Introduction

The calcium ion (Ca²⁺) concentration gradient is known to be a crucial second messenger signal in all eukaryotic cells. The calcium release channels encoded by ryanodine receptor genes are essential to maintaining correct Ca²⁺ dynamics across biological membranes. RYR proteins are cellular sites of interactions with ryanodine, an alkaloid isolated from the stem wood of the plant Ryania speciosa (Jenden and Fairhurst, 1969) that was used for several decades as an insecticide. Because of its muscle-paralyzing effect in humans, ryanodine has been replaced by insect-specific derivatives that are non-toxic for mammals. Analysis of the RYR multi-domain structures and phylogenetic relations among different taxa yields a model suggesting that it evolved from inositol 1,4,5-trisphosphate receptor (IP₃R)-like ancestral Ca²⁺ release channels. The RYR activity is strictly controlled and may respond to the presence of ions (Ca²⁺, Mg²⁺, and Zn²⁺), proteins (calmodulin (Cam) and FK-506 binding protein (FKBP12/12.6)), and small molecules such as ATP, caffeine, and ryanodine. The domain structure is highly conserved. For example, the SPIa kinase and ryanodine receptor (SPRY) domain shares high sequence identity between vertebrates and invertebrates and is engaged in protein-protein interactions with several protein families (Hadiatullah et al., 2022).

The number of RYR copies ranges across taxa. Mammalian genomes carry three RYR genes. For example, in humans, RYR1, RYR2 and RYR3 are located on chromosomes 19q13.2, 1q43 and 15q13.3-14, respectively. Non-mammalian vertebrates, such as Xenopus laevis and chicken, have two RYR copies (Ottini et al., 1996) while in a zebrafish genome, there are five genes: RYR1a, RYR1b, RYR2a, RYR2b, and RYR3 displaying high similarity to other vertebrate RYR genes (Wu et al., 2011). The greater number of RYR genes in zebrafish than in mammals and birds is thought to result from teleost-specific whole-genome duplication (Howe et al., 2013; Postlethwait et al., 1998). In the invertebrates Drosophila melanogaster, Caenorhabditis elegans, live scallop (Placopecten magellanicus), and lobster (Homarus americanus), a single RYR gene has been identified (Hasan and Rosbash, 1992; Maryon et al., 1996; Murayama & Kurebayashi, 2011; Quinn et al., 1998, Xu et al., 2000). However, through alternative splicing, the single invertebrate RYR gene produces several isoforms, thus increasing the diversity of the available protein pool (De Mandal et al., 2019; George et al., 2007).

It is well known that RYR genes are expressed in muscle cells and play a crucial role in muscle contraction, which results from excitation-contraction (E-C) coupling, a series of events involving the conversion of electrical stimulus to Ca-dependent mechanical response. However, RYRs are also expressed in many other animal tissues including the central nervous system (Klatt Shaw et al., 2018; Q. Liu et al., 2005) and are involved in housekeeping functions in the cells of the adult organism and in developmental processes (Fill & Copello, 2002).

RYR1 mutations underlie an array of diseases including muscle-impairing central core disease (CCD), one of the most frequent congenital myopathies, and malignant hyperthermia susceptibility (MHS), characterized by severe reaction to anesthetics, excessive heat or exercise, which can be fatal. In addition, several RYR2 mutations have also been implicated in cardiopathic catecholaminergic polymorphic ventricular tachycardia (CPVT) and arrhythmogenic right ventricular dysplasia of type 2 (ARVD2) (Lanner, 2012). Of the numerous mutations identified to date in RYR genes, most are missense mutations with single amino acid substitutions. However, several deletions, duplications and frame shift mutations have also been identified (Lanner, 2012).

