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First published online 27 June 2007
doi: 10.1242/dev.004531
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1 Graduate School of Science, Nagoya University, Nagoya 464-8602, Japan.
2 Department of Molecular, Cellular and Developmental Biology, University of
Michigan, Ann Arbor, MI 48109-1048, USA.
3 Institute of Molecular Embryology and Genetics, Kumamoto University, Kumamoto
860-0811, Japan.
Author for correspondence (e-mail:
hhirata{at}bio.nagoya-u.ac.jp)
Accepted 29 May 2007
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SUMMARY |
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 TOP SUMMARY INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key words: Zebrafish, Ryanodine receptor, Muscle, Calcium, Multi-minicore disease
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INTRODUCTION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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).
Second, embryos are transparent, facilitating the visualization of dynamic
events, such as cell migration, axonal outgrowth and calcium transients, in
live fish (Fetcho and Bhatt,
2004; Wilson et al.,
2002). Third, electrophysiological techniques can be used to
analyze the physiology of embryonic neurons and muscles
(Drapeau et al., 2002;
Fetcho, 2006). Fourth,
zebrafish embryos exhibit readily assayable and well-characterized behaviors
(Eaton and Farley, 1973;
Saint-Amant and Drapeau,
1998). Fifth, zebrafish mutants can potentially serve as animal
models of human motor disorders (Bassett
and Currie, 2003; Kunkel et
al., 2006; Lieschke and
Currie, 2007).
Zebrafish embryos display three stereotyped behaviors by 36 hours
post-fertilization (hpf) (Saint-Amant and
Drapeau, 1998). The earliest behavior consists of repetitive, slow
and alternating coiling of the trunk and tail. This coiling is independent of
sensory stimulation and observed from 17 to 26 hpf. After 21 hpf, embryos
respond to mechanosensory stimulation with two or three rapid C-bends of the
trunk and tail. By 26 hpf, embryos swim in response to tactile stimulation.
The frequency of muscle contractions during swimming increases from 7 Hz at 26
hpf to 30 Hz at 36 hpf, the latter being similar to the frequency of swimming
by adult zebrafish (Buss and Drapeau,
2001).
The process of touch-induced swimming involves a number of steps, starting
with the sensing of tactile stimuli and ending with the contraction of
muscles. Touch is sensed by Rohon-Beard neurons in the trunk and tail or
trigeminal sensory neurons in the head
(Drapeau et al., 2002). Once
triggered by sensory inputs, interneuronal networks located in the hindbrain
and spinal cord create the appropriate motor pattern that alternately
activates motor neurons in each side of the spinal cord
(Fetcho, 1992;
Gahtan et al., 2002). Motor
terminals release acetylcholine at the neuromuscular junction (NMJ) to
depolarize the muscle membrane (Buss and
Drapeau, 2001; Wen and Brehm,
2005) and the change of membrane potential is converted to muscle
movement by excitation-contraction (E-C) coupling
(Franzini-Armstrong and Protasi,
1997). Depolarizations of the plasma membrane spread down the
transverse-tubules (t-tubules), which are invaginations of the plasma
membrane, and cause conformational changes of the dihydropyridine receptor
(DHPR), a voltage sensor located in the t-tubule membrane. DHPRs then trigger
the opening of ryanodine receptor 1 (RyR1) in the adjacent sarcoplasmic
reticulum (SR) to allow Ca2+ release from the SR to the cytosol
(Meissner, 1994). Elevated
cytoplasmic Ca2+, in turn, activates the sliding of actin/myosin to
produce muscle contraction.
