOMP-1 and zOtolin-1 and their similarity to other known proteins. (A) Exon/Intron organization of the zebrafish omp-1
gene with the corresponding protein domain structure of zebrafish OMP-1 (zOMP-1), rainbow trout OMP-1 (rtOMP-1) and human melanotransferrin (hMTf);
white boxes, cDNA sequence determined in this study; light gray, sequence from zebrafish genome data. Inside the boxes depicting protein structure: green,
signal peptide; blue and blue-outlined bars, conserved and non-conserved amino acid residues involved in iron binding, respectively; red bars, potential N-
linked glycosylation sites; yellow bars, zinc binding consensus sequences. The similarities of rtOMP-1 and the C-lobe of hMTf (bolded) with zOMP-1 are
indicated by percentage. (B) Genome structure of the zebrafish otolin-1 gene, and the corresponding domain structure of zebrafish Otolin-1 (zOtolin-1), chum
salmon Otolin-1 (csOtolin-1), bluegill sunfish saccular collagen (bsSC), human C1q (hC1q), human type VIII (hCOL VIII) and X (hCOL X) collagens. White
boxes, cDNA sequence determined in this study; light gray, sequence from zebrafish genome data. Inside the boxes depicting protein structure: green,
signal peptide; bolded, collagenous domain (COL). Red bars indicate potential N-linked glycosylation sites. The similarity of all these proteins to zOtolin-1 in
the C-terminal non-collagenous (C-NC) domain is indicated by percentage. N-NC, N-terminal non-collagenous domain. (C) Multiple sequence alignment of
the C-NC domains. Identical and similar amino acid residues are shown in blue and gray bold types, respectively. The amino acid residues involved in calcium
binding are shown in red.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 793
the C-lobe of hMTf is unable to bind iron because of
changes in two of five iron-binding residues relative to
transferrin (Baker et al., 1992). zOMP-1 and rtOMP-1
conserve only one of these amino acid residues, indicating
that these molecules play a role other than iron transport
(Fig. 1A). On the other hand, both zOMP-1 and rtOMP-1
contain the consensus sequence for zinc binding, HEXXH,
found in many zinc-carrying metalloproteases such as
thermolysin (Jongeneel et al., 1989) (Fig. 1A).
The predicted amino acid sequence of zebrafish Otolin-
1 (zOtolin-1) shows 84% overall similarity with that of
chum salmon Otolin-1 (csOtolin-1; Murayama et al.,
2002) and 86% with that of bluegill sunfish saccular
collagen (bsSC; Davis et al., 1995), a probable homolog
of Otolin-1 in this species (Fig. 1B). Two potential
N-glycosylation sites are conserved among zOtolin-1,
csOtolin-1 and bsSC. These molecules belong to a family
of collagenous proteins that contains collagens type VIII
and X (Muragaki et al., 1991; Apte et al., 1991; Thomas
et al., 1991) and C1q (Reid, 1985). The N-terminal non-
collagenous (N-NC) domain is not significantly similar
among these proteins, neither among zOtolin-1, csOtolin-1
and bsSC, nor to other proteins thus far identified. In
contrast, their C-terminal non-collagenous (C-NC)
domains share about 60% similarity. Type VIII and X
collagens are non-fibrillar short chain collagens found in
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803794
basement membranes (type VIII; Kapoor et al., 1988) and
hypertrophic chondrocytes (type X; Schmid and Conrad,
1982). Their N-NC and C-NC domains form nodes, which
are important for the helical chain association (Chan et al.,
1995), while the collagenous (COL) domain forms the
interconnecting spacers. These molecules oligomerize
supramolecularly into a three-dimensional, hexagonally
arranged lattice (Sawada et al., 1990; Kwan et al., 1991).
The size of the collagenous domain of zOtolin-1 is about
half of that of type VIII and X collagens (Fig. 1B),
suggesting that the lattice size is smaller in the otolith than
in the basement membranes and the hypertrophic
chondrocytes.
Type X collagen is believed to provide a temporal
pericellular matrix during endochondral ossification, in
which cartilage is replaced by trabecular bone (Chan and
Jacenko, 1998). Although type X collagen is found at later
stages only in the context of fracture repair and osteoarthritis
(Grant et al., 1987; von der Mark et al., 1992), previous
reports indicate that type X collagen plays a role in
calcification at least at the level of cartilage-bone conver-
sion (Chan and Jacenko, 1998; Sutmuller et al., 1997).
Another link to calcification is the calcium binding property
of type X collagen (von der Mark et al., 1992; Kirsch and
von der Mark, 1991; Bogin et al., 2002). The amino acid
Fig. 2. Developmental expression of zebrafish omp-1 (zomp-1) and otolin-1 (zoto
zomp-1 transcripts are detected only in the otic vesicles (arrowheads). (C) Magnifi
ventro-posterior part (arrowhead). (D) zomp-1 in situ hybridization at 72 hpf. The e
and pm, outlined by dotted lines) than in the non-sensory parts of the epithelium (
vesicle of a 26-somite embryo labeled with an anti-OMP-1 antibody. Both otoliths
anti-OMP-1 antibody. The ventro-lateral epithelium posterior to the anterior macul
three increasingly deeper focal planes of the same ear; zotolin-1 transcripts are not
and ventral sides of the latter (I, arrowheads); am, anterior macula; pm, posteri
juxtaposed for focusing the two otoliths. All panels show lateral views, anterior to t
residues involved in Ca-binding are conserved between
zOtolin-1 and type X collagen (Fig. 1C), suggesting that
zOtolin-1 may act as a mediator for the interaction of
organic matrix with inorganic materials in otolith formation.
Interestingly, like type X collagen, MTf is also expressed in
chondrocytes in mouse and rabbit (Kawamoto et al., 1998;
Nakamasu et al., 1999). It is notable that the two proteins,
respectively, related to OMP-1 and Otolin-1 share a cellular
source and potential function in chondrogenesis.