Here, we analyze the phylogenetic origin, expression and function of the Drosophila dRyR gene. Our data show that dRyR is not only required for contractile properties of differentiated somatic and cardiac Drosophila muscle but also influences muscle structure and plays an instructive role in muscle development. Structural muscle changes in a dRyR loss-of-function context suggest its implication in age-associated muscle decline, while severe developmental muscle defects observed in dRyR mutant embryos provide insights into early-onset RYR-related myopathies. Considering extensive structural and functional dRyR conservation, we generated a Drosophila model of one undiagnosed human RYR1 variant mutation (p.M4881Ile) and found that it negatively impacted muscle structure and function, making it likely pathogenic. We report a detailed functional analysis of dRyR. Our findings pinpoint the under-investigated role of dRyR in embryonic muscle development and demonstrate that Drosophila could be used for assessing impacts of human RYR1 variant mutations of unknown significance.

Results

Drosophila dRyR belongs to the RYR gene family and shows conserved somatic and cardiac muscle-associated expression and function

dRyR is the sole RYR gene family member in Drosophila. We applied the maximum likelihood method and the Kimura 2-parameter model (Kimura, 1980) to infer the evolutionary history of dRyR. Based on evolutionary distance studies, RYR2 is the most ancient of three human RYR genes (Ding et al., 2017) and shows the highest sequence homology with Drosophila dRyR. We compared sequences of RYR2 from selected mammalian and non-mammalian vertebrates with the sequence of single invertebrate RYR, including fruit fly dRyR to illustrate its phylogenetic origin (Fig. 1A) (Ding et al., 2017; Mackrill, 2012; McKay & Griswold, 2014; Takeshima et al., 1994). Evolutionary analyses were conducted in MEGA X (Kumar et al., 2018; Stecher et al., 2020). We observed that single invertebrate RYR genes, here from Drosophila and C. elegans, clustered into a separate branch connected to the vertebrate RYR2 branch that clustered 100% (Fig. 1A). The vertebrate RYR2 branch was then subdivided clonally from Xenopus through zebrafish, chick and mouse to human. All clonal branches in vertebrates clustered 100% (Fig. 1A), indicating that RYR2 genes are closely related and most probably evolved from a single invertebrate RYR.

Figure 1.

Expression of RYR genes in vertebrates has been extensively studied and described in various excitable cells including skeletal and cardiac muscles and neurons, and in non-excitable cells such as pancreatic beta cells and lymphocytes (Rossi & Sorrentino, 2002). By contrast, in spite of early works by Hasan and Rosbash (1992) and Sullivan et al., (2000) no systematic analyses have yet been performed to assess the developmental expression pattern of the sole Drosophila dRyR gene.

We first tested the expression of dRYR at protein level. In differentiated body wall muscle of 3^rd instar larva (Fig. 1B,C) dRyR was detected in a discrete striated pattern (Fig. 1B), which in a zoom view revealed highly ordered punctate dRyR protein localization in close vicinity to discs large (Dlg)-positive T-tubules (Fig. 1C and scheme in Fig. 1D). Thus, in Drosophila muscle, like in vertebrates, dRyR localizes at the T-tubule interface, a sub-cellular localization consistent with its calcium release role at SR and E-C coupling function during muscle contraction.

We also assessed muscle-associated expression of dRyR transcripts (Fig. S1). Of ten dRyR transcript isoforms, all coded for proteins of similar amino acid (aa) length, ranging from 5113 to 5134 aa and molecular weight about 580 kDa (http://flybase.org/reports/FBgn0011286.htm). To test dRyR isoform expression, we applied the FISH-HCR technique (Choi et al., 2016) and four probes targeting alternative exons 10, 11, 22 and 23 (Fig. S1A). In this setup we detected the expression of the A,B,F,G,H,J isoforms with the dRyR Ex10 probe, A,B,C,D,E,I isoforms with the Ex11 probe, A,B,C,D,E,F,G isoforms with the Ex22 probe and H,I,J isoforms with the Ex23 probe (Fig. S1A).