The membranes of t-tubules and SR are juxtaposed and permit direct physical
interactions between DHPR and RyR1 in skeletal muscle
(Block et al., 1988). The
skeletal muscle DHPR is composed of the voltage-sensing and pore-forming
1S subunit, intracellular modulatory ß1 subunit, and auxiliary
2
1 and 
1 subunits
(Catterall, 2000;
Flucher et al., 2005). A
tetrad, which is a cluster of four DHPRs, associates with a
Ca2+-releasing RyR1 channel, which is formed with four RyR1
monomers. The RyR1 protein, the largest known ion channel protein, weighs 560
kDa (Takeshima et al., 1989)
and, thus, RyR1 channels can be observed in electron micrographs as dots of
high electronic density
(Franzini-Armstrong and Protasi,
1997). Although a hydrophobic C-terminus domain might contain
several transmembrane domains as well as the channel pore, the exact number
and position of membrane domains is not known. Three RyR isoforms (RyR1, RyR2
and RyR3) are encoded by different genes in mammals
(Fill and Copello, 2002). RyR1
is the most abundant isoform in skeletal muscle. RyR2 predominantly functions
in cardiac muscle and RyR3 is expressed by many tissues, but at relatively low
levels. RyR1-deficient mice do not move because of the absence of E-C coupling
and die from dysfunction of the diaphragm muscles shortly after birth
(Takeshima et al., 1994). The
formation of DHPR tetrads is also impaired in RyR1-deficient myotubes
(Takekura et al., 1995).
Mutations in RyR1, encoded by the RYR1 gene in humans, are involved in a
pharmacogenetic muscle disorder, malignant hyperthermia (MH), and two
congenital myopathies, central-core disease (CCD) and multi-minicore disease
(MmD). Inherited as a dominant trait, MH appears as a hypermetabolic crisis
when a susceptible individual is exposed to certain anesthetics, such as
halothane (MacLennan and Phillips,
1992). CCD is caused by a dominant RYR1 mutation, with two
recessive exceptions, and is characterized by infantile hypotonia and muscle
weakness (Zhang et al., 1993).
Amorphous central cores, which run along the long axis of the muscle fibers,
can be observed in histological sections of CCD muscle. MmD is characterized
by muscle weakness, scoliosis and respiratory insufficiency, but is inherited
through a recessive RYR1 mutation (Engel,
1967; Jungbluth et al.,
2004). MmD is defined by the presence of multiple small cores in
histological sections. Although little is known about the development of
cores, it has been proposed that cores are formed as a secondary cellular
response to isolate regions of defective Ca2+ regulation from
regions of normal Ca2+ homeostasis
(Lyfenko et al., 2004).
In this paper, we characterized the relatively relaxed (ryr) mutant, which was identified as a spontaneous mutation in our breeding stock of zebrafish. Mutants displayed slow swimming due to weak muscle contractions despite normal output from the CNS. Ca2+ transients in the muscle cytosol and RyR1 at the t-tubules were dramatically decreased in mutant fast muscles, suggesting a defect in E-C coupling. In fact, most of the ryr1b mRNA, encoding RyR1, carried a nonsense mutation in relatively relaxed mutants. Analysis of genomic DNA found an insertion in an intron of the ryr1b gene that resulted in aberrant splicing and a premature stop codon. Similar to human MmD, relatively relaxed mutants displayed small amorphous cores in muscle fibers. Interestingly, application of antisense morpholino oligonucleotides against ryr1b that blocked the aberrant splicing in mutants restored normal swimming. These findings suggest that analysis of the relatively relaxed mutant may be useful for understanding the development of amorphous cores and the physiology of MmD.
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MATERIALS AND METHODS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Westerfield,
2000), which meet the guidelines set forth by the University of
Michigan and Nagoya University. ryrmi340 was identified as
a spontaneous mutation in our breeding stock of zebrafish.
Video-recording of zebrafish behavior
Embryonic behaviors were observed and video-recorded using dissection
microscopy. Mechanosensory stimuli were delivered to the tail with forceps.
Videos were captured with a CCD camera (WVBP330, Panasonic) and a frame
grabber (LG-3, Scion Corporation), and were analyzed with Scion Image on a G4
Macintosh (Apple).
