Myo18b is essential for sarcomere assembly in fast skeletal muscle

Abstract

Congenital myopathies are muscle degenerative disorders with a broad clinical spectrum. A number of myopathies have been associated with molecular defects within sarcomeres, the force-generating component of the muscle cell. Whereas the highly regular organization of the myofibril has been studied in detail, in vivo assembly of sarcomeres remains a poorly understood process. Therefore, a more detailed knowledge of sarcomere assembly is crucial to better understand the pathogenic mechanisms within myopathies. Recently, mutations in myosin XVIIIB (MYO18B) have been associated with cases of myopathies, although the underlying mechanism for the resulting pathology remains to be defined. To analyze the role of myosin XVIIIB in skeletal muscle disease, zebrafish mutants for myo18b were generated. Full loss of myo18b function results in a complete lack of sarcomeric structure, revealing a highly surprising and essential role for myo18b in sarcomere assembly. Importantly, scattered thin and thick filaments assemble throughout the sarcoplasm; but fail to organize into recognizable sarcomeric structures in myo18b null mutants. In myo18b partial loss-of-function mutants sarcomeric structures are assembled, but thin and thick filaments remain misaligned within these structures. These observations suggest a novel model of sarcomere assembly where Myo18b coordinates the integration of preformed thick and thin filaments into the sarcomere. Disruption of this highly coordinated process results in a block in sarcomere biogenesis and the onset of myopathic pathology.

Introduction

Knowledge about the myofibril is crucial for a better understanding of the molecular basis of the pathological onset of myopathies. The organization of the myofibril and its basic subunit the sarcomere is well characterized. The main components of sarcomeres are myosin-rich thick filaments interdigitated with thin filaments that are mainly comprised of actin. Thin filaments are anchored in the Z-disk and by sliding past thick filaments contraction force is generated. In contrast to sarcomere organization, the initial process of sarcomere assembly remains poorly understood. Two prominent models of sarcomere assembly have been proposed: One model suggests that randomly scattered I-Z-I bodies of Z-disk associated proteins and thin filaments recruit titin, which subsequently associates with independently assembled thick filaments to form sarcomeres (1,2). Another model suggests that thin filaments, actinin and nonmuscle myosin II form regularly patterned minisarcomeres called premyofibril (3). Nonmuscle myosin II is subsequently replaced by muscle myosin II and titin is added. Also the distance between Z-bodies is widened, as they align in register and mature into Z-disks (4,5). While these two models partially overlap, they are mainly distinguished by the proposed role of titin and the spacing of the Z-bodies, which are suggested to be organized in either randomly scattered I-Z-I bodies or regularly arranged in minisarcomeres. In addition, other models have been brought forward that propose integrin adhesion sites serve as nucleation sites for the assembly of Z-bodies of the premyofibril (6) or a more recent model that emphasizes the impact of mechanical tension enabled by muscle-tendon attachment for myofibrillogenesis (7).

Congenital myopathies comprise a diverse group of muscle disorders that are diagnosed by skeletal muscle weakness and hypotonia with absence of dystrophic characteristics, such as myofiber detachment followed by myofiber necrosis and fibrosis. However, classification of myopathies can be challenging, because the genetic basis of many myopathies remains unresolved to date (8). Members of the myosin superfamily, which comprises ATP-dependent motor proteins that generate force by sliding over actin filaments, have been implicated in with different types of myopathy (9). One of the best-described myosins is myosin II, which in skeletal muscle forms hexameric complexes of two heavy and four light chains (10). Myosin heavy chains share three common regions: the motor domain that hydrolyses ATP and interacts with actin filaments, the neck region that binds light chains via IQ motifs to enable motor domain movement and finally the tail domain that forms rod-shaped coiled-coil structures by dimerization (11). In skeletal muscle, multiple myosin complexes are assembled to form the thick filaments that are subsequently integrated into sarcomeres, the basic unit of the myofibril.

Phylogenetic analysis suggests that class II myosins share common origins with class XVIII myosins as they have a typical motor domain, one IQ motif and a tail region with putative coiled-coil domains in common (12). In contrast to the extensively studied myosin II, however, myosin XVIIIB (MYO18B) has only recently been discovered and its function still remains elusive (13). Myo18b-deficient mice are characterized by myocardial defects and disordered alignment of thick and thin filaments within sarcomeres (14). However, although myo18b is expressed also in skeletal muscle of the mouse, skeletal muscle defects have not been reported in Myo18b-deficient mice.

MYO18B function has been implicated in diverse human syndromes and cellular processes. Statistical analysis of a genome-wide association study has suggested that a specific single nucleotide polymorphism (SNP) in MYO18B might contribute to mathematical disability (15), a result brought into doubt by a more recent replication study (16). MYO18B function is often lost in cancer cell lines, suggesting MYO18B as a tumor suppressor (17). Importantly, null mutations in MYO18B have also been detected in patients suffering from myopathies. A non-sense mutation in the last coding exon of MYO18B was discovered by exome sequencing in a patient presented with severe nemaline myopathy and cardiomyopathy (18). Interestingly, MYO18B protein was only detected in the patient’s muscle biopsies by antibodies against the N-terminal region of MYO18B, but not with antibodies against the C-terminus. In another case report, two patients from unrelated families were presented with characteristics of nemaline myopathy, Klippel-Feil anomaly and facial abnormalities (19). Mapping followed by exome sequencing revealed the same non-sense mutation in the last coding exon of MYO18B in both patients. In addition, real-time PCR revealed near-complete loss of MYO18B transcript, indicating nonsense-mediated decay of the transcript (19).

In order to study the process of sarcomere assembly and gain novel insights into human myopathies, we performed a genetic screen in zebrafish to systematically identify novel molecules that play a role in sarcomere assembly. By utilizing birefringence as a direct indicator for muscle integrity, a novel mutant was isolated that we linked to a chromosomal region that contained partially predicted fragments of myo18b. Analysis of partial and full loss-of-function of myo18b revealed that in skeletal muscle, Myo18b is integrated into sarcomeres and thereby plays an essential role for sarcomere assembly.