2.2. Expression of zomp-1 and zotolin-1
We examined the distribution of zomp-1 and zotolin-1
transcripts in developing zebrafish embryos. zomp-1 mRNA
was first detected at the 14-somite stage (16 hpf) in the otic
placode (Fig. 2A), which forms at about that stage (Haddon
and Lewis, 1996). At 24 hpf, zomp-1 mRNA was detected
throughout the otic epithelium (Fig. 2B) with more intense
expression in the ventro-posterior side (Fig. 2C). At 72 hpf,
zomp-1 expression was most intense in a region outside the
maculae (Fig. 2D, arrowheads). zomp-1 transcripts were
never detected in any other tissues through the stages
examined in this study. In contrast to the intense expression
of zomp-1 mRNA from the onset of otocyst development,
the zOMP-1 protein was first immunodetected at
lin-1). zomp-1 in situ hybridization at 14-somite stage (A) and 24 hpf (B).
cation of the otic vesicle at 24 hpf. zomp-1 expression is more intense in the
xpression level of zomp-1 is lower at the anterior and posterior maculae (am
arrowheads). A dotted line indicates the outline of the otic vesicle. (E) Otic
are slightly labeled. (F) At 35 hpf, both otoliths are heavily stained with the
a is also labeled (arrowhead). (G–I) zotolin-1 in situ hybridization at 72 hpf;
detected in the anterior (G) nor in the posterior macula (H), but at the dorsal
or macula. In panels (E) and (F), two photos at different focal planes are
he left, dorsal to the top. Scale bar in (C) indicates 25 mm in panels (C) to (I).
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 795
the 26-somite stage (22 hpf) in both anterior and posterior
otoliths (Fig. 2E), suggesting that the translation of the
zomp-1 mRNA is blocked until the 26-somite stage. At 27
and 35 hpf, the otoliths were intensely labeled with the anti-
OMP-1 antibody, and the ventro-lateral part of the
epithelium just posterior to the anterior macula was also
labeled (Figs. 5B,2F). The absence of zOMP-1 immunola-
beling of the precursor particles and nascent otoliths before
the 26-somite stage suggests that zOMP-1 is mainly
involved not in otolith seeding, but in the subsequent
phase of otolith growth from solute material, which begins
by that stage (data not shown).
In contrast to zomp-1, zotolin-1 mRNA was first detected
faintly at 48 hpf in a restricted, postero-medial part of the
otic vesicle (data not shown), and became clear by 72 hpf,
although still at a low level, essentially at the ventral and
dorsal edges of the posterior macula (Fig. 2G,H,I). The
zOtolin-1 protein was immunodetected in the otoliths at the
same stage (Fig. 5F). This result is reminiscent of our
previous immunohistochemical study using chum salmon,
in which Otolin-1 expression was only detected in
transitional epithelial cells just adjacent to the sensory
epithelium (Murayama et al., 2002). Like zomp-1
expression, zotolin-1 expression was never observed in
any other tissues through the stages examined in this study.
2.3. zomp-1 and zotolin-1 knockdown phenotypes
To probe the function of zOMP-1 and zOtolin-1, we used
morpholino antisense oligonucleotides (MO) to block
translation of their mRNA in zebrafish embryos. Injection
of zomp-1 MO or zotolin-1 MO into 1–2 cell embryos
reproducibly induced phenotypes of abnormal otolith
morphology. Otolith seeding appeared normal in both
zomp-1 and zotolin-1 MO-injected embryos (Fig. 3A–C).
The kinocilia of the tether cells were formed normally in all
morphants and the nascent otoliths formed at their tips (Fig.
3D–F), indicating that the otolith seeding phase was not
affected by the absence of zOMP-1 or zOtolin-1. This is
consistent with our previous study on the rainbow trout,
which revealed that OMP-1 and Otolin-1 were not present in
the core of the embryonic otoliths (Murayama et al., 2004).
zomp-1 MO-injected embryos displayed a phenotype
marked mainly by the smaller size of both otoliths
(‘small’ phenotype). The difference with control embryos
increased with time, indicating a reduced otolith growth rate
(Fig. 3I,M,Q). This phenotype was observed in both otic
vesicles of each embryo (Table 1).
We examined this otolith growth defect more closely by
measuring the longest linear dimension of the otoliths
(otolith length) in the live embryos at several stages of their
development (Fig. 4 and supplemental Table S1). In control
embryos, the anterior and posterior otoliths grew at the same
rate until 35 hpf. After this stage, the growth rate of the
anterior otolith became much slower than that of the
posterior otolith (Fig. 4A). In zomp-1 MO-injected embryos,
the growth of both otoliths proceeded slowly in contrast to
control embryos (Fig. 4B). At the 28-somite stage, the
otolith length was only 6% smaller than in controls for
the anterior (P!0.05) and 14% smaller for the posterior
(P!0.001) otolith. This relative difference increased to
26% (P!0.001) for the anterior, and 44% (P!0.001) for
the posterior otolith at 72 hpf (Table S1). In terms of otolith
volume, this translates into a 4-fold lower growth rate for
the anterior, and a 20-fold lower growth rate for the
posterior otolith between 35 and 72 hpf.
Injection of zotolin-1 MO mainly resulted in a fusion of
the two otoliths. Typically, at 4 ng of injected MO, the two
otoliths became closer to each other by 35 hpf (Fig. 3J),
came into contact by 55 hpf (‘contact’ phenotype, Fig. 3N),
and then were fused by 72 hpf (‘fused’ phenotype, Fig. 3R).
On an average, this progression towards fusion occurred
faster upon increasing the MO dose (between 1 and 8 ng).
Thus, at 72 hpf, 53.5% of embryos injected with 4 ng MO
displayed fused otoliths and 21.7% showed coalescent
otoliths, while only 28.7% of those injected with 1 ng MO
were already fused, and 33.9% still contact (Table 1).
Whatever the MO dose and developmental time, the stage in
this phenotypic progression was always similar in both otic
vesicles of each embryo, strengthening the conclusion that
variations in the timing of this phenotypic progression
reflected variations in the actual MO dose present in the
embryo’s cells. In zotolin-1 MO-injected embryos, the
posterior otolith was also significantly smaller than in the
controls, mainly due to a slower growth rate between 35 and
55 hpf (Fig. 4A,C and Table S1).
In both zomp-1 and zotolin-1 MO-injected embryos,
beside the otolith phenotypes, the development of support-
ing cells and hair cells appeared normal through the stages
examined, both in the maculae (Fig. 3G,K) and cristae (data
not shown). In contrast, the otic vesicle of both zomp-1 MO
and zotolin-1 MO-injected embryos appeared slightly
smaller, and rounder with time, than that of control
embryos, and the semicircular canals did not form; an
incomplete protrusion of anterior and posterior canals was
present by 72 hpf, while the lateral protrusion did not occur,
even past 5 dpf (data not shown). No abnormalities were
found in zomp-1 MO and zotolin-1 MO-injected embryos
outside the otocysts.