Muscle recording
The dissection protocols for in vivo patch recordings have been described
elsewhere (Buss and Drapeau,
2000). Briefly, 48 hpf zebrafish embryos were anaesthetized in
0.02% tricaine (ethyl 3-aminobenzoate methanesulfonate, Sigma), pinned on a
Sylgard dish and immersed in Evans solution (134 mM NaCl, 2.9 mM KCl, 2.1 mM
CaCl2, 1.2 mM MgCl2, 10 mM glucose and 10 mM Hepes at
290 mOsm and pH 7.8). The skin was peeled off to allow access to the
underlying muscles. For electrophysiological recordings, embryos were
partially curarized in Evans solution containing 3 µM d-tubocurarine
(Sigma) without tricaine. Patch electrodes were pulled from borosilicate glass
(Narishige) to yield electrodes with resistances of 3-10 M
. The
electrode was visually guided to patch muscle cells using Hoffman modulation
optics (40x water immersion objective). The electrode solution consisted
of 105 mM potassium gluconate, 16 mM KCl, 2 mM MgCl2, 10 mM Hepes,
10 mM EGTA and 4 mM Na3ATP at 273 mOsm and pH 7.2. Recordings were
performed with an Axopatch 200B amplifier (Axon Instruments), low-pass
filtered at 5 kHz and sampled at 10 kHz. Data were collected with Clampex 8.2
and analyzed with Clampfit 9.0 (both Axon Instruments). Mechanosensory
stimulation was delivered by ejecting bath solution (20 psi, 20 milliseconds
pulse) from a pipette with a 20 µm tip to the tail of the pinned embryo
using a Picospritzer III (Parker Hannifin Corporation) to induce fictive
swimming.
Ca2+ imaging in muscle
The protocols for Ca2+ imaging have been described previously
(Hirata et al., 2004).
Briefly, we injected Calcium Green-1 dextran (10,000 Mr,
Molecular Probes) into one blastomere of 8- to 16-cell-stage progeny of
ryr carrier in-crosses. At 48 hpf, embryos were anaesthetized with
0.02% tricaine and pinned to a Sylgard dish with tungsten wires. The tricaine
was washed out and embryos were bathed in Evans solution with 5 mM of the
muscle myosin inhibitor, N-benzyl-p-toluene sulphonamide (Sigma)
(Cheung et al., 2002) to
immobilize embryos. Mechanosensory stimulation was applied by ejecting bath
solution using a Picospritzer III to the tail of the pinned embryos.
Line-scanning of a Calcium Green-1 dextran-labeled muscle cell at 800 Hz was
performed by confocal microscopy (FV-500, Olympus). After imaging, the
genotype of the embryos was determined by genomic PCR.
Pharmacological treatment
Ruthenium red (Sigma), an inhibitor of RyR1
(Pessah et al., 1985), was
diluted to 0.5 mg/ml in Evans solution and applied to embryos 1 hour before
behavioral assay.
Mapping
ryr carrier fish were crossed with wild-type WIK fish to generate
mapping carriers that were crossed to identify mutants for meiotic mapping to
microsatellites (Gates et al.,
1999; Shimoda et al.,
1999), as described previously
(Bahary et al., 2004). The LN54
radiation hybrid panel was used for the physical mapping
(Hukriede et al., 1999).