Results

Identification of the muscle mutant schläfrig

To study unresolved congenital muscle disorders, we performed a forward genetic screen based on muscle birefringence to identify novel genes that regulate sarcomere biogenesis. Birefringence is a light effect provoked by the diffraction of polarized light through the pseudo-crystalline array of the muscle myofibril, which causes muscle fibers to appear bright in an otherwise dark environment. Therefore, any reduction in the amount of myofibril, either by muscle atrophy or damage, can be detected by quantification of birefringence (20). Random mutations were introduced in male zebrafish by the mutagen N-ethyl-N-nitrosourea (ENU) and 126 F2 families, established by out-crossing of ENU-treated males were screened for a reduction in birefringence at 3 days post fertilization (dpf) (21). One of the mutants identified appeared unremarkable under bright-field microscopy (Fig. 1A). However, polarized light revealed a strong reduction of birefringence in homozygous mutants, which indicates deficiencies in muscle integrity (Fig. 1B). Subsequent quantification of the birefringence revealed a highly significant reduction to 52±1% within homozygous mutants compared to their siblings (Fig. 1C). In line with the reduction in birefringence, mutants also showed impaired motility in a touch response assay (Supplementary Material, Movies 1, 2) and homozygotes did not survive 14 dpf. Accordingly, these mutants were named schläfrig (sig), which is German for tired. Our previous analyses indicated focal loss of birefringence was a common feature of dystrophic models that exhibit a stochastic detachment of fibers from the myosepta, whereas a global loss of birefringence was found to occur in mutations that disrupt sarcomere assembly or maintenance (20). Furthermore, dystrophic musculature is characterized by myofiber detachment and subsequent myofiber necrosis and fibrosis (22). Thus, to better to define the nature of the muscle defect in sig mutants, whole mount immunohistochemistry with antibodies against dystrophin was undertaken, which revealed that dystrophin localized to the sarcolemma as in siblings and myofibers were attached to the myotendinous junction (Fig. 1D). Cross sections at 3 dpf appeared comparable between sig mutants and wildtype siblings, with neither fibrotic tissue nor necrotic fibers evident in sig mutants (Fig. 1E). Overall, this analysis suggested that the sig phenotype most likely resembles a myopathy like pathology rather than a dystrophic condition.

Figure 1

Muscle birefringence is reduced in schläfrig (sig). (A) Under bright-field microscopy, homozygous sig mutants appear comparable to their siblings. (B) Polarized light reveals a reduction of birefringence in sig homozygotes compared to their siblings. (C) The birefringence of sig mutants is significantly reduced after normalization to siblings. Data are represented as mean ± s.e.m, ∗∗P < 0.01 (Student’s t test). (D) Antibodies against dystrophin mark somite borders in both genotypes, indicating that the muscle in sig is not dystrophic. (E) Myofibers of siblings and sig homozygotes appear comparable on cross sections of 3 dpf-old larvae stained with H&E.

Splicing of myo18b is altered in schläfrig mutants

sig mutants were subjected to a SNP-based linage analysis to identify the causative mutation. 25 sig homozygotes and siblings were phenotyped based on their reduction in birefringence and their isolated genomic DNA was sequenced. Obtained sequences were analyzed with the web-based software SNPtrack that located a region of homozygosity on chromosome 10 between 41 and 46 million base pairs (Mbp) (Fig. 2A) (23). Within this region of homozygosity, the Ensemble genome assembly (Zv9) predicted the presence of three gene fragments based on their sequence homology to human myosin XVIIIB (MYO18B): CABZ01087088.1, myo18b (2 of 3) and CABZ01073112.1 (Fig. 2B). RT-PCR and RACE were performed with oligonucleotides targeting predicted gene fragments and subsequently combined to generate myo18 cDNA (submitted under accession KX485371). Full-length zebrafish myo18b cDNA is comprised of 6713 base pairs (bp) that encode for Myo18b that is 47.7% identical and 77.2% similar to human MYO18B. In the current genome assembly GRCz10, the myo18b gene is scattered on the forward and reverse strand of chromosome 10 within 44.3 to 45.6 Mbp and one internal exon is designated to chromosome 12, which provides an explanation for the broad region of homozygosity identified by the linkage analysis (Supplementary Material, Fig. S1). Sequence analysis using the InterPro software suggests that zebrafish Myo18b contains domains typically found within myosin heavy chains (24): a motor domain, one IQ motif and five coiled-coil regions (Fig. 2C). RT-PCR followed by densitometric analysis of myo18b transcript revealed that the splicing of 41±2% of the myo18b transcript is altered in 3-dpf-old sig homozygotes (Fig. 2D). Subsequent sequencing revealed that the intron between exons 21 and 22 is not fully removed and a residual fragment of 105 bp remains in the altered myo18b transcript. This insertion harbors additional in-frame pre-mature stop codons, indicating that the aberrant myo18b transcript does not encode for functional Myo18b (Fig. 2E and F). However, wildtype (WT) myo18b transcript was still detected in sig homozygotes, indicating that sig mutants harbor a partial loss-of-function allele of myo18b. To identify the molecular cause of the aberrant splicing of myo18b in sig homozygotes, genomic PCR was performed with primers annealing to exon 21 and the insertion. However, genomic PCR only produced an amplicon with genomic DNA isolated from siblings but not sig homozygotes, suggesting that the splice aberration is caused by a larger genomic insertion (Fig. 2G). Based on the discovered splice alteration, the novel muscle mutant sig is subsequently referred to as myo18b^sig.