2.4. Immunostaining of the otoliths in control and MO-
injected embryos
To examine the localization of zOMP-1 and zOtolin-1 in
the otoliths of control embryos, and the extent of
suppression of zomp-1 and zotolin-1 expression by the
MOs, we stained the injected embryos with antibodies
previously raised against recombinant rtOMP-1 and csOto-
lin-1 (Murayama et al., 2004). In wild type embryos, the
anti-OMP-1 antibody first labeled both otoliths at the 26-
somite stage (22 hpf, Fig. 2E), and then more strongly at
27 hpf (Fig. 5B) and 35 hpf (Fig. 2F). It is notable that the
Fig. 3. Otolith phenotypes observed in live control, zomp-1 and zotolin-1 MO-injected embryos by DIC videomicroscopy. Embryos were injected with 4 ng
MO. (A–C) 28-somite stage; otolith seeding occurs normally in control MO (A), zomp-1 MO (B) and zotolin-1 MO (C) injected embryos. (D–F) Magnification
of the anterior otolith and two tethering kinocilia shown in (A–C). (G–K) At 35 hpf, the otoliths of zomp-1 MO (I) and zotolin-1 MO (J) injected embryos are
slightly smaller, and closer to each other, respectively, than those of control MO-injected embryos (H); the anterior otolith of zotolin-1 MO-injected embryos
does not stay in contact with the anterior macula (arrowhead in J); (G, K) Anterior macula, otolith and tethering kinocilia of the zomp-1 MO ear shown in (I) and
zotolin-1 MO ear shown in (J). (L–N) At 55 hpf, zomp-1 MO and zotolin-1 MO injection cause a ‘small otoliths’ phenotype (M) and a ‘fused otoliths’
phenotype (N), respectively. (O) Magnification of the cilia of the anterior macula shown in (N). At 72 hpf (P–S), the otoliths of control MO (P) and zotolin-1
MO (R) injected embryos keep growing, especially the posterior otolith, while both otoliths of zomp-1 MO-injected embryos stay small (Q). (S) Magnification
of the cilia of the anterior macula shown in (R). In panels (A–C), (H), (I), (L), (M), (P) and (Q), the pictures of posterior otoliths are inlayed, from a deeper focal
plane, onto the main pictures, which are focused on the anterior otoliths. Through the panels, anterior is to the left, dorsal to the top. Scale bar in (A) indicates
25 mm in (A–C), in (D) indicates 10 mm in (D–F), in (H) indicates 25 mm in (H–J, L–N, P–R), in (G) indicates 10 mm in (G), (K), (O) and (S).
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803796
size of the otoliths observed at these stages by immunos-
taining of fixed embryos was larger than observed in live
embryos (Fig. 5A,B). At this stage, Otolin-1 was not
detected in the otoliths yet (Fig. 5C). At 72 hpf, the otoliths
of control MO-injected embryos (Fig. 5D) were intensely
immunostained with the anti-OMP-1 antibody (Fig. 5E).
They were also stained by the anti-Otolin-1 antibody,
although less strongly (Fig. 5F). In zomp-1 MO-injected
embryos (Fig. 5G), both otoliths were immunonegative with
the anti-OMP-1 antibody (Fig. 5H), demonstrating that the
zomp-1 MO effectively suppressed OMP-1 expression.
Similarly, the otoliths of zotolin-1 MO-injected embryos
Table 1
Dose dependence of otolith phenotypes at 72 hpf
Dose of
MO
injected
Otolith phenotypes of normal embryos (%) Curved
body axis
(%)Normal Fusion Contact Small
Control MO
1 ng 96.8G0.6 0 0 1.9G0.8 1.4G0.2
4 ng 95.9G0.1 0 0 2.1G0.3 1.7G0.4
zomp-1 MO
1 ng 41.2G5.6 0 0 56.8G5.4 2.1G0.2
4 ng 20.9G1.1 0 7.2G3.3 68.1G3.7 3.9G1.2
8 ng 10.2G2.3 0 5.9G1.8 73.9G1.2 10.1G1.7
zotolin-1 MO
1 ng 24.6G8.8 28.7G2.5 33.9G3.2 5.8G2.1 7.2G1.1
4 ng 7.2G0.8 53.5G2.7 21.7G2.6 2.2G0.4 15.6G1.1
8 ng 3.2G0.8 64.1G6 14.8G0.6 3.15G0.8 16.5G4.2
The numbers shown are the averages of two independent experiments. In
each experiment, at least 50 embryos were phenotyped for each dose of
MO. Otolith phenotypes were always similar in both ears. About 5% of the
embryos had an additional otolith in one ear, irrespective of MO nature and
dose.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 797
were immunonegative with the anti-Otolin-1 antibody
(Fig. 5K). Surprisingly, the anti-Otolin-1 antibody also
failed to stain the otoliths of zomp-1 MO-injected embryos
(Fig. 5I). This result suggests that the presence of zOMP-1
in the growing otolith is a prerequisite for the deposition of
zOtolin-1. Our previous work showed that Otolin-1 and
OMP-1 were co-localized in the otoliths of adult rainbow
trout (Murayama et al., 2004), and that purified rainbow
trout OMP-1 and Otolin-1 proteins interacted with each
other in vitro (Murayama, 2002b). These earlier obser-
vations support the present results. In contrast, the fused
otoliths of zotolin-1 MO-injected embryos (Fig. 5J) were
strongly immunolabeled by the anti-OMP-1 antibody at
72 hpf (Fig. 5L). Intriguingly, the immunostained otoliths
appeared substantially larger than in the live embryos
(Fig. 5J,L), just as we had found before with control
embryos of 22–35 hpf (Figs. 2E,F,5A,B). In other words,
the otolith matrix appeared to swell during the immunos-
taining procedure only when zOtolin-1 was absent, either
because not present yet in the otoliths (22–35 hpf embryos)
or because suppressed by the zotolin-1 MO (such swelling
did not occur, e.g. in 72 hpf control embryos, Fig. 5D–F).
Fig. 4. Otolith growth rates in control, zomp-1 and zotolin-1 MO-injected embryos
of anterior (blue squares) or posterior (orange circles) otoliths measured at various
each viewed laterally. Error bars show SD. The numbers are given in Table S1.
Correlatively, the zOMP-1 immunostaining was always
more intense in these swelled otoliths (Figs. 2F,5B,L),
which may be due to increased accessibility of zOMP-1
epitopes within the swollen organic matrix. This obser-
vation suggests that the presence of zOtolin-1 may stabilize
the otolith organic matrix in a compact form. Another
observation suggests that it may also stabilize its association
with the mineral component. Upon fixation, otoliths are
known to decalcify spontaneously with time. We observed
that this process occurs clearly faster in otoliths of zotolin-1
MO-injected embryos than in control embryos (all fixed at
7 dpf; see supplemental Fig. S1). These various obser-
vations suggest that the incorporation of the collagenous
protein zOtolin-1 in the otoliths may stabilize both the
mineral and the organic matrix components of the otoliths.