FISH-HCR experiments performed on differentiated 3^rd instar larval muscles revealed muscle-associated specific signals with all four probes (Fig. S1B-E’). dRyR transcripts were detected at the periphery of nuclei, in the sarcoplasm and in a repeated striated pattern following sarcomeres. Similar signals observed with the four probes indicated that at least the isoforms A and B that are commonly targeted by the Ex10, Ex11 and Ex22 probes and one of the isoforms H,I or J targeted by the Ex23 probe were present in the functional body wall muscle. We could not, however, rule out the possibility that most or even all dRyR isoforms were expressed in the differentiated body wall muscles. Supporting this, we note that nine out of ten dRyR isoforms (isoform I being an exception) share the same transcription start site, suggesting that a common core promoter regulates their expression.

Detected muscle-associated expression of dRyR prompted us to test its involvement in muscle contraction and larva mobility. We observed that muscle-targeted attenuation of dRyR expression resulted in reduced muscle performance with a significantly longer time required for larvae to switch from the dorsal to the ventral position (Fig. 1E – righting test) and a reduced number of muscle contraction waves compared to control recorded in 1 minute (Fig. 1F – motility test).

Finally, dRyR-RNAi larvae crawled inefficiently and were unable to move over a longer distance (Fig. 1G – locomotor test). In parallel, overexpressing dRyR in larval muscles also impaired muscle function, with fewer contraction waves in 1 minute compared to control and a slower locomotion (Fig. 1F,G), while the time required to switch from the dorsal to the ventral position remained unchanged (Fig. 1E).

Altogether, our data extend previous observations of affected muscle contractility in RyR mutants (Sullivan et al., 2000) and suggest that Drosophila dRyR, like its vertebrate counterparts, ensures correct muscle function, likely acting as a sarcoplasmic reticulum (SR) calcium release channel essential for muscle contraction and E-C coupling.

Figure S1.

Given that vertebrate RYR genes (RYR2 in humans) also play an instrumental role in cardiac muscle function, we sought to determine whether dRyR protein could be detected in the fly heart and whether it could influence heartbeat variables. In the adult Drosophila heart, dRyR protein was detected predominantly in the circular muscle fibers ensuring cardiac contractions (Fig. 2A-A’’’), which appear structurally affected (Fig. 2C, E) in a heart-specific dRyR RNAi context (Hand>dRyR RNAi). Consistent with this, heart-targeted attenuation of dRyR resulted in an abnormal M-mode heart profile (Fig. 2B,D) associated with a longer heart period (Fig. 2F) and a slow heart rate (Fig. 2G), with significantly increased diastolic interval (Fig. 2H). Attenuation of cardiac dRyR expression also led to arrhythmic heartbeat (Fig. 2J), increased systolic diameter (Fig. 2K) and reduced fractional shortening (Fig. 2M). As the diastolic diameter remained unchanged, we conclude that cardiac dRyR knockdown affects cardiac performance without causing dilated cardiomyopathy. In contrast to dRyR-RNAi, increasing dRyR cardiac expression had only a minor influence on cardiac variables (Fig. 2F-M) with increased systolic diameter (Fig. 2K) but no effect on cardiac contractility (Fig. 2M). Overall, observed dRyR loss-of-function adult fly heart phenotypes with a slow heart rate and increased arrhythmia correlate with impaired cardiac function in RyR mutant larvae (Sullivan et al., 2000). We hypothesize that dRyR RNAi-induced impairment of Ca2+ homeostasis could contribute to cardiac aging, for which Drosophila is a recognized model (Nishimura et al., 2011).

Figure 2.