Figure 2

sig mutants harbor a splice site mutation in myo18b. (A) Linkage analysis of sig mutants located a region of homozygosity on chromosome 10. (B) Allocated within the linked region of Zv9 were three predicted gene fragments that were identified based on their homology to human MYO18B, namely CABZ01073112.1, CABZ01087088.1 and myo18b (2 of 3). (C) Cloning of zebrafish myosin18b resulted in a transcript encoding for a 2103 amino acid protein containing a motor domain, an IQ motif and five coiled-coil regions. (D) RT-PCR using oligonucleotides targeting exons 21 and 27 resulted in an 819-bp amplicon with wild type larvae and an additional 924-bp amplicon with sig homozygotes. (E) Sequencing revealed that, in contrast to the WT myo18b transcript, (F) the aberrant transcript from sig carried a 105-bp insertion that harbored in-frame pre-mature stop codons. (G) Genomic PCR using primers targeting exon 21 and the insertion produces a 474-bp amplicon only with genomic DNA isolated from siblings but not sig homozygotes. (H) At 1 dpf, in situ hybridization detected robust myo18b expression in the trunk musculature of WT larvae (arrowhead), but absence of myo18b in the superficial slow muscle (arrowhead) as well as the spinal cord and the notocord. (I) In addition to the trunk muscle, the fin muscle (arrow) as well as the heart expressed myo18b at 2 dpf. (J) At 3 dpf, myo18b transcript was detected in head as well as extraocular muscles.

Next, in situ hybridization was performed to examine if the expression pattern of myo18b is correlated with the phenotype of myo18b^sig. In line with the muscle specific defects evident in myo18b^sig homozygotes, robust myo18b expression was detected in the trunk muscle of WT larvae at 1 dpf. However, cross sections revealed that myo18b transcript was restricted to differentiated fast muscle and absent in the superficial layer of slow-twitch myofibers as well as the central nervous system and the notochord (Fig. 2H). At 2 dpf, myo18b transcript was additionally identified in the fin and heart muscle (Fig. 2I). Expression of myo18b within the head muscle and the extraocular muscles becomes more pronounced at 3 dpf (Fig. 2J). Thus, the myo18b expression in zebrafish is similar to that described in human tissue and its muscle specific expression is in accordance with the reduction in birefringence evident within myo18b^sig homozygotes (13).

myo18b ^{− 9 + 3} harbors a nonsense mutation in myosin XVIIIb

To further validate the myo18b^sig mutant, the CRISPR/Cas9 system was utilized to introduce mutations into zebrafish myo18b. A single guide RNA targeting exon 6 of myo18b was co-injected with Cas9 into WT eggs (Fig. 3A). Germline transmission was achieved for an allele harboring a deletion of 9 bp and an insertion of an in-frame pre-mature stop downstream of the E219 codon (Fig. 3B). Importantly, the robust myo18b expression within 3-dpf-old siblings was markedly reduced in myo18b ^{− 9 + 3} homozygotes, which is indicative of nonsense-mediated decay of the myo18b transcript (Fig. 3C). In line with the heart expression of myo18b described above, myo18b ^{− 9 + 3} homozygotes exhibit a pronounced heart edema at 3 dpf (Fig. 3D). In addition, myo18b ^{− 9 + 3} do not survive to 11 dpf and do not inflate their swim bladders, indicating that the phenotype of myo18b ^{− 9 + 3} is more severe compared to myo18b^sig. Whereas the birefringence of heterozygous myo18b ^{− 9 + 3} was comparable to WT siblings, homozygous myo18b ^{− 9 + 3} presented a significantly reduced birefringence that was further reduced compared to myo18b^sig, another indication that the loss of muscle integrity is more evident in myo18b ^{− 9 + 3} homozygotes (Fig. 3E and G). Compound myo18b ^{− 9 + 3};myo18b^sig heterozygous larvae presented with a significant reduction in birefringence that was midway between the birefringence evident in single myo18b^sig and myo18b ^{− 9 + 3} homozygotes (Fig. 3F and G). Taken together, these results confirm that the phenotype of myo18b^sig is caused by a myo18b partial loss-of-function allele and suggests that myo18b ^{− 9 + 3} mutants harbor a full loss-of-function allele.

Figure 3

Muscle phenotype of myo18b ^{− 9 + 3}. (A) CRISPR/Cas9 technology was utilized to target exon 6 of myo18b, (B) which resulted in a 9 bp deletion and simultaneous pre-mature stop codon insertion. (C) At 3 dpf, whole-mount in situ hybridization shows myo18b expression in siblings that is strikingly reduced within myo18b ^{− 9 + 3} mutants, suggesting nonsense-mediated decay. (D) In comparison to their siblings, myo18b ^{− 9 + 3} mutants are bent and present with a pronounced heart edema. (E) Homozygotes of myo18b ^{− 9 + 3} showed a reduction in birefringence compared to their siblings. (F) Also 3 dpf-old heterozygous myo18b^sig; myo18b ^{− 9 + 3} compound heterozygotes showed a reduction in birefringence, confirming that the muscle phenotype of myo18b^sig results from a mutation in myo18b. (G) Quantification of the birefringence of myo18b^sig/^−9 + 3 at 3 dpf revealed a highly significant reduction after normalization to siblings, which was a stronger reduction than detected in myo18b^sig homozygotes. myo18b ^{− 9 + 3} homozygotes have an even further reduced birefringence, indicating that insertion of the premature stop codon leads to myo18b loss of function. Data are represented as mean ± s.e.m, ∗∗P < 0.01 (Student’s t test). (H) Transgenic Tg(acta1:lifeact-GFP) labels thin filaments that in siblings are arranged in the typical striated pattern of the myofibril. In myo18b ^{− 9 + 3}, however, marked thin filaments appear devoid of any structure. (I) Transmission electron micrographs of myofibers of siblings depict highly organized sarcomeres. (J) Homozygotes of myo18b ^{− 9 + 3} are devoid of sarcomere structures. (J’) Higher magnification of the box in I reveals bundles of thick and thin filaments dispersed in different directions (red and green arrowheads, respectively). (K) Abundant filaments deposits, in which thin (yellow arrowheads) and thick (red arrowheads) filaments could be distinguished via their electro-density, were found throughout myofibers. (L) In posterior somites of 18-somite embryos of myo18b ^{− 9 + 3} homozygotes and siblings, antibodies against α-Actinin mark Z-bodies that in myo18b ^{− 9 + 3}, however, were not regularly spaced. (M) Transiently expressed GFP-tagged Myo18b integrated into sarcomeres of both myo18b ^{− 9 + 3} and siblings, as identified by the striated fluorescence pattern of GFP. (N) Immunohistochemistry using antibodies against α-Actinin revealed that Myo18b-GFP was integrated in-between sarcomeric Z-disks.