In trout otoliths, we previously found Otolin-1 and OMP-
1 co-localized in the daily organic increment layers
(Murayama et al., 2004). In zebrafish embryos, the adult
mode of otolith growth by alternate daily increments
appears to start by 3 dpf (J.Y. Sire, personal communi-
cation), i.e. about when we start to detect Otolin-1 clearly in
the otoliths. So we suggest that the switch to this growth
mode requires the participation of zOtolin-1, possibly
forming a 3D-lattice similar to what the related proteins
collagen VIII and X do, and that the resulting layered
structure brings increased stability to the otolith.
2.5. Relationship of the fused otoliths to their respective
sensory maculae in zotolin-1 MO-injected embryos
The ‘fused otoliths’ phenotype observed in vivo with the
zotolin-1 MO suggests that the otolith matrix has self-
aggregation property, causing the fusion of otoliths when
they are in contact with each other. This recalls a common
observation done on wild type embryos. A fraction of
them (about 5% in the stocks used for the present study—see
Table 1—but it can be quite higher in occasional stocks)
initially make one additional otolith; in most cases, this
additional otolith fuses sooner or later with the anterior or the
posterior otolith (the closer one; when this additional otolith
is unanchored, it most often fuses with the posterior otolith).
More intriguing here is the progressive displacement of
. Each point represents the mean of longest linear dimension (otolith length)
stages of development. Size measurements were made from 15 specimens
Fig. 5. Whole-mount immunostaining of the otoliths of wild type and MO-injected embryos. (A, D, G, J) Live embryos; all other panels show fixed,
immunostained embryos. (A–C) Wild-type embryos at 27 hpf; (B) zOMP-1 is detected in both otoliths and in the ventro-lateral epithelium posterior to the
anterior macula (arrowheads), but zOtolin-1 is not observed yet (C). (D–O) Embryos at 72 hpf. In control MO-injected embryos, anti-OMP-1 (E) and anti-
Otolin-1 (F) antibodies recognize both otoliths. (G–I) zomp-1 MO-injected embryos; zOMP-1 is effectively knocked down (H); the anti-Otolin-1 antibody fails
to detect zOtolin-1 in the small otoliths (I). (J–L) zotolin-1 MO-injected embryos; zOtolin-1 is not detected (K) while the anti-OMP-1 antibody reacts strongly
with the fused otoliths (L). (M–O) Pre-immune serum shows non-specific staining (arrowheads) of the sensory hair cells of the anterior macula (M), posterior
macula (N) and anterior and lateral cristae (O). In panels (A–G), the posterior otoliths were inlayed from a deeper focal plane onto the main pictures, which are
focused on the anterior otoliths. Dotted lines outline otoliths in panels (H), (I) and (K) and the posterior macula in (N). Through the panels, anterior is to the left,
dorsal to the top. Scale bar in (A) and (O) indicates 25 mm in (A–N) and (O), respectively.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803798
the anterior and posterior otoliths of zotolin-1 MO-injected
embryos that bring them into contact with each other.
Initially, they nucleate normally on the tips of the kinocilia of
the tether cells (Fig. 3F), which lie in the middle of the
prospective maculae. However, by 35 hpf, the anterior
otolith has already moved somewhat away from the anterior
macula (the position of the posterior otolith relative to its
macula is less easy to evaluate in vivo from this stage, due to
the more medial position of the latter) and by 72 hpf this
distance has increased (Fig. 3R). Yet it seems that the otolith
is still connected to the corresponding macula, the kinocilia
are longer than normal (Fig. 3K,O,S). To clarify the
topographical relationship of the fused otoliths to their
respective maculae and associated stereo- and kino-cilia in
zotolin-1 MO embryos, we fixed them at 72 hpf and
performed triple immunofluorescence confocal analysis
through the depth of the otocyst, labeling the otolith with
OMP-1 antibody, the kinocilia with anti-acetylated tubulin
Fig. 6. Relationship of the fused otoliths in zotolin-1 MO-injected embryos to the corresponding sensory maculae. Triple-labeling with anti-OMP-1 (red), anti-
acetylated tubulin (blue) antibodies, and phalloidin (green). Confocal fluorescence images at 72 hpf. Phalloidin (green) reveals the hair (stereocilia) bundles of
the sensory cells and the contours of cells through their actin cortex. Acetylated tubulin (blue, false color) is found in the kinocilia and in the apical cytoplasm of
the sensory hair cells (Riley et al., 1997). (A) Anterior macula and otolith of a control MO-injected embryo, lateral view. (B–D) zotolin-1 MO-injected embryo,
with fused otoliths. (B) and (C) are the same z-stacks (3D reconstruction available upon request). (B) Anterior macula and anterior otolith with the fused
posterior otolith; note the long kinocilia (arrowhead). (C) Projection of the confocal series along the z-axis (22 images, taken every 2 mm). Both maculae are
visible through their hair bundles, in relation to both otoliths. (D) Far-red (blue-coded in A–C) fluorescence channel extracted from the image shown in (C), to
show the acetylated tubulin immunostaining of kinocilia at both maculae. ao, anterior otolith; po, posterior otolith. Dorso-lateral view, anterior to the left,
dorsal to the top. Scale bars, 20 mm.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 799
antibody, and the stereocilia with phalloidin (Fig. 6). This
analysis confirmed that both maculae display well-organized
stereocilia and kinocilia. It also showed that each of the two
fused otoliths remains associated with its respective macula
(Fig. 6B,C; 3D reconstruction available upon request). They
did not appear distant from the maculae as in live embryos,
possibly due to the swelling of the otolith matrix discussed
above. However, the kinocilia between the macula and the
otolith seemed particularly long (Fig. 6A,B,D), as the in vivo
observation had suggested.
In adult fish, the otoliths are bound to the sensory macula
via a gelatinous material called otolithic membrane
(Dunkelberger et al., 1980). Otolin-1 was also identified
as a component of the otolithic membrane in adult rainbow
trout (Murayama et al., 2002, 2004) and bluegill sunfish
(Davis et al., 1997), suggesting that Otolin-1 may be
involved in the positioning of the otoliths via the otolithic
membrane. It is tempting to link these data from adult fish
with the present observation that the otoliths of zotolin-1
MO-injected embryos do not stay in contact with the
corresponding maculae. The development of the otolithic
membrane has not been documented in the zebrafish (nor in
other fish to our knowledge). The fact that the otoliths are
already moving away from the maculae by 35 hpf in the
zotolin-1 MO-injected embryos would suggest that some
form of Otolin-1-containing otolithic membrane is already
present by that stage in control embryos, and that it
contributes to maintaining the otoliths close to their
respective maculae. Our failure to immunodetect such an
Otolin-1-containing structure would not be surprising given
the moderate affinity of our anti-chum salmon Otolin-1
antibody for the zebrafish protein.