In view of the reduced motility of 3^rd instar larva with the attenuated dRyR (Fig. 1E-G) we examined whether structural properties of body wall muscles were adversely affected. We first found that muscle-targeted attenuation of dRyR (C57>dRyR RNAi) led to a significantly reduced larva body length (Fig. 3B, Q) compared to control (Fig. 3A, Q, an observation that correlates with previously observed (Sullivan et al., 2000) reduced body size of dRyR¹⁶ mutant larvae. Though to a lesser extent, the overexpression of dRyR in body wall muscles also impacted larva length (Fig. 3D, Q). These changes in larva size in loss and gain of dRyR function correlated with a reduced longitudinal muscle length (Fig. 3 E-H, R), which in turn correlated with shortening of Kettin/D-Titin-labelled sarcomeres (Fig.3 I-L, T) and reduced number of myonuclei (Fig.3 E-H, S). Because RYR-mediated calcium homeostasis involves dynamic interactions between the sites of calcium release from the SR and calcium uptake by the mitochondria (Li et al., 2025) we examined whether in Drosophila dRyR loss and/or dRyR gain of function could adversely affect mitochondria pattern in the larval muscles. We noted that the I band-associated striated mitochondria pattern was lost in the C57>dRyR RNAi context (Fig. S2 B) and appeared irregular when dRyR was overexpressed in muscles (Fig. S2 D) compared to the wild-type larval muscles (Fig. S2A, B, E).

Figure 3.

Both muscle-targeted attenuation and gain of dRyR function led not only to impaired muscle functions but also to overall reduction of muscle size and myofibrillar disarray associated with a downsizing of sarcomeres and mitochondrial mismatch. Altered structural muscle features observed in dRyR-attenuated Drosophila larvae are reminiscent of myofibrillar and mitochondrial pattern defects reported in mice harboring a pathogenic RYR1 mutation (Elbaz et al., 2019).

Figure S2.

dRyR is expressed during embryonic muscle development and is required for correct myogenic differentiation

Previous reports provide evidence that RYR-dependent elevation of intracellular calcium promotes late steps of myogenic differentiation and in particular fusion of myoblasts to myotubes (Eigler et al., 2021; Sinha et al., 2022). In parallel, a recent study on RYR1-depleted primary myoblasts revealed the calcium-independent inhibitory role of RYR1 in myogenic differentiation (Tourel et al., 2025). To further explore myogenic roles of RYRs, we tested dRyR expression and function during embryonic development.

Sullivan et al. (2000) reported embryonic dRyR transcript expression in body wall and visceral muscle precursors. Here, we tested dRyR protein expression and found that it was prominently expressed in the mesodermal derivatives in embryos. We detected dRyR protein in the developing visceral, somatic and cardiac muscle cells (Fig. 4). Regarding body wall muscles, dRyR could be detected in the somatic muscle precursors starting from embryonic stage 12 (Fig. 4A, A’), accumulated in the growing myotubes at mid-stage embryos (Fig. 4B, B’) and continued to be expressed in the developing muscle fibers at later embryonic stages (Fig. 4C-D’). At embryonic stage 16, dRyR protein was distributed in a discrete granular pattern within the cytoplasm of myofibers and appeared excluded from the myonuclei (Fig. 4D,D’). Thus, dRyR protein was detected from the early phase of myogenic differentiation that encompasses specification of muscle founders and first myoblast fusion events and continues during the second phase of fusion and myofiber growth and maturation. We also applied HCR-FISH to test dRyR transcript isoform expression in the developing somatic muscle (Fig. S3). We found that dRyR A, B, F and G isoforms harboring alternative exons 10 and 22 were actively transcribed in the developing muscle, whereas the remaining dRyR isoforms were barely detected (Fig. S3).

Figure 4.

Figure S3.