Cloning of ryr1b cDNA
Twelve overlapping cDNA fragments covering the coding region of
ryr1b were cloned by reverse transcriptase (RT)-PCR with the
following primers: forward primer 1,
5'-GAGAAAACGCACGGATTTTCTGATTTCTCC-3'; reverse primer 1,
5'-CTGTGTAAAAAGGCCTGTGGTGCTGTAGAT-3'; forward primer 2,
5'-CTGCTCTCGCTCTCAGACAGAAGAGTC-3'; reverse primer 2,
5'-AGCAGTGTCAACAGGACAGGGAGTGAATG-3'; forward primer 3,
5'-CCTCCTCCTGGTTATGCACCATGTTATGAG-3'; reverse primer 3,
5'-TCAATGTGGTTGTGATCAGTGGGAACGGGA-3'; forward primer 4,
5'-CATCTGTGGCCTCCAAGAGGGCTTTG-3'; reverse primer 4,
5'-ACACACGGCGCAATAAAGCATCAGAGTGTG-3'; forward primer 5,
5'-TCCTGGAACTATCTGAGCAACATGACCTGC-3'; reverse primer 5,
5'-AAGACTGAACATGAGTCGCACCAGCTCTG-3'; forward primer 6,
5'-GTTGATATCCCACACAATGATCCACTGGGC-3'; reverse primer 6,
5'-AACAGTGGGGCACATTTAGTGAGCAGAGG-3'; forward primer 7,
5'-CACCACAGAAATGGCTCTGGCTCTGAAC-3'; reverse primer 7,
5'-CTTAAAGTACTGGTTTATGAGCGGGAGCAG-3'; forward primer 8,
5'-GGAAGAGTTGAAAAATCCCCACACGAACAG-3'; reverse primer 8,
5'-ATCACCACAAAGTTCTGCTCCTCTCGCTTG-3'; forward primer 9,
5'-GAAGTCTTCATCTTCTGGTCTAAATCGCAC-3'; reverse primer 9,
5'-TGCAGTCTGAGCAGAAAGTCTACTGTGCAG-3'; forward primer 10,
5'-AACTACCGGCGCACACAAACAGGCAG-3'; reverse primer 10,
5'-ACGTCATCTTTTTAGCTCCCTCCACGAGAC-3'; forward primer 11,
5'-CTTGCTGATCTGGAACACTCTCTTTGGAG-3'; reverse primer 11,
5'-GTGTATTAAGGACTAGTCGGTCCCATTGG-3'; forward primer 12,
5'-CTGGCCCGTAAACTGGAGTTTGATGGTC-3'; reverse primer 12,
5'-AGGAGAGTTAAGTCAAGGCTACAGTCGGTC-3'.
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Knockdown by morpholino
To knockdown RyR1b protein synthesis
(Nasevicius and Ekker, 2000),
antisense morpholino (MO)1 was designed against splice donor site of exon48:
ryr1b MO1, 5'-ATGATTGAGTTTACCGTATCCAGAG-3'.
Standard control MO (randomized sequence available from Gene Tools) was used for control MO: control MO, 5'-CCTCTTACCTCAGTTACAATTTATA-3'.
For inhibition of aberrant splicing of ryr1b mRNA, antisense MO2 and MO3 were respectively designed against acceptor and donor sites of the aberrant exon: ryr1b MO2, 5'-ATTGGTTGACTCCTGATACTCAATG-3'; ryr1b MO3, 5'-TATCTTACACTTACCTTTAAATAAG-3'.
Injections were performed as described previously
(Nüsslein-Volhard and Dahm,
2002).
Immunostaining
Zebrafish embryos were anaesthetized in 0.02% tricaine and pinned on a
Sylgard dish with tungsten wires. After peeling off the skin at trunk region,
embryos were fixed in 4% paraformaldehyde at room temperature for 20 minutes
and then subjected to immunostaining as described previously
(Hirata et al., 2005). The
following primary antibodies were used: anti-RyR: 34C, IgG1 isotype, Sigma,
1/1000 (Airey et al., 1990);
anti-DHPR1: 1A, IgG1 isotype, Affinity BioReagents, 1/200
(Morton and Froehner, 1987);
anti-DHPRß: VD21B12, IgG1 isotype, Developmental Studies
Hybridoma Bank, 1/2 (Leung et al.,
1988); anti-DHPR2: 20A, IgG2a isotype, Affinity
BioReagents, 1/200 (Morton and Froehner,
1989). Alexa Fluor 488-conjugated anti-mouse IgG, Alexa Fluor
488-conjugated anti-mouse IgG1 and Alexa Fluor 555-conjugated anti-mouse IgG2a
were used as secondary antibodies (1/1000, Molecular Probes).