2.6. Behavioral defects in the zomp-1 and zotolin-1
MO-injected embryos
Finally, we examined the behavioral changes caused by
the loss of zOMP-1 or zOtolin-1. Vestibular defects became
Table 2
Behavioral defects at 4 dpf embryos injected with 4 ng MOs
MO Control MO zomp-1 MO zotolin-1 MO
Phenotype Normal otoliths Small otoliths Fused otoliths
No. of tested
animals
38 71 67
Swimming 21 (55.3%) 0 0
Laying down 17 (44.7%) 71 (100%) 67 (100%)
No reaction 0 22 (31.0%) 18 (26.8%)
Short
movements
17 (44.7%)a 22 (31.0%)b 20 (29.9%)b
Circular
movements
0 27 (38.0%) 29 (43.3%)
Stimulus: tapping the edge of the Petri dish with tweezers.a Linear movements.b Non-linear movements.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803800
obvious after 4 dpf. At this stage, nearly half of the non-
injected wild type embryos or control MO-injected embryos
keep an upright position and swim spontaneously (Table 2).
In contrast, all zomp-1 and zotolin-1 MO-injected embryos
lay down on the bottom of the Petri dish and remained still.
When the edge of the Petri dish was tapped, the half of
control MO-injected embryos that were laying down were
very sensitive to the stimulus and started swimming in a
straight manner (Movie S1). In contrast, about 30% of
zomp-1 MO or zotolin-1 MO-injected embryos showed only
short movements in response to the stimulus, and never kept
an upright position (Movie S2). Another 40% of zomp-1 MO
or zotolin-1 MO-injected embryos exhibited circular move-
ments (Movie S3). The remaining 30% of zomp-1 MO or
zotolin-1 MO-injected embryos did not respond to the
tapping stimulus (Table 2), whereas they did show an escape
response upon stimulation by touch with a tweezer tip. Most
otolith mutants (Whitfield et al., 1996) and some sensory
hair cell mutants (Nicolson et al., 1998) show similar
phenotypes, such as circular movement including looping
and/or rolling motion. The normal development of vestib-
ular function in zebrafish larvae is dependent on stimulus by
otolith weight (Moorman et al., 1999). Since the develop-
ment of the sensory maculae appears normal in zomp-1 MO
and zotolin-1 MO-injected embryos, the observed beha-
vioral defects are most likely caused by the lack or reduction
of hair cell stimulation due to the malformation or
malpositioning of the otoliths.
This work addressed the molecular mechanisms of
otolith formation through the roles of two otolith matrix
proteins, zOMP-1 and zOtolin-1. zOMP-1 is especially
involved in otolith growth and appears to be required for the
deposition of zOtolin-1. In contrast, zOtolin-1 is required
for the correct anchoring of the otoliths on the sensory
maculae, and as a component of the otolith it seems to
stabilize the otolith matrix from about 72 hpf, presumably
by providing a collagenous scaffold. Thus, so far, three
proteins, OMP-1 (Murayama et al., 2000), Otolin-1
(Murayama et al., 2002) and Starmaker (Sollner et al.,
2003), are known to be components of fish otoliths. Future
analyses of the interaction of these proteins with each other
and with calcium carbonate should contribute further
insights into the process of biomineralization. In addition,
comprehensive studies of other components contained in the
otolith, such as minor components of the protein matrix,
proteoglycans (Borelli et al., 2003) and polysaccharides
(Pisam et al., 2002), as well as of the otolithic membrane
and endolymph, will be needed for a precise understanding
of the molecular mechanism of otolith formation.
3. Experimental procedures
3.1. Fish strains and maintenance
AB and Tu wild type strains of the zebrafish, Danio rerio,
were used throughout this study. Fishes were kept under a
photoperiod of 14 h light/10 h dark at 28.5 8C. Eggs were
obtained by random crosses and kept at the same
temperature. Embryos were staged as hours post-fertiliza-
tion (hpf) at 28.5 8C and using developmental landmarks
(Kimmel et al., 1995).
3.2. Cloning of zebrafish omp-1 and otolin-1 cDNAs
A cDNA was synthesized with a SMART RACE cDNA
Amplification kit (Clontech) according to the manufac-
turer’s instruction using 1 mg of total RNA isolated from the
inner ear of adult zebrafish. To amplify the cDNA fragments
encoding zebrafish omp-1 (zomp-1) and otolin-1 (zotolin-1),
the following sets of degenerate oligonucleotide primers
were used.
zomp-1-F (5 0-GARGCIGARGARCARAARTG-3 0)
zomp-1-R (5 0-NWSICKCTCRTGCATCTC-3 0)
zotolin-1-F (5 0-TAYAAYGGCGARGGICAYTG
GGA-3 0)
zotolin-1-R (5 0-GCYTGRTCDATRTCYTGICC-3 0)
These primers were designed based on the deduced
amino acid sequence of rainbow trout OMP-1 (Murayama
et al., 2000) and chum salmon Otolin-1 (Murayama et al.,
2002), respectively. PCR reactions were carried out as
described previously (Murayama et al., 2002) using
zebrafish inner ear cDNA as a template. Briefly, the reaction
mixtures were denatured at 94 8C for 3 min followed by 40
cycles of 94 8C for 30 s, 54 8C for 30 s, and 72 8C for 30 s.
To isolate the 5 0-end regions, a 5 0 RACE reaction was
performed using gene specific primers, zomp-1-5R1 (5 0-
GTTGTCCGCGAAGGTTCCTGGCTGTGGC-3 0) and
zotolin-1-5R1 (5 0-AAGAGAGTCACGCGTTCGCAG-
TTTCCG-3 0). For zotolin-1, the PCR reaction was carried
out using NUP primer (Clontech) and zoto1-5R2 (5 0-
ACGTACAGTTATGTAGTATGAAAAGACG-3 0) as
described previously (Murayama et al., 2002). To obtain
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 801
the complete 5 0 end of zotolin-1 cDNA, an additional RACE
reaction was done using the following primers:
zotolin-1-5R3 (5 0-GTTCCATTCCAGCCAGGCT-
CTCCGCGC-3 0)
zotolin-1-5R4 (5 0-CGCGCTCCCCTGGGTCTC-
CTTTCAGCC-3 0)
The amplified cDNA fragments were subcloned into a
pCR2.1 TOPO vector (Invitrogen) according to the
manufacturer’s instructions. The nucleotide sequences
were determined using a Thermo Sequenase Cy5 dye
terminator cycle sequencing kit (Amarsham Biosciences).