Previous analyses (Sullivan et al., 2000) showed that muscle contraction was compromised in larvae carrying a hypomorphic dRyR¹⁶ mutant allele. dRyR mutant larvae were also smaller in size and died before the pupation stage. However, whether dRyR embryonic expression has a functional impact on larval muscle development has not yet been assessed. Accordingly, we analyzed embryonic muscle pattern in late-stage dRyR¹⁶ mutant embryos (Fig. 5). We observed a wide range of developmental somatic muscle defects with predominant phenotype of thin myofibers present in 64% of abdominal A2-A5 hemisegments and more severely sphere-shaped (arrowheads in Fig. 5B) or missing myofibers (asterisks in Fig. 5B) observed in 25% of hemisegments. In addition, in 10% of hemisegments with dRyR loss of function led to supernumerary lateral transverse (LT) muscles (arrows in Fig. 5B), a phenotype that could arise from LT muscle splitting (Bertin et al., 2021). Predominant thin/misshaped/missing myofiber phenotype in a dRyR loss-of-function context suggests a pro-myogenic role during development. To further characterize the role of dRyR during myogenesis, we analyzed embryonic LT muscle phenotypes in LT-targeted dRyR attenuation and gain-of-function contexts (Fig. 5C-E). Like in dRyR¹⁶ mutant embryos, dRyR RNAi knockdown in LT muscles resulted in thin and/or misshaped LT muscles (Fig. 5D, H) observed in 88% of hemisegments and in rare cases of LT splitting, found in 8% of hemisegments (Fig. 5D, H). We noted no loss of LT muscles in dRyR RNAi embryos. Consistent with the promyogenic role of dRyR, LT-targeted overexpression of dRyR appears to promote LT splitting phenotype (Fig. 5E, H) found in 18% of hemisegments. Calmodulin (Cam is the major calcium dependent RYR regulator. The Ca2+-bound Cam at high calcium levels acts as RYR inhibitor (Fruen et al., 2003). We thus tested effects of Cam attenuation in LTs and found that it results in a dRyR overexpression-like phenotype (Fig. 5F, H). This suggests that during myogenesis Cam is present mainly in a calcium-bound form that represses dRyR. Another major regulator of calcium homeostasis, the endoplasmic reticulum calcium pump SERCA is required to maintain a high calcium levels in the ER lumen (Suisse & Treissman, 2019). To test its role in myogenesis we analysed Lms>SERCA RNAi embryos. We observe that SERCA-depleted LT muscles display growth defects with predominant thin myofiber phenotype (Fig. 5G, H) also observed in Lms>dRyR RNAi context (Fig. 5D, H). Thus, both dRyR-regulated cytosolic and SERCA-regulated ER lumen calcium levels are required to promote muscle development.

Figure 5.

Interestingly, live imaging (Fig. S4) shows that the sphere-shaped muscle phenotype arises from the impaired LT myotube extension and not from retraction of already extended myotubes. Also myonuclei remained on LT extremities and did not spread along the abnormally thin myofibers. The reduced number of myonuclei (2-4 per dRyR RNAi LT myofiber (Fig. S4B, lower panel) instead of 4-6 in control LTs (Fig. S4A lower panel)) points to the role of dRyR in the second wave of fusion (Eigler et al., 2021). This observation is consistent with the fact that overexpression of dRyR induced the LT split phenotype (Fig. 4E) known to be promoted by an excessive myoblast fusion (Bertin et al., 2021). Because the embryonic LT muscle defects in dRyR-mutant embryos are associated with a reduced calcium signal in LTs (Fig. S4 C, D), we hypothesize that dRyR acts as a promyogenic factor ensuring correct calcium levels in the developing myotubes.

Figure S4.

Assessing the impact of the RYR1 undiagnosed variant mutation in Drosophila

Drosophila dRyR shares 45% aa sequence identity with human RYR1 and RYR2, and all protein domains are conserved, with up to 75% of identity for the most C-terminal RIH domain (Fig. S5A). Thus, although the 3D conformation of the Drosophila dRyR has not yet been established, high sequence and positional conservation of functional domains suggest that the conformation of dRyR protein is similar to that of its vertebrate counterparts. Also, the distribution of pathogenic mutations identified in human RYR1 and RYR2 genes (reviewed by Lanner et al., 2010), clustered in three hot spots, correlates with the positions of conserved domains. As revealed by the identity heat map (Fig. S5B) the hot spot regions of human RYR1 and RYR2 mutations align with the most conserved portions of the Drosophila dRyR indicating suitability of the Drosophila dRyR for modeling human RYR gene mutations and their impact on muscular and cardiac systems.