myo18b ^{− 9 + 3} mutants are devoid of sarcomeric structures within fast-twitch fibers

In accordance to the significant reduction in birefringence detected in myo18b ^{− 9 + 3} homozygotes, the touch response assay revealed that the mobility of myo18b ^{− 9 + 3} mutants is severely impaired (Supplementary Material, Movies 3, 4). To further analyze the myofibril defect, myo18b ^{− 9 + 3} was out-crossed to the transgenic line Tg(acta1:lifeact-GFP), in which the Lifeact-GFP protein binds and thereby marks thin filaments (25). Whereas GFP fluorescence in siblings presented the typical striation pattern of the myofibril, myo18b ^{− 9 + 3} larvae showed abundant fluorescence that was devoid of any striation, indicative of a vast amount of isolated thin filaments in all myo18b ^{− 9 + 3} homozygotes examined (n = 3) (Fig. 3H). Transmission electron microscopy (TEM) revealed organized sarcomeres in myofibres of 3 dpf-old WT siblings (Fig. 3I). In myo18b ^{− 9 + 3} homozygotes, however, sarcomeric structures were not detected at all (Fig. 3J). Instead, deposits of dispersed thick and thin filaments were apparent in myo18b ^{− 9 + 3} homozygotes that featured various orientations and seemed devoid of any pattern or organization (Fig. 3J’ and K). Similar to the vast amount of isolated thin filaments detected in the background of Tg(acta1:lifeact-GFP), serial sections of three different myo18b ^{− 9 + 3} homozygous larvae analyzed by TEM revealed abundant deposits of isolated filaments and striated myofibril was not detected.

As a first step of myofibrillogenesis, the Sanger model proposes formation of an intermediary premyofibril that is characterized by regularly spaced Z-bodies (3). The spacing between these α-actinin-positive Z-bodies is subsequently widened while they fuse and mature into Z-disks (5). To analyze if intermediary structures of myofibrillogenesis are formed within myo18b ^{− 9 + 3} mutants, immunohistochemistry with antibodies against α-Actinin was performed in larvae at the 18-somite stage, in which skeletal muscle formation is initiating within the posterior somites. Within siblings at the 18-somite stage, linear bands of α-actinin-positive Z-bodies were detected, in which the distance between Z-bodies was 1.34±0.03 µm (Fig. 3L). This distance between the Z-bodies of siblings of the 18-somite stage was significantly shorter compared to the distance of 2.04±0.01 µm between mature Z-disks of 3-dpf-old siblings, which were also marked by α-Actinin antibody (Student’s t-test, P < 0.01, n = 3 embryos per age group). This result indicates that the linear bands of actinin-positive Z-bodies detected within siblings of the 18-somite stage likely represent premyofibril. Actinin-positive Z-bodies were also evident within myo18b ^{− 9 + 3} homozygotes at the 18-somite stage (Fig. 3L). Intriguingly however, these Z-bodies were not in registered alignment, as their spacing was irregular.

To study the molecular basis of sarcomere deficiency in myo18b ^{− 9 + 3}, GFP-tagged Myo18b was transiently expressed in muscle fibers under the control of the muscle-specific unc-45b promoter (26). Fluorescence of Myo18b-GFP in 3-dpf-old siblings revealed a striated pattern within myofibers, demonstrating that Myo18b integrates into sarcomeres (Fig. 3M). The same fluorescence pattern was also detected in homozygous myo18b ^{− 9 + 3} larvae, indicating that expression of Myo18b-GFP rescued the loss of sarcomere formation evident within myo18b ^{− 9 + 3} homozygotes confirming that integration of Myo18b into the sarcomere is essential for the sarcomere assembly. To identify the location of Myo18b-GFP in sarcomeres, immunohistochemistry with antibodies against the Z-disk marker α-Actinin was performed. Fluorescence analysis revealed that α-Actinin did not colocalize with Myo18b-GFP in transiently expressing myo18b ^{− 9 + 3} homozygotes, confirming that Myo18b-GFP integrates between Z-disks within sarcomeres, where also the A-band is located (Fig. 3N).

In summary, ablation of myo18b leads to loss of sarcomeres. Transgenic Myo18b-GFP rescues loss of sarcomeres by its integration into sarcomeric A-bands that harbors myosin thick filaments. Nonetheless, dispersed single thick and thin filaments were detected in myo18b ^{− 9 + 3} homozygotes, indicating that myo18b is essential for sarcomere rather than filament assembly.

Sarcomeres of myo18b^sig mutants are disorganized

As the null mutation myo18b ^{− 9 + 3} results in a complete failure of sarcomere assembly, it provides limited insights into the mechanistic basis of this failure. We therefore sought to examine sarcomere assembly defects in the partial loss-of-function allele myo18b^sig more fully. To analyze the sarcomeres, myo18b^sig was crossed to the double-transgenic background of Tg(acta1:lifeact-GFP) and Tg(acta1:mCherryCaaX), so that in addition to thin filaments labeled by lifeact-GFP the sarcolemma and t-tubules of skeletal muscle are also marked by mCherry via the use of the CaaX-tag. Transgenic Lifeact-GFP revealed a highly organized myofibril in WT siblings together with regular spaced t-tubules marked by mCherryCaaX (Fig. 4A). In contrast, thin filaments marked by Lifeact-GFP were largely unorganized in myo18b^sig homozygotes and myofibril striation as well as the regular pattern of t-tubules was only occasionally observed, indicating residual sarcomere formation within myo18b^sig mutants (Fig. 4A). To analyze sarcomere organization in more detail, TEM was performed. Highly organized sarcomeres were found in siblings (Fig. 4B). In myo18b^sig homozygotes only slow fibers contained highly organized sarcomeres (Fig. 4C). The fast fibers of myo18b^sig homozygotes, in contrast, were devoid of substantial higher-order structures and featured sarcomeres with misaligned thin and thick filaments (Fig. 4C–C’’’). Interestingly, the disorganized sarcomeres of myo18b^sig featured regularly spaced Z-disks, indicating that Z-disk architecture is not affected by myo18b partial loss-of-function (Fig. 4C). In addition to disorganized sarcomeres, myo18b^sig homozygotes presented abundant thin and thick filaments scattered throughout the sarcoplasm, which indicates that polymerization of both filaments occurs independent from each other and does not require myo18b function (Fig. 4C’,C’’). Importantly, in addition to misorganised sarcomeres, Z-disk structures with attached filaments were detected within myo18b^sig homozygotes that appeared to be randomly scattered throughout the sarcoplasm (Fig. 4C’’’).