In situ hybridization
In situ hybridizations to wholemounted zebrafish were performed as
described previously (Hirata et al.,
2004). For sectioning after color development, embryos were
equilibrated in 15% sucrose/7.5% gelatin in PBS at 37°C and then embedded
in it at -80°C. Sections (10 µm) were cut with a cryostat (CM3050S,
Leica). An ryr1b probe covering 1089 bp of the C-terminus amino acids
and 148 bp of the 3'-UTR was used for in situ hybridization. An
ryr1a probe covering 1157 bp of the C-terminus was cloned with the
following primers: ryr1a forward primer,
5'-ATGGTGAGAAGGCTGAGAAGGAAGTGGAGG-3'; ryr1a reverse
primer, 5'-CGTCATCATGTCGTCACACTTCATGTCCGG-3'.
Transmission electron microscopy
The protocols for transmission electron microscopy have been described
elsewhere (Hatakeyama et al.,
2004; Schredelseker et al.,
2005). Briefly, embryos were fixed with 6% glutaraldehyde-2%
paraformaldehyde in 0.1 M sodium cacodylate buffer, pH 7.2, overnight at
4°C. After being washed in 0.1 M sodium cacodylate buffer, the embryos
were post-fixed with 1% OsO4 for 60 minutes, and then dehydrated
and embedded in Epon 812. Ultrathin sections (80 nm) were cut and examined
using an electron microscope (H-7000, Hitachi) operated at 75 kV.
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RESULTS |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Ca2+ transient is smaller in ryr mutant fast muscle
Depolarization of the muscle membrane causes a transient increase in
cytoplasmic Ca2+ mediated by E-C coupling that results in
actin/myosin sliding and the contraction of muscle
(Franzini-Armstrong and Protasi,
1997). Because muscles were defective in ryr mutants, we
examined whether the increase in cytosolic Ca2+ was perturbed in
mutant muscles by injecting live embryos with Ca2+ indicator dye,
Calcium Green-1 dextran (Fig.
3A). The amplitude of Ca2+ transients in fast muscle
was 3.3-times smaller in ryr mutants compared with wild-type siblings
at 48 hpf [wild-type relative level of Calcium Green-1 fluorescence
(
F/F): 0.43±0.13, n=7; ryr F/F:
0.13±0.05, n=7; Student's t-test,
P<0.001, Fig. 3B].
By contrast, Ca2+ transients in slow muscle were not perturbed in
ryr mutants (wild-type F/F: 0.35±0.08, n=5;
ryr F/F: 0.35±0.07, n=5;
Fig. 3C). Furthermore,
Ca2+ transients in mutants were comparable with wild-type siblings
at 24 hpf (data not shown), when all mutants responded normally to tactile
stimulation. Thus, a defect in E-C coupling in fast muscles appears to be the
basis for weak contractions in ryr mutant muscles.
Proteins for E-C coupling are not clustered at t-tubule-SR junctions in ryr mutants
E-C coupling is mediated by direct interaction between DHPRs and RyRs, both
of which are clustered at the juxtaposed membranes of t-tubule-SR junctions.
To see how E-C coupling might be defective in mutant fast muscles, the
distribution of RyRs and DHPRs were examined in ryr mutant muscles.
Labeling with anti-RyR showed that RyRs were distributed in a striated pattern
in wild-type fast muscles, which presumably represented the t-tubule-SR
junctions (Fig. 4A), whereas
RyR labeling was significantly reduced in mutant fast muscles
(Fig. 4B). Similarly,
anti-DHPR1, anti-DHPRß and anti-DHPR2, respectively, showed
that DHPR1, DHPRß and DHPR2 were probably localized to
t-tubule-SR junctions in wild-type fast muscles
(Fig. 4C,E,G). Double labeling
with anti-RyR and anti-DHPR2 confirmed that RyRs and DHPRs were
colocalized at presumptive t-tubule-SR junctions
(Fig. 4I). On the other hand,
DHPR1, DHPRß and DHPR2 labeling were significantly reduced
in mutant fast muscles (Fig.