The nucleotide sequences of zomp-1 and zotolin-1 were
submitted to DDBJ/EMBL/GenBank and have been
assigned the accession numbers AB124553 and
AB124554, respectively.
3.3. Whole-mount in situ hybridization
Whole-mount in situ hybridization was performed
according to the procedure described by Thisse et al.
(1993) with slight modifications, using the digoxygenin-
UTP labeled RNA probes of zomp-1 and zotolin-1. The
cDNA fragments of zomp-1 (nucleotides 137–436) and
zotolin-1 (nucleotides 1227–1408) were subcloned into a
pCR4-TOPO vector (Invitrogen). Both constructs were
digested with Not I and transcribed with T3 RNA
polymerase. For embryos older than 48 hpf, heads were
cut off before hybridization. Hybridization was performed
overnight at 70 8C with 150 ng DIG-labeled riboprobe in
200 ml hybridization buffer. After a series of washes with
PBT (0.1% Tween-20 in PBS), embryos were incubated
with preabsorbed anti-DIG-alkaline phosphatase Fab frag-
ments (1:4000) overnight at 4 8C.
3.4. Whole-mount immunostaining and phalloidin staining
Embryos were fixed with 4% paraformaldehyde/PBS
overnight at 4 8C. After washing with PBST (0.1% TritonX-
100 in PBS), the samples were dehydrated overnight with
methanol and stored at K20 8C. The samples were
rehydrated with PBST, the heads were cut off, then treated
with the blocking solution (5% sheep serum, 0.2% BSA in
PBST) for 2 h at room temperature. Primary and secondary
antibodies were diluted in blocking solution. Primary
antibodies were (1) anti-rtOMP-1 polyclonal antibody
(Murayama et al., 2004) at 1:1000 dilution, (2) anti-
csOtolin-1 polyclonal antibody (Murayama et al., 2004) at
1:500 dilution, or (3) anti-acetylated tubulin monoclonal
antibody (Sigma) at 1:500 dilution. The embryos were
incubated with these antibodies overnight at 4 8C. After
washing with PBST for at least 4 h, the embryos were
blocked again then incubated with the corresponding
secondary antibodies; (1 0) biotin-conjugated anti-rabbit
IgG antibody or (2 0) horseradish peroxydase linked
anti-rabbit IgG at 1:800 dilution (Amersham Biosciences).
The signals were detected with the ABC staining kit
(Vector) using 3,3 0-diaminobenzidine (Sigma) as a sub-
strate, or directly visualized with 3-amino-9-ethyl calbazole
(Sigma) as a substrate. For the confocal fluorescence
analysis, the following secondary antibodies were used:
(1 00) Cy3-conjugated anti-rabbit IgG Fab fragments at 1:800
dilution (Jackson laboratory) and (3 0) Cy5-conjugated anti-
mouse IgG Fab fragments at 1:800 dilution (Jackson
laboratory), mixed with Alexa Fluor 488-phalloidin
(1:200, Molecular Probes). They were applied overnight
in the dark at 4 8C. The embryos were thoroughly rinsed,
mounted in glycerol, and viewed under the 40! oil
objective of a Zeiss LSM 510 confocal fluorescence
microscope (Zeiss Axioskop 2FSM). Optical sections
were taken every 2 mm.
3.5. Microinjection of morpholino oligonucleotides
Morpholino oligonucleotides (MOs) were designed to
target the initiation codons of zomp-1 and zotolin-1 mRNAs.
zomp-1 MO (5 0-CAAGATGTCCTCCTGGAAGATC-
CAT-3 0)
zotolin-1 MO (5 0-TGAACGGGTGGAGAATATTGGG-
CAT-3 0)
GeneTools control MO (5 0-CCTCTTACCTCAGTTA-
CAATTATA-3 0)
These MOs were dissolved in distilled water at 10 mg/ml
and diluted to 8 or 1 mg/ml with 200 mM KCl, containing
phenol red (2.5 mg/ml) before the injection. MOs (0.5–1 nl)
were injected into the yolk of 1–2 cell stage embryos
together with rhodamine-dextran 10000 conjugate (Mol-
ecular Probes) as a fluorescent tracer to check for the
homogenous distribution of the injected material in the
resulting embryos. After injection, embryos were incubated
in Volvic water at 28.5 8C until desired developmental
stages. Embryos with non-homogenous rhodamine-dextran
distribution at 22 hpf were discarded.
3.6. Imaging and otolith measurements on live embryos
Differential Interference Contrast (DIC) video-
microscopy was performed on live embryos as described
previously (Herbomel et al., 1999). To determine the size of
otoliths in MO-injected embryos at successive develop-
mental stages, the embryos were anesthetized and then
observed and video-recorded at 28-somite stage (29 hpf),
35, 55 and 72 hpf, by DIC video-microscopy (stage 28-
somite), or under a Leica MZ16 stereomicroscope at
maximum magnification (all other stages). Otolith measure-
ments were made in a total of 15 pairs of inner ears for both
anterior and posterior otoliths. Differences between zomp-1
(or zotolin-1 MO) injected embryos and control embryos
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803802
were tested using the Student’s t test at each developmental
stage.
3.7. Behavioral tests
The MO-injected embryos at 4 dpf were placed in a 96-
well plate (one embryo per well) and left without
stimulation for at least 30 s. After verifying their inert
status, the edge of the well was tapped with a forceps. Their
reaction was observed and video-recorded for 10 s follow-
ing stimulation.
Acknowledgements
We are grateful to Kazuki Horikawa for many valuable
discussions, and to Karima Kissa-Marin and Pascal Roux
for their help with the confocal microscopy. We also thank
the members of Unite Macrophages et Developpement de
l’Immunite for their kind support. This work was supported
by Grants-in-Aid for Creative Basic Research (#12NP0201)
and for Scientific Research (Nos. 12876025, 13660176 and
13660178) from the Ministry of Education, Culture, Sports,
Science and Technology of Japan. E. M. was supported by a
Research Fellowship of the Japan Society for the Promotion
of Science for Young Scientists.
Supplementary material
Supplementary data associated with this article can
be found, in the online version, at doi:10.1016/j.mod.2005.
03.002
References
Aisen, P., Leibman, A., 1972. Lactoferrin and transferrin: a comparative
study. Biochim. Biophys. Acta 257, 314–323.