Figure S5.

Over the last decades, whole genome sequencing has identified large numbers of variant mutations within the RYR genes, most of which are classified as variants of unknown significance (VUS). The recessive c.14643G>A / p.Met4881Ile missense RYR1 mutation was identified in a young patient with a phenotype of congenital myopathy with a delayed acquisition of motor function. At the histopathological level, filamentous aggregates were present in muscle biopsies. This very rare mutation for which a link with a muscle disorder has not yet been evaluated, is located in the RYR1 region encoding calcium pore.

We made use of the conserved muscle function and structural similarity between dRyR and human RYRs (Fig. S5A) demonstrated here to generate a Drosophila model of variant p.Met4881Ile RYR1 and assess its impact on larval muscle function and structure (Fig. 6). The p.Met4881Ile RYR1

Figure 6.

VUS-carrying larvae were homozygous-viable but were significantly smaller (Fig. 6B,I) than control larvae (Fig. 6A,I). They had shorter ventral longitudinal muscles (Fig. 6 D,J), which harbored fewer myonuclei (Fig. 6D,K) and were characterized by shorter sarcomeres (Fig. 6F,H) compared to control (Fig. 6C,E,G). As revealed by the larva motility tests (Fig. 6M, O) the p.Met4881Ile RYR1 VUS impacted not only structural but also contractile muscle properties, leading to a reduced frequency of peristaltic body wall muscle contraction (Fig. 6N), slower larva locomotion (Fig. 60) and compromised muscle performance revealed by the righting test (Fig. 6M).

These structural and functional muscle phenotypes are reminiscent of those observed in dRyR RNAi larvae, suggesting that p.Met4881Ile variant mutation negatively influences dRyR expression and/or function and could likely contribute to RYR-related myopathies in humans.

Discussion

The ryanodine receptors encoding calcium release channels are key regulators of intracellular calcium level, largely conserved among vertebrate and invertebrate species. By their involvement in many calcium-dependent biological processes, RYR genes are vital for normal cellular functions, while mutations in RYR genes underlie a broad spectrum of human diseases including neuromuscular and cardiac disorders (Marks, 2023). Because of their large size (over 500 kDa), structural complexity, many isoforms and broad spectrum of interacting factors, even though their role in promoting muscle contraction via E-C coupling is well-characterized, other RYR functions remain only partially understood. Here, we focus on dRyR, a single Drosophila member of the RYR gene family and characterize its expression and function in differentiated and developing muscle.

Drosophila dRyR shares 45% aa sequence identity with its human counterparts. Considering previous evolutionary distance studies (Ding et al., 2017) we show that the phylogenetic dRyR branch is connected to RYR2, the most ancient of the three human RYRs. Further, aa sequence alignment reveals an extensive conservation of all protein domains between dRyR and human RYRs, while a heatmap of conserved regions highlights previously identified hot spots of pathogenic RYR1 and RYR2 mutations. Remarkably, several aa residues at the functionally relevant channel pore domain (Wang et al., 2012) are conserved between Drosophila and humans.

Consistent with potential conservation of dRyR function in triggering Ca²⁺ dynamics across ER membranes, we detected ER-associated punctate dRyR protein expression in differentiated body wall muscles. Our data extend previously reported analyses (Sullivan et al., 2000; Vázquez-Martínez et al., 2003) providing evidence that dRyR is not only expressed in visceral muscle and the nervous system but also prominently in differentiated striated somatic and cardiac muscle. We also wanted to know whether different dRyR transcript isoforms were differentially expressed in body wall muscles. However, we did not detect any such differential expression with 4 HCR-FISH probes targeting alternative dRyR exons. The fact that most dRyR transcript isoforms are present in differentiated larval muscles is consistent with their common transcription start sites. However, whether all dRyR isoforms are collectively required for larval muscle function requires further investigation. In line with muscle-associated dRyR expression and the excitation-contraction coupling role of its vertebrate orthologs, RNAi knockdown of dRyR leads to reduced muscle contractility and severely impaired larva mobility. Interestingly, in addition to impaired muscle function, dRyR attenuation causes extensive structural muscle defects including reduced muscle size, smaller and aberrant sarcomeres and degraded mitochondria pattern. All these functional and structural muscle defects are reminiscent of those of aged muscle, indicating that maintenance of dRyR-triggered calcium management could prevent muscle aging. This applies also to cardiac muscle, which in the dRyR RNAi context shows aging-associated slow heart rate and arrhythmia (Blice-Baum et al., 2019).