4D,F,H,J). However, the distribution of RyRs and of DHPR subunits
were unperturbed by the mutation in slow muscles
(Fig. 4K-T). Electron
micrographs verified the immunohistochemistry results. Patterned
electron-dense structures that presumably represent juxtaposed RyRs and DHPRs
were present at t-tubule-SR junctions in wild-type fast muscles
(Fig. 4U) but not in
ryr mutant fast muscles (Fig.
4V). The distribution of presumptive RyR/DHPR particles in slow
muscles was comparable between wild type and ryr mutants
(Fig. 4W,X). Thus, ryr
mutants are deficient in E-C coupling in their fast muscles because of a lack
of clustering by RyRs and DHPRs at the t-tubule-SR junctions.
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). Reverse transcriptase (RT)-PCR with primers flanking the
insertion was performed to confirm that the insertion was specifically found
in ryr mutants (Fig.
5C). Both longer and shorter PCR fragments were amplified using
the cDNA from a group of wild-type siblings as template
(Fig. 5C, lane 1), whereas the
longer fragment was predominant in mutants
(Fig. 5C, lane 2). Only the
shorter fragment was amplified from the cDNA of wild-type embryos of another
strain, the AB strain, of zebrafish (Fig.
5C, lane 3), confirming that the shorter band corresponded to
wild-type ryr1b. Sequencing of the longer and shorter fragments
verified that the 32-bp insertion was found only in the longer product. Thus,
the ryr phenotype is very likely to be due to mutation in
ryr1b.
Genomic sequencing revealed that ryr mutants carry a 4046-bp DNA
insertion, including the 32-bp cDNA insertion, in the intron between exon 48
and 49 of the ryr1b gene (Fig.
5D). Sequences flanking the 32 bp in the genomic insert contained
splicing acceptor and donor sites, confirming that the 32-bp sequence acts as
an additional exon in the mutant ryr1b gene. This genomic insertion
might represent a transposable element, because it contained a repeated motif
at both ends that are characteristic of Tc1/mariner family transposons (data
not shown) (Ivics et al.,
2004; Kawakami,
2005). Because the aberrant splicing results in a premature stop
codon that predicts a truncated RyR1b lacking the channel domains, the great
majority of fast muscle RyR1b would probably be non-functional.
To confirm whether a loss of RyR1b is responsible for the ryr
phenotype, we attempted to phenocopy slow swimming by antisense knockdown of
RyR1b and application of a specific inhibitor of RyR. We injected antisense
morpholino oligonucleotides (MO1), which were complementary to the splice
donor site of exon 48, into wild-type embryos and assayed touch responses at
36 hpf. MO1-injected wild-type embryos swam more slowly than control
MO-injected wild-type embryos (MO-1 injected: 0.81±0.20 cm/s,
n=12; control MO-injected: 1.78±0.35 cm/s, n=12;
Student's t-test, P<0.001), much like mutant embryos.
Interestingly, most of the MO1-injected embryos also exhibited weak coils of
the trunk and tail following tactile stimulation at 24 hpf (82.0±7.8%,
n
43-76, five trials) rather than the normal fast, vigorous coils,
suggesting that RyR1b is required for touch-induced coiling at earlier stages
as well as for swimming at later stages. Correlated with the behavioral
defects, the amount of ryr1b mRNA with normal splicing at 24, 36 and
48 hpf in MO1-injected embryos was reduced compared with control MO-injected
embryos (Fig. 5E), confirming
the efficacy of knockdown by MO1. Treatment of wild-type embryos with
Ruthenium red, an inhibitor of RyR (Pessah
et al., 1985), also phenocopied touch-induced slow swimming at 36
hpf (n=20). Thus, RyR1 is essential for normal muscle function in
zebrafish, as it is in mammals.