Figure 4

Myofibril of the fast musculature is disorganized in myo18b^sig mutants. (A) Transgenic Tg(acta1:lifeact-GFP) and Tg(acta1:mCherryCaaX) larvae, which mark thin filaments and sarcolemma respectively, demonstrate that the typical striation of the myofibril detected in 3 dpf-old siblings is rarely found in myo18b^sig (arrowhead). (B) Organized sarcomeres are visualized in TEM micrographs of 3 dpf-old siblings. (C) Whereas slow myofibers of 3 dpf-old myo18b^sig homozygotes feature organized sarcomeres (bracket), sarcomeres of fast myofibers appear disorganized; arrowheads mark the sarcolemma. (C’) Abundant deposits of isolated filaments were found in myo18b^sig. (C’’) Isolated thin and thick filaments (yellow and red arrowheads, respectively) of different electron-density were detected in myo18b^sig mutants. (C’’’) In myo18b^sig, thin and thick filaments often form Z-disks that are devoid of other sarcomeric structures, indicating that Z-disks might be the first assembly scaffold assembled in the forming sarcomeres. (D) The single-twitch contraction generated by 5-dpf-old single larvae of both myo18b mutants (myo18b ^{− 9 + 3} and myo18b^sig) was weaker compared to their siblings that feature an overlapping active force trace. (E) Whereas siblings generated a maximal active force of 0.98±0.07 mN, myo18b^sig homozygotes produced significantly less force of 0.27±0.01 mN. Ablation of the force generated by the fast muscle via administration of BTS significantly reduced the active force of myo18b^sig homozygotes further to 0.025±0.003 mN. (F) The active force production of myo18b ^{− 9 + 3} homozygotes was significantly reduced to 0.006±0.002 mN compared to their siblings which generate 1.04±0.09 mN, indicating that the musculature of myo18b ^{− 9 + 3} homozygotes is weaker compared to that of myo18b^sig homozygotes. Homozygotes myo18b ^{− 9 + 3} larvae treated with BTS produced an active force of 0.008±0.002 mN, which was not further reduced compared to untreated myo18b ^{− 9 + 3} homozygotes. However, the force generated by BTS treated siblings was significantly higher than the one generated by myo18b ^{− 9 + 3} homozygotes. Data are represented as mean±s.e.m, ∗∗P<0.01 (Student’s t test).

Overall, the partial loss-of-function mutant myo18b^sig confirms that mutations within myo18b can directly affect sarcomere assembly. In addition, independently polymerized thin and thick filaments and randomly scattered complexes of Z-disks with filaments attachments were detected in myo18b^sig homozygotes that arrange into disorganized sarcomeric structures. However, filaments within these structures remain misaligned and highly organized sarcomeres do not form within fast fibers of myo18b^sig mutants.

The muscle pathology of myo18b^sig and myo18b ^{− 9 + 3} mutants results in muscle weakness

The physiological consequences of defective myofibril formation in myo18b mutants were analyzed by subjecting whole-mount 5-dpf-old larvae to mechanical force measurements using a muscle physiology force transducer (27). Single twitch stimulation revealed that the active force generated by both myo18b mutants was reduced compared to their siblings, with myo18b ^{− 9 + 3} homozygotes showing the weakest active force production (Fig. 4D). Subsequent quantification of the maximal active force showed that the reduced force generation by myo18b^sig homozygotes was highly significant (Fig. 4E). To analyze if the disorganized sarcomeres evident within the fast muscle of myo18b^sig homozygotes are still capable of force generation, N-benzyl-p-toluene sulphonamide (BTS), a potent and specific inhibitor of fast muscle myosin II, was added to specifically block the contraction of fast muscle only (28). Treatment of myo18b^sig homozygotes with BTS significantly reduced the active force generated by mutants to levels below the one generated by untreated homozygotes. Similarly, the maximal active force generated by myo18b ^{− 9 + 3} homozygotes was significantly reduced compared to their siblings, with myo18b ^{− 9 + 3} homozygotes generating a residual active force of 0.006±0.002 mN (Fig. 4F). However, administration of BTS to myo18b ^{− 9 + 3} homozygotes did not further reduce the active force generated by untreated myo18b ^{− 9 + 3} mutants, indicating that the residual force generated by myo18b ^{− 9 + 3} homozygotes can be attributed to the slow muscle. In comparison to BTS-treated siblings BTS-treated myo18b ^{− 9 + 3} homozygotes generated a significantly lower active force, which could be suggestive of weaker slow-twitch fibers in myo18b ^{− 9 + 3} compared to siblings (Fig. 4F). It is important to note that force transduction was measured over whole mounts and therefore external factors such as compromised fast musculature could contribute to the reduced force generated by slow fibres of myo18b ^{− 9 + 3} mutants.

The results therefore indicate that the defective sarcomere formation evident within the fast-twitch musculature of myo18b^sig mutants results in skeletal muscle weakness and that loss of sarcomeres evident within the fast-twitch muscle of myo18b ^{− 9 + 3} homozygotes leads to loss of force generation. By contrast, the sarcomeres of the slow-twitch fibers detected within both myo18b mutants are capable of force generation.