dRyR transcripts and dRyR protein were also detected in the embryonic muscle precursors from embryonic stage 12 to late-stage embryos, indicating that dRyR could be involved in managing Ca²⁺ levels throughout myogenesis. In contrast to differentiated larval muscles, HCR-FISH experiments show that only a subset of dRyR splice isoforms is expressed in the developing muscles. Our developmental dRyR expression data, focusing on somatic muscle, extend previous more general analyses of dRyR expression and function (Takeshima et al., 1994; Sullivan et al., 2000; Vázquez-Martínez et al., 2003). Consistent with spatiotemporal embryonic dRyR expression and the role of cytoplasmic calcium management (Li et al., 2025), we show that dRyR loss of function and RNAi knockdown in developing muscles cause severe developmental muscle defects. We found that dRyR promoted myogenic differentiation and was required for myotube growth associated with myoblast fusion and followed by myonuclear spreading within the myotubes. Our observations in Drosophila are consistent with the promoting role of RYR1 in the calcium-dependent myoblast-to-myotube fusion process reported by the Avinoam lab in an in vitro myogenic differentiation system (Eigler et al., 2021). Interestingly, our analyses of embryonic muscle phenotypes of hypomorph dRyR¹⁶ embryo reveal that dRyR, in addition to its major promyogenic role, may also negatively influence myogenic differentiation. A negative influence on myogenic differentiation and in particular on myoblast fusion has recently been reported in mouse RYR1 mutant myoblast culture (Tourel et al., 2025). It was suggested that this early developmental role of RYR1 was calcium-independent. dRyR might thus play a dual role in myogenesis: (i) as a calcium-independent negative regulator of first myoblast-to-myoblast fusion events and (ii) as a positive regulator of myogenic differentiation acting in later steps of myogenesis in a calcium-dependent way to promote myoblast-to-myotube fusion and muscle fiber growth. This major pro-myogenic dRyR function is further supported by the dRyR gain-of-function phenotypes.

The knowledge gained on dRyR and its muscle-associated functions prompted us to use Drosophila to test the impact of one human RYR1 mutation with unknown significance. We chose undiagnosed recessive p.Met4881Ile variant mutation identified in a patient with congenital myopathy phenotypes, severely impaired mobility and accumulation of filamentous aggregates in muscle fibers. We found that Drosophila larvae carrying p.Met4881Ile mutation in the dRyR gene showed dRyR RNAi-like phenotypes with impaired larval mobility and significantly impaired sarcomeric muscle structure. In all, we assessed eight different functional and structural muscle variables showing that p.Met4881Ile variant mutation consistently impaired larval muscle performance and changed muscle size and structure. This suggests that the p.Met4881Ile mutation impairs dRyR function and is likely pathogenic.

To conclude, we report functional analysis of dRyR, the sole fruit fly RyR gene and show that in addition to ensuring contractile properties of differentiated striated muscle it plays a key pro-myogenic role during muscle development. Our findings advocate Drosophila for modeling and testing the impact of human RYR1 variant mutations of unknown significance.

Materials & Methods

Drosophila strains and genetics

Fly stocks were maintained at 25°C on standard fly food.