The fact that weak muscle contractions in mutants were obvious after 36 hpf but not earlier than 30 hpf, whereas wild-type embryos in which ryr1b was knocked down exhibited weak contractions at 24 hpf, suggests that defective splicing was stage-dependent. To test this possibility, the head and trunk of individual embryos from a cross of two ryr carriers were subjected to genomic PCR and RT-PCR, respectively, to assay the genotype and splicing at 24 and 48 hpf (Fig. 5F). Wild-type embryos of the wt/wt genotype showed only a wild-type (short) RT-PCR fragment at both stages (Fig. 5F, lanes 1, 4), whereas wt/ryr embryos exhibited both wild-type (short) and mutant (long) fragments at both stages (Fig. 5F, lanes 2, 5). Mutant embryos (ryr/ryr) gave both wild-type and mutant fragments with comparable intensity at 24 hpf (Fig. 5F, lane 3), whereas the mutant product became predominant at 48 hpf (Fig. 5F, lane 6). These results suggest that the mutant behavioral phenotype was due to stage-dependent aberrant splicing in mutants.
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) and identified as RyR1, RyR2 or RyR3 based on homology with
the human sequences. This classification suggested that there were at least
five different ryanodine receptor genes in zebrafish: two for RyR1
[ryr1a in linkage group (LG)10 and ryr1b in LG18], two for
RyR2 (ryr2a in LG12 and ryr2b in LG17) and one for RyR3
(ryr3 in LG20).
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) (Fig.
6F-I). These results indicate that the potential duplicates of
RyR1 were both expressed by muscles but that only RyR1b is expressed by fast
muscles.
The ryr mutant is a disease model of MmD
MmD, a recessive myopathy, is caused by RYR1 mutations in human and is
pathologically defined by multiple amorphous cores in muscle fibers
(Engel, 1967;
Jungbluth et al., 2004). To
examine whether ryr mutants displayed morphological defects in
muscle, transverse sections of larval axial muscles were analyzed by
transmission electron microscopy. Superficial slow muscles appeared comparable
between wild type and mutant (data not shown). Well-formed actin/myosin
bundles and SR were observed in wild-type fast muscle at 2, 7 and 14 dpf
(Fig. 7A-C). Mutant fast
muscle, however, displayed small amorphous cores (50-100 nm in diameter) at 2
dpf (Fig. 7D). The diameter of
the cores in mutant fast muscles increased with development (50-500 nm at 7
dpf; 100-800 nm at 14 dpf; Fig.
7E,F). Disorganization of the SR was also evident at 7 dpf and, in
some cases, the SR was missing at 14 dpf. Thus, ryr mutant fast
muscles displayed ultrastructural defects similar to those seen in MmD
muscles.
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DISCUSSION |
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TOPSUMMARYINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The zebrafish immotile mutant, relaxed (also known as
cacnb1 - Zebrafish Information Network), is deficient in E-C coupling
because of a null mutation in DHPRß1
(Schredelseker et al., 2005;
Zhou et al., 2006). In
relaxed mutant muscles, DHPR1 was significantly reduced, but
RyR1 was correctly targeted to the t-tubule-SR junctions. In ryr
mutants, by contrast, there was a dramatic decrease in both DHPR and RyR1 even
when the t-tubules and the SR were not damaged. Taken together, the phenotypes
of the two zebrafish mutations corroborate the finding that the formation of
DHPR tetrads requires the presence of RyR1
(Takekura et al., 1995).
Indeed, a cytoplasmic domain of RyR1 is essential for the physical interaction
with an intracellular loop of DHPR1 in cultured mammalian myotubes
(Kugler et al., 2004;
Proenza et al., 2002). The
failure of DHPR to localize to the t-tubule-SR junctions is probably a direct
consequence of the failure of RyR1b to be targeted to the SR in ryr
mutants.