Myofibril defects within myo18b ^{− 9 + 3} and myo18b^sig mutants are specific for the fast-twitch fibers

Slow muscle in zebrafish forms a separate, superficial layer of myofibers that, in contrast to fast muscle, is oriented parallel to the axis of body. As the in situ hybridization results described above indicated that expression of myo18b is restricted to the fast musculature, immunohistochemistry with antibodies against α-Actinin and slow muscle specific F59 antibodies was performed to analyze if sarcomeres of the slow-twitch fibers of myo18b ^{− 9 + 3} and myo18b^sig mutants exhibit any defects. In-line with the analyses performed in the transgenic background of Tg(acta1:lifeact-GFP) and on TEM micrographs, fast-twitch fibers of myo18b ^{− 9 + 3} homozygotes did not show a regular pattern of actinin-positive Z-disks comparable to their siblings, thereby confirming loss of sarcomeres in myo18b ^{− 9 + 3} (Fig. 5A). Superficial slow fibers of myo18b ^{− 9 + 3} homozygotes appeared irregular in register. However, slow fibers of both genotypes featured regularly spaced actinin-positive Z-disks, indicating that sarcomeres form normally in slow myofibers also in myo18b ^{− 9 + 3} homozygotes. This result was confirmed on sagittal sections utilizing the slow fiber specific antibody F59, which revealed myofibril striation within slow fibers of myo18b ^{− 9 + 3} homozygotes and siblings (Fig. 5A). Similarly, siblings of myo18b^sig showed a regular pattern of actinin-positive Z-disks in both fiber types, which was severely compromised in fast but not slow myofibers of myo18b^sig homozygotes (Fig. 5B). Therefore, the musculature defects in both myo18b mutants are specific for the fast and not slow muscle, as foreshadowed by the restricted expression of myo18b.

Figure 5

In both myo18b mutants slow muscle is unaffected, but the head cartilage is misshaped in myo18b ^{− 9 + 3} mutants (A) Immunohistochemistry using antibodies against α-Actinin revealed regularly spaced Z-disks in slow and fast myofibers of 3-dpf-old siblings. In contrast to the fast muscle, the superficial slow fibers that orientate in parallel to the horizontal myoseptum show the typical myofibril striation in myo18b ^{− 9 + 3} homozygotes (arrowheads). Slow fiber specific F59 antibodies, applied to sagittal sections of 3-dpf-old larvae, confirmed intact myofibril striation in slow fibers of myo18b ^{− 9 + 3} homozygotes. Magnified areas are indicated by boxes. (B) Within 3-dpf-old myo18b^sig homozygotes, the regular spacing of actinin-positive Z-disks was detected in the slow-twitch fibers. However, in contrast to siblings, the fast-twitch fibers rarely featured regular Z-disk spacing. (C) At 3 dpf, immunohistochemistry with antibodies against myosin revealed organized myofibrils in the head musculature of siblings and loss of myofibril organization within myo18b ^{− 9 + 3} homozygotes. (D) At 6 dpf, Alcian blue and Alizarin red staining visualized malformed head cartilages within myo18b ^{− 9 + 3} homozygotes compared to their siblings. Label of pharyngeal cartilages: ceratohyal (ch); Meckel’s cartilage (m); palatoquadrate (pq). Note the increased angle between ceratohyal cartilage and midline indicated by dashed lines. (E) In contrast to their 3-dpf-old siblings, the myofibril of myo18b^sig homozygotes appears wavy and reduced in its amount, as revealed by antibodies against myosin. (F). Homozygotes of myo18b^sig did not present with marked malformations of the pharyngeal cartilage that was stained with Alcian blue.

Pharyngeal cartilage is malformed within myo18b ^{− 9 + 3} homozygotes

Within zebrafish, the morphology of the pharyngeal cartilage depends on the contraction force generated by the attached muscle (29). Therefore, zebrafish mutants with weakened head musculature are characterized by malformations of the pharyngeal cartilage (30). To analyze the head musculature of myo18b mutants, which expresses myo18b as shown above, immunohistochemistry was performed with antibodies against myosin. As similarly detected in the transgenic background of Tg(acta1:lifeact-GFP), the typical myofibril striation detected in siblings is lost in the head muscles of myo18b ^{− 9 + 3} homozygotes (Fig. 5C). Accordingly, Alcian blue and Alizarin red staining performed on 6-dpf-old larvae revealed that the head cartilage of myo18b ^{− 9 + 3} homozygotes was characterized by severe malformations in comparison to their siblings (Fig. 5D). Specifically the ceratohyal cartilage within myo18b ^{− 9 + 3} mutants displayed a pronounced increase of the angle between the ceratohyal cartilage and the midline, which is in accordance to weakened head muscles. Importantly, the malformations of the head cartilages evident within myo18b ^{− 9 + 3} homozygotes were similarly to those found within patients affected by MYO18B deficiency, who presented with characteristic facies, including ptosis with compensated arching eyebrows, everted lower lip and bulbous nose with hypoplastic alae nasi (19). In contrast to myo18b ^{− 9 + 3} homozygotes, the head muscles of myo18b^sig homozygotes feature myofibril, which nonetheless appeared wavy and reduced in its amount when compared to siblings (Fig. 5E). Accordingly, myo18b^sig mutants were not characterized by the severe malformations of the head cartilage evident within myo18b ^{− 9 + 3} homozygotes (Fig. 5F).

Discussion

A deeper knowledge of the sarcomere assembly is required to unravel the molecular basis of the pathologies associated with genetic defects of the human sarcomere. The current literature associates a wide range of pathological consequences to mutations within MYO18B, including myopathy. Our analyses on the function of zebrafish myo18b identified an essential role for myo18b in the assembly of sarcomeres and the associated skeletal muscle phenotype of the zebrafish model resembles the progression of the human myopathy resulting from MYO18B deficiency. Our results therefore provide considerable evidence to support the notion that MYO18B dependent myopathy results primarily from defects in sarcomere formation within skeletal muscle and further suggest that zebrafish mutations we have generated are novel models for the study of the human disease.