Apte, S., Mattei, M.G., Olsen, B.R., 1991. Cloning of human alpha 1(X)
collagen DNA and localization of the COL10A1 gene to the q21–q22
region of human chromosome 6. Fed. Eur. Biochem. Soc. Lett. 282,
393–396.
Baker, E.N., Baker, H.M., Smith, C.A., Stebbins, M.R., Kahn, M.,
Hellstrom, K.E., Hellstrom, I., 1992. Human melanotransferrin (p97)
has only one functional iron-binding site. Fed. Eur. Biochem. Soc. Lett.
298, 215–218.
Balsamo, G., Avallone, B., Del Genio, F., Trapani, S., Marmo, F., 2000.
Calcification processes in the chick otoconia and calcium binding
proteins: patterns of tetracycline incorporation and calbindin-D28K
distribution. Hear. Res. 148, 1–8.
Bever, M.M., Fekete, D.M., 2002. Atlas of the developing inner ear in
zebrafish. Dev. Dyn. 223, 536–543.
Bogin, O., Kvansakul, M., Rom, E., Singer, J., Yayon, A., Hohenester, E.,
2002. Insight into Schmid metaphyseal chondrodysplasia from the
crystal structure of the collagen X NC1 domain trimer. Structure
(Cambridge) 10, 165–173.
Borelli, G., Mayer-Gostan, N., Merle, P.L., De Pontual, H., Boeuf, G.,
Allemand, D., Payan, P., 2003. Composition of biomineral organic
matrices with special emphasis on turbot (Psetta maxima) otolith and
endolymph. Calcif. Tissue Int. 72, 717–725.
Burge, C., Karlin, S., 1997. Prediction of complete gene structures in
human genomic DNA. J. Mol. Biol. 268, 78–94.
Campana, S.E., Neilson, J.D., 1985. Microstructure of fish otoliths. Can.
J. Fish Aquat. Sci. 42, 1014–1032.
Chan, D., Jacenko, O., 1998. Phenotypic and biochemical consequences of
collagen X mutations in mice and humans. Matrix Biol. 17, 169–184.
Chan, D., Cole, W.G., Rogers, J.G., Bateman, J.F., 1995. Type X collagen
multimer assembly in vitro is prevented by a Gly618 to Val mutation in
the alpha 1(X) NC1 domain resulting in Schmid metaphyseal
chondrodysplasia. J. Biol. Chem. 270, 4558–4562.
Davis, J.G., Oberholtzer, J.C., Burns, F.R., Greene, M.I., 1995. Molecular
cloning and characterization of an inner ear-specific structural protein.
Science 267, 1031–1034.
Davis, J.G., Burns, F.R., Navaratnam, D., Lee, A.M., Ichimiya, S.,
Oberholtzer, J.C., Greene, M.I., 1997. Identification of a structural
constituent and one possible site of postembryonic formation of a
teleost otolithic membrane. Proc. Natl Acad. Sci. USA 94, 707–712.
Dunkelberger, D.G., Dean, J.M., Watabe, N., 1980. The ultrastructure of
the otolithic membrane and otolith in the juvenile mummichog,
Fundulus heteroclitus. J. Morphol. 163, 367–377.
Grant, W.T., Wang, G.J., Balian, G., 1987. Type X collagen synthesis
during endochondral ossification in fracture repair. J. Biol. Chem. 262,
9844–9849.
Haddon, C., Lewis, J., 1996. Early ear development in the embryo of the
zebrafish, Danio rerio. J. Comp. Neurol. 365, 113–128.
Herbomel, P., Thisse, B., Thisse, C., 1999. Ontogeny and behaviour of early
macrophages in the zebrafish embryo. Development 126, 3735–3745.
Jongeneel, C.V., Bouvier, J., Bairoch, A., 1989. A unique signature
identifies a family of zinc-dependent metallopeptidases. Fed. Eur.
Biochem. Soc. Lett. 242, 211–214.
Kapoor, R., Sakai, L.Y., Funk, S., Roux, E., Bornstein, P., Sage, E.H., 1988.
Type VIII collagen has a restricted distribution in specialized
extracellular matrices. J. Cell Biol. 107, 721–730.
Kawamoto, T., Pan, H., Yan, W., Ishida, H., Usui, E., Oda, R., et al., 1998.
Expression of membrane-bound transferrin-like protein p97 on the cell
surface of chondrocytes. Eur. J. Biochem. 256, 503–509.
Kimmel, C.B., Ballard, W.W., Kimmel, S.R., Ullmann, B., Schilling, T.F.,
1995. Stages of embryonic development of the zebrafish. Dev. Dyn.
203, 253–310.
Kirsch, T., von der Mark, K., 1991. Ca2C binding properties of type X
collagen. Fed. Eur. Biochem. Soc. Lett. 294, 149–152.
Kwan, A.P.L., Cummings, C.E., Chapman, J.A., Grant, M.E., 1991.
Mascromolecular organization of chicken type X collagen in vitro.
J. Cell Biol. 114, 597–604.
Labermeier, U., Demlow, T.A., Kenney, M.C., 1983. Identification of
collagens isolated from bovine Descemet’s membrane. Exp. Eye Res.
37, 225–237.
Lowenstein, O., 1971. The labyrinth. In: Hoar, W.S., Randall, D.J. (Eds.),
Fish Physiology, Sensory System and Electric Organs, vol. V.
Academic Press, New York, pp. 207–240.
Manning, F.B., 1924. Hearing in the goldfish in relation to the structure of
its ear. J. Exp. Zool. 41, 5–20.
Moorman, S.J., Burress, C., Cordova, R., Slater, J., 1999. Stimulus
dependence of the development of the zebrafish (Danio rerio)
vestibular system. J. Neurobiol. 38, 247–258.
Mugiya, Y., 1987. Phase difference between calcification and organic
matrix formation in the diurnal growth of otoliths in the rainbow trout,
Salmo gairdneri. Fish. Bull. 85, 395–401.
Muragaki, Y., Mattei, M.G., Yamaguchi, N., Olsen, B.R., Ninomiya, Y.,
1991. The complete primary structure of the human alpha 1 (VIII) chain
and assignment of its gene (COL8A1) to chromosome 3. Eur.
J. Biochem. 197, 615–622.
Murayama, E., 2002. Studies on molecular mechanism of otolith
formation in salmonids. PhD dissertation, the University of Tokyo,
Tokyo, Japan.
E. Murayama et al. / Mechanisms of Development 122 (2005) 791–803 803
Murayama, E., Okuno, A., Ohira, T., Takagi, Y., Nagasawa, H., 2000.