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;
Kawakami, 2005). Although
ryr1b was expressed by fast muscles at 24 hpf and was essential for
E-C coupling at later stages, ryr mutants responded to touch with
normal C-bends at 24 hpf and exhibited slow swimming after 36 hpf. This lack
of a mutant phenotype at 24 hpf can be explained by the possibility that
splicing of ryr1b might be developmentally regulated and that the
aberrant splicing may only occur during later stages. In fact, half of the
ryr1b transcripts were normally processed at 24 hpf, whereas most
ryr1b mRNA transcripts were aberrantly spliced at 48 hpf. A splicing
factor, which determines exon preference, might be differentially regulated
between 1 and 2 dpf. The finding that knockdown of RyR1b translation in
wild-type embryos leads to weak muscle contractions at 24 hpf is consistent
with this hypothesis.
We found that zebrafish have two genes encoding RyR1; ryr1a
expressed by slow muscles and ryr1b by fast muscles. A similar
division of labor between duplicated RyR1s in slow and fast muscles appears in
other fish, such as blue marlin (Makaira nigricans) and yellowfin
tuna (Thunnus albacares) (Franck
et al., 1998; Morrissette et
al., 2000; Morrissette et al.,
2003). Although single-channel analysis indicates that channel
activity of RyR1 in fast muscle is higher than that in slow muscle
(Morrissette et al., 2000),
any functional difference in E-C coupling has not been examined. It would be
interesting to examine potential differences in E-C coupling between slow and
fast muscles to see how the requirements of the two muscle types dictate
divergent RyR1s and how these differences may have evolved.
The ryr mutant is an animal model for MmD
The zebrafish ryr mutant phenotype shares several crucial features
with human MmD. First, mutations in genes encoding RyR1 are responsible for
both the ryr phenotype and MmD. Second, both the ryr
phenotype and MmD are inherited as autosomal recessives. Third, both
ryr mutants and individuals with MmD exhibit muscle weakness. Fourth,
both ryr mutants and individuals with Mmd display myopathy
characterized by minicores in histological sections. The amorphous cores in
MmD (2-25 µm in diameter) are associated with labeling for reductase
activity in mitochondria with NADH-TR (tetrazolium reductase)
(Engel, 1967;
Martin et al., 1986;
Swash and Schwartz, 1981).
Unfortunately, NADH-TR staining appears to not be useful in fish
(Johnston et al., 1975;
Matsuoka and Iwai, 1984).
However, electron microscopy clearly demonstrated amorphous cores in zebrafish
ryr mutant muscles and showed that they are evident in embryonic
muscles. The fact that RyR1-deficient mice die on the day of birth limits
their usefulness as an animal model of MmD
(Takeshima et al., 1994). By
contrast, zebrafish ryr mutants die 7-15 dpf, but their fast
development and accessibility might be useful for detailed physiological and
pathological analysis of the consequences of MmD.
We succeeded in treating muscle weakness in ryr mutants by the
application of an antisense morpholino that increased the normal splicing of
ryr1b and restored normal swimming. Germane to our finding,
antisense-mediated exon skipping restored normal dystrophin expression in
certain dystrophin mutant mice and in cultured muscle cells from Duchenne
muscular dystrophy individuals harboring splicing defects
(Goyenvalle et al., 2004;
van Deutekom et al., 2001).
This novel therapeutic strategy might represent an effective treatment for the
human genetic disease and is under clinical trials
(Muntoni et al., 2005;
Wilton and Fletcher, 2006).
Interestingly, Monnier and her colleagues reported a hypomorph in human RYR1
that was responsible for MmD (Monnier et
al., 2003), similar to the zebrafish ryr mutants. In this
case, an aberrant splice donor site was generated by a point mutation in an
intron, resulting in incorrect splicing and in the generation of a nonsense
codon in most of the human RYR1 transcripts. Given the similarity to
ryr mutants, this case of MmD might be treated by blocking aberrant
splicing with antisense reagents.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/134/15/2771/DC1
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ACKNOWLEDGMENTS |
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Shimoda, N., Knapik