The function of myo18b is essential for sarcomere assembly

The two currently prominent models of sarcomere assembly diverge regarding the proposed patterning of the Z-bodies. One model proposes premyofibril formation as a first step of myofibrillogenesis (4). During the transition of the premyofibril into mature myofibril, the space in-between Z-bodies widens while premyofibrils align and Z-bodies fuse to mature into Z-disks (3). An alternative model proposes that randomly scattered I-Z-I bodies associate with titin, which subsequently recruits independently formed thick filaments to enable regular spacing of Z-disks (1,2). Analysis of the partial loss-of-function mutant myo18b^sig revealed isolated thin and thick filaments within the sarcoplasm but also scattered Z-disk structures with attached filaments, which resembled I-Z-I bodies. Furthermore, sarcomeric structures with misaligned thin and thick filaments but regularly spaced Z-disks were detected. Independently formed Z-disks associate with thin and thick filaments to form sarcomeric structures with regular Z-disks but misaligned filaments in myo18b^sig homozygotes, indicating that Myo18b is specifically involved with the integration and alignment of preformed thick filaments into sarcomeres. This role of myo18b is also supported by our findings that transgenic Myo18b-GFP localizes into sarcomeres. Interestingly, detection of the randomly scattered I-Z-I bodies within myo18b^sig also indicates that Z-disk formation can occur independent from their assembly into premyofibril. Importantly, early Z-bodies were detected within myo18b-deficient myo18b ^{− 9 + 3} mutants at the 18-somite stage, but Z-bodies were not assembled in regularly spaced premyofibril. This result indicates that alignment of Z-bodies in premyofibril is an essential step during myofibrillogenesis.

Based on the essential role that Myo18b plays during myofibrillogenesis, we propose a novel model of sarcomere assembly where Myo18b coordinates the integration of preformed thick and thin filaments into Z-bodies to enable formation of premyofibril and mature myofibril.

In zebrafish, integration of Myo18b into sarcomeric a-bands is essential for myofibril formation

The expression pattern of myo18b in zebrafish detected by in situ hybridization in heart and skeletal muscle matched the expression of the orthologous genes in human and mice (13,14). However, the reported subcellular localization of MYO18B is currently subject to controversy. Overexpression studies of myc-tagged MYO18B in primary cultures of rat cardiomyocytes demonstrated the integration of MYO18B into A-bands (13). Another study performed with two independent antibodies against MYO18B reported localization of MYO18B within the sarcomeric I-band adjacent to the A-band within skeletal muscle (14). In addition, transgenic MYO18B has also been reported to localize to nuclei in undifferentiated C2C12 myoblasts and endogenous MYO18B was detected by immunohistochemistry in the nuclei of rat cardiomyocytes maintained in confluent cultures (13). Localization to nuclei, however, could not be confirmed by other mouse studies (14). To analyze the localization of zebrafish Myo18b, we expressed GFP-tagged Myo18b in the skeletal muscle of live fish, which revealed the integration of Myo18b-GFP into the sarcomeric A-bands. Importantly, forced expression of Myo18b-GFP was able to restore sarcomere formation in myo18b ^{− 9 + 3} homozygotes that are otherwise devoid of any sarcomeric structures, indicating that transgenic Myo18b-GFP is functional in skeletal myofibers in vivo. In contrast a nuclear localization of Myo18b-GFP within myofibres was not detected.

Zebrafish myo18b mutants feature aspects of myopathies associated with human MYO18B

As discussed above, analysis of myo18b expression revealed that it is mainly expressed in the skeletal musculature of zebrafish, resembling the expression of myosin XVIIIB in humans and mice. Both zebrafish mutants for myo18b are characterized by a reduction in muscle integrity that results in a reduction of maximal force generated by the mutants. In addition, further analyses revealed that Myo18b constitutes part of the sarcomeres and that a reduction or loss of myo18b function results in loss of sarcomere organization. Thus zebrafish myo18b mutants reflect the myopathy pathology evident in human patients carrying mutations in MYO18B. Similarly, mutations in myosin XVIIIB in mammals and zebrafish result in heart defects (14,18). Also the characteristic facies associated with MYO18B in humans is an aspect of the phenotype detected within myo18b-deficient zebrafish. However, we did not detect any malformations of the spinal cord in either of the myo18b mutants possibly due to the early lethality these mutants display. However, the lack of expression of this gene in spinal cord or skeletal tissue suggests that such phenotypes, were they to occur, are unlikely to be the result of a cell autonomous requirement for Myo18b function in these tissues. Therefore, both zebrafish models do not provide further insights into the spinal cord defects reported in human patients exhibiting Klippel-Feil anomaly. Similarly, analysis of myo18b expression did not reveal a marked expression within the central nervous system of developing zebrafish and, accordingly, obvious neuronal defects were not detected by analysis of the tectum utilizing antibodies against acetylated α-tubulin (data not shown). Thus, the zebrafish myo18b mutants do not provide further contributions to the discussion of mathematical disabilities caused by MYO18B in humans.

Interestingly, zebrafish myo18b is not expressed in embryonic slow muscles and accordingly the sarcomere defects that characterize fast muscle are not found within slow muscle of myo18b mutants. In contrast to zebrafish, that feature compartmentalization of the fast and slow muscle, mammalian musculature is a complex and varied mixture of fiber types within individual muscle. The clear distinction of fast and slow muscle within the myo18b zebrafish model might therefore be a valuable tool to study the function of myo18b within a fully penetrant defect within fast muscle that may not be evident in mixed fiber types that might be characterized by various levels of myo18b function within mammalian muscle.

Overall, our findings suggest that Myo18b is a novel sarcomere assembly factor essential for filament alignment during sarcomere biogenesis. Accordingly, the zebrafish myo18b mutants are characterized by a severe sarcomere defect and heart edema, which reflects symptoms found in human patients carrying mutations within MYO18b. Therefore the establishment of zebrafish myo18b mutants might serve as valuable tools to gain further insights into human pathologies associated with deficiencies in MYO18B function.

Materials and Methods

F3 mutant screen

For the F3 generation screen, 48 adult male zebrafish in TU background were treated with ENU as approved by the Monash Animal Service (MAS/2009/05) and described previously (21). Mutagenized males were subsequently out-crossed over two generations to establish F2 families, which were in-crossed and their 3 dpf-old offspring screened for a reduction in birefringence. One of the mutants identified, named schläfrig, was outcrossed over 11 generations to remove background mutations prior to phenotype analysis. All animal breeding was approved by MAS/2009/02BC.