Molecular cloning and expression of an otolith matrix protein cDNA
from the rainbow trout, Oncorhynchus mykiss. Comp. Biochem.
Physiol. 126B, 511–520.
Murayama, E., Takagi, Y., Ohira, T., Davis, J.G., Greene, M.I.,
Nagasawa, H., 2002. Fish otolith contains a unique structural protein,
otolin-1. Eur. J. Biochem. 269, 688–696.
Murayama, E., Takagi, Y., Nagasawa, H., 2004. Immunohistochemical
localization of two otolith matrix proteins in the otolith and inner ear of
the rainbow trout, Oncorhynchus mykiss: comparative aspects between
the adult inner ear and embryonic otocysts. Histochem. Cell Biol. 121,
155–166.
Nakamasu, K., Kawamoto, T., Shen, M., Gotoh, O., Teramoto, M.,
Noshiro, M., Kato, Y., 1999. Membrane-bound transferrin-like protein
(MTf): structure, evolution and selective expression during chondro-
genic differentiation of mouse embryonic cells. Biochim. Biophys. Acta
1447, 258–264.
Nicolson, T., Rusch, A., Friedrich, R.W., Granato, M., Ruppersberg, J.P.,
Nusslein-Volhard, C., 1998. Genetic analysis of vertebrate sensory hair
cell mechanosensation: the zebrafish circler mutants. Neuron 20, 271–
283.
Pisam, M., Jammet, C., Laurent, D., 2002. First steps of otolith formation of
the zebrafish: role of glycogen? Cell Tissue Res. 310, 163–168.
Piscopo, M., Balsamo, G., Mutone, R., Avallone, B., Marmo, F., 2003.
Calbindin D28K is a component of the organic matrix of lizard
Podarcis sicula otoconia. Hear. Res. 178, 89–94.
Pote, K.G., Hauer 3rd., C.R., Michel, H., Shabanowitz, J., Hunt, D.F.,
Kretsinger, R.H., 1993. Otoconin-22, the major protein of aragonitic
frog otoconia, is a homolog of phospholipase A2. Biochemistry 32,
5017–5024.
Reid, K.B., 1985. Molecular cloning and characterization of the
complementary DNA and gene coding for the B-chain of subcomponent
C1q of the human complement system. Biochem. J. 231, 729–735.
Riley, B.B., Moorman, S.J., 2000. Development of utricular otoliths, but
not saccular otoliths, is necessary for vestibular function and survival in
zebrafish. J. Neurobiol. 43, 329–337.
Riley, B.B., Zhu, C., Janetopoulos, C., Aufderheide, K.J., 1997. A critical
period of ear development controlled by distinct populations of ciliated
cells in the zebrafish. Dev. Biol. 191, 191–201.
Sawada, H., Konomi, H., Hirosawa, K., 1990. Characterization of the
collagen in the hexagonal lattice of Descemet’s membrane: its relation
to type VIII collagen. J. Cell Biol. 110, 219–227.
Schmid, T.M., Conrad, H.E., 1982. Metabolism of low molecular
weight collagen by chondrocytes obtained from histologically distinct
zones of the chick embryo tibiotarsus. J. Biol. Chem. 257,
12451–12457.
Shuttleworth, C.A., 1997. Type VIII collagen. Int. J. Biochem. Cell Biol.
29, 1145–1148.
Sollner, C., Burghammer, M., Busch-Nentwich, E., Berger, J., Schwarz, H.,
Riekel, C., Nicolson, T., 2003. Control of crystal size and lattice
formation by starmaker in otolith biomineralization. Science 302, 282–
286.
Sumanas, S., Larson, J.D., Miller Bever, M., 2003. Zebrafish chaperone
protein GP96 is required for otolith formation during ear development.
Dev. Biol. 261, 443–455.
Sutmuller, M., Bruijn, J.A., de Heer, E., 1997. Collagen types VIII and X,
two non-fibrillar, short-chain collagens. Structure homologies, func-
tions and involvement in pathology. Histol. Histopathol. 12, 557–566.
Thisse, C., Thisse, B., Schilling, T.F., Postlethwait, J.H., 1993. Structure of
the zebrafish snail1 gene and its expression in wild type, spadetail and
no tail mutant embryos. Development 119, 1203–1215.
Thomas, J.T., Cresswell, C.J., Rash, B., Nicolai, H., Jones, T., Solomon, E.,
et al., 1991. The human collagen X gene. Complete primary translated
sequence and chromosomal localization. Biochem. J. 280, 617–623.
Verpy, E., Leibovici, M., Petit, C., 1999. Characterization of otoconin-95,
the major protein of murine otoconia, provides insights into the
formation of these inner ear biominerals. Proc. Natl Acad. Sci. USA 96,
529–534.
von der Mark, K., Kirsch, T., Nerlich, A., Kuss, A., Weseloh, G.,
Gluckert, K., Stoss, H., 1992. Type X collagen synthesis in human
osteoarthritic cartilage. Indication of chondrocyte hypertrophy. Arthri-
tis Rheum. 35, 806–811.
Wang, Y., Kowalski, P.E., Thalmann, I., Ornitz, D.M., Mager, D.L.,
Thalmann, R., 1998. Otoconin-90, the mammalian otoconial matrix
protein, contains two domains of homology to secretory phospholipase
A2. Proc. Natl Acad. Sci. USA 95, 15345–15350.
Watabe, N., Tanaka, K., Yamada, J., Dean, J.M., 1982. Scanning electron
microscope observations of the organic matrix in the otolith of the
teleost fish Fundulus heteroclitus and Tilapia nilotica. J. Exp. Mar.
Biol. Ecol. 58, 127–134.
Whitfield, T.T., Granato, M., van Eeden, F.J., Schach, U., Brand, M.,
Furutani-Seiki, M., et al., 1996. Mutations affecting development of the
zebrafish inner ear and lateral line. Development 123, 241–254.
Woodbury, R.G., Brown, J.P., Yeh, M.Y., Hellstrom, I., Hellstrom, K.E.,
1980. Identification of a cell surface protein, p97, in human melanomas
and certain other neoplasms. Proc. Natl Acad. Sci. USA 77, 2183–2187.
Yaoi, Y., Suzuki, M., Tomura, H., Sasayama, Y., Kikuyama, S., Tanaka, S.,
2003. Molecular cloning of otoconin-22 complementary deoxyribonu-
cleic acid in the bullfrog endolymphatic sac: effect of calcitonin on
otoconin-22 messenger ribonucleic acid levels. Endocrinology 144,
3287–3296.