Birefringence analysis

Birefringence is a light scattering effect provoked by the pseudo-crystalline structure of the myofibril that enables visualization of the larvae’s muscle under polarized light conditions. Because the birefringence effect depends on the angle of the larvae and the polarizing filter arrangement, the automated Abrio LS2.2 polarizing microscope (Cri, USA) was utilized to take unbiased pictures of larvae following established methods (20). Images were subsequently subject to densitometric analysis via the Fiji software to measure the average grey values of the first 20 somites. Obtained values were normalized against control siblings that were set to 100%. Three independent clutches of six larvae per genotype were analyzed for their muscle birefringence. Data are represented as means ± SEM and statistical significance was determined by Student’s t-test.

Linkage analysis of schläfrig

To establish the mapping cross, sig mutants in the TU background were out-crossed to WIK. Offspring of one F1 mapping cross was phenotyped by birefringence analysis. Twenty-five larvae per genotype were pooled and their genomic DNA was sequenced via an Illumina HiSeq 100-bp paired-end sequencer (Illumina, USA). Subsequent linkage analysis was performed using the software SNPtrack and the region of homozygosity analyzed in IGV (23).

Cloning and transcript analysis of myo18b

The predicted transcript sequences ENSDART00000122824, ENSDART00000124853 and ENSDART00000129974 were used to clone zebrafish myo18b by RT-PCR. 5- and 3-prime ends of the myo18b transcript were cloned using the SMARTer RACE 5’/3’ kit (Takara, Japan). RT-PCR amplicons were sequence verified and the full coding sequence of myo18b was cloned into pCR-2.1-topo (Invitrogen, USA) by recombination using an In-Fusion HD Cloning kit (Takara, Japan), resulting in pmyo18bCDS. The cDNA of myo18b was submitted as accession number KX485371. The coding sequence of myo18b was cloned downstream of the unc-45b promoter in pcryGFP-600uncGFP to clone the expression plasmid pcryGFP_Umyo18bGFP, which was subsequently used for transient expression of GFP-tagged Myo18b by microinjection (26). To identify the altered myo18b^sig transcript, total RNA was extracted from 3 dpf-old larvae with TriReagent (Sigma) and cDNA was prepared with the SuperScript III First-Strand-Synthesis System (Invitrogen) and myo18b_13R (5’-atgtgtcagagcctgggtca-3’). Subsequent PCR was performed with myo18b_13F (5’-ttgttgagtggttggctggt-3’) and myo18b_12R (5’- cctcctctgcttcctgtcca-3’) and amplicons were sequence identified. Subsequent densitometric analysis was performed on images of three independently performed RT-PCRs as described earlier (21).

Generation and genotyping of myo18b ^{− 9 + 3} mutants

Mutations in myo18b were generated using the CRISPR/Cas9 system following established methods (31). In short, the CRISPR target region within myo18b was selected with the web-based ZiFiT Targeter software and the designed oligonucleotides myoCRISPR_F (5’-taggcacgccggagctccccga-3’) and myoCRISPR_R (5’- aaactcggggagctccggcgtg-3’) were cloned into pDR274 linearized with BsaI (31). The resulting plasmid was transcribed in vitro using the Maxiscript T7 kit (Invitrogen, USA) to generate the sgRNA, which was injected together with Cas9 (NEB, England) into fertilized WT eggs.

To genotype myo18b ^{− 9 + 3} mutants, DNA was isolated from clipped fins or whole embryos and used in a PCR reaction with the oligonucleotides myo18b_F (5’-acctgtctcagccactcagc-3’) and myo18b_R (5’-aaagtgggcgggcactaaca-3’). Whereas SacI digests the resulting 210-bp WT amplicon into 162- and 48-bp, the myo18b ^{− 9 + 3} allele produced a 204-bp amplicon that is not cut.

In situ hybridization, histology, TEM and immunohistochemistry

In situ hybridization, immunohistochemistry as well as Alcian blue and Alizarin red staining was performed according to standard methods. 10µm cryofrozen sections of 3 dpf old larvae were stained with H&E or subject to immunohistochemistry as described (22). Unless described otherwise, all experiments were performed with a minimum of three independent biological replicates. Primary antibodies were used against α-Actinin (1:1000, A7811, Sigma), myosin (A4.1025, DSHB, USA), F59 (1:20, DSHB, USA) and dystrophin (1:20, Mandra1, DSHB). For electron microscopy, 3-dpf-old larvae were fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate over night at 4 °C. Electron micrographs of ultrathin sections were taken on a Hitachi H7500 transmission electron microscope (Hitachi, Japan). Fluorescence images were recorded using a Zeiss LSM 510 Meta fluorescence confocal microscope (Zeiss, Germany).

Muscle force measurements

5 dpf whole larval trunk muscles were prepared and mounted horizontally using aluminum clips onto a small intact muscle test apparatus (model 801C-1900, force transducer model 403A, Aurora Scientific, Ontario, Canada). The muscle preparations were immersed in MOPS-buffered physiological solution, and excited with single-twitch stimulation. Between contractions, the length was stepwise increased by external stretch to a length that generates maximal active force (27). N-benzyl-p-toluene sulphonamide (BTS, Tocris Bioscience, UK) was dissolved in DMSO at the concentration of 50 mM. To inhibit fast-fiber contraction larvae were pretreated in fish water containing 50 μM BTS for 1 h. Subsequently, active force was measured in the presence of 50 μM BTS in physiological solution using a single-twitch stimulation. All force measurements were performed at room temperature.

Supplementary Material

Supplementary Material is available at HMG online.

Acknowledgements

We thank Monash Micro Imaging and the Monash Ramaciotti Centre for Cryo Electron Microscopy for provision of instrumentation and technical support.

Conflict of Interest statement. None declared.

Funding

This work was supported by the National Health and Medical Research Council [APP1084944 to P.D.C. and J.B.; APP1041885 to P.D.C.]. The Australian Regenerative Medicine Institute is supported by grants from the State Government of Victoria and the Australian Government.