The targeted expression experiments were performed using the UAS-GAL4 system (Brand and Perrimon, 1993) on the following GAL4 and UAS lines: C57-GAL4 (Bl32556); Hand-GAL4 (kindly provided by L. Perrin; TAGC, Aix-Marseille University, France); UAS-dRyR RNAi (BL29445); Lms-GAL4 (BL46861); UAS-mCherry RNAi (BL35785); UAS-dRyR (kindly provided by H. Nash University of Maryland College Park, Rockville, USA), UAS-lifeAct-GFP (BL35544), UAS-dsRed NLS (BL8547; BL8546), UAS-LacZ (BL1776), UAS-Cam RNAi (Bl34609), UAS-SERCA RNAi (BL44581) and UAS-GCaMP3 (BL32236). The RyR¹⁶/CyO Wg LacZ (BL6812) was used as hypomorphic mutant and the w¹¹¹⁸ strain was used as wild type.

Phylogenetic analysis

Initial trees for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood (MCL) approach and then selecting the topology with superior log likelihood value (Kumar et al., 2018). The tree is drawn to scale, with branch lengths measured in number of substitutions per site. This analysis involved seven nucleotide sequences. Codon positions included were 1st+2nd+3rd+Noncoding. There were 17,322 positions in the final dataset.

Immunohistochemistry

Antibody staining was performed using standard protocol. Embryos were fixed in 4% formaldehyde and blocked in NGS serum to remove non–specific epitopes. They were incubated overnight at 4°C with primary antibodies followed by secondary antibodies for 2 h at RT.

3^rd. instar larvae were dissected and fixed in 4% paraformaldehyde for 20 minutes as previously described (Lavergne et al., 2020). The fly hearts were dissected (Fink et al., 2009; Ocorr et al., 2007) and fixed for 15 min in 4% formaldehyde and the immunostaining procedure was performed as described (Auxerre-Plantié et al., 2019).

The following primary antibodies were used in this study: guinea pig anti-dRYR antibody (1:200, kindly provided by Robert Scott and Benjamin White from NIH/NIMH Institute and previously described in Gao et al., 2013), mouse anti-Dlg (1-50, DSHB, 4F3), rat anti-actin (1-500, Abcam, ab 50591), mouse anti-kettin (1:50 DSHB 1B8-3D9), mouse anti-ATP5A (1:200 Abcam, ab 14748), goat anti-GFP (1:500, Abcam, ab 5450), and chicken anti-β galactosidase (1:1000, Abcam, ab 9361). Rhodamine phalloidin (ThermoFischer Scientific) was used to reveal actin filaments in the heart and muscles. Fluorescent secondary antibodies (Jackson ImmunoResearch) were used to detect primary antibodies.

Muscle characteristics measurements

All analyses of muscle length and sarcomere size were performed on fixed larval muscle preparations in a relaxed state. Acquired confocal images were analysed in Fiji using the line tool. Analyze – Measure tool was then applied to obtain muscle length values and measurements were analysed with Prism. Sarcomere size and number were calculated using Analyze – Plot profile Fiji tool. The sarcomere size was measured between peaks corresponding to Z-disc (revealed with Z-line specific marker) on approximatively 100 μm of muscle length. Sarcomere measurements were then analysed with Prism. DAPI-stained nuclei were counted in Z-stacks of confocal views of VL3 larval muscle and data analysed with Prism. About 30 larval muscles from 6-8 larval filets were analysed for each measurement.

In situ hybridization chain reaction – HCR

In this study we used the two-stage in situ HCR protocol described by Choi et al. (2016). This technique detects and amplifies specific transcripts by the direct binding of probes to nucleic target sequences without additional long-lasting enzymatic reaction. We planned four different mRNA probes targeting alternative exons numbered 10,11,22 and 23.

We used a Molecular Instruments HCR kit containing a DNA probe set, a DNA HCR amplifier B1-Alexa fluor 488, B2-fluor 532 and hybridization, wash and amplification buffers.

Fixed samples were pre-hybridized at 65°C for 2 h followed by hybridization steps overnight at 45°C. After several washes, the amplification step was performed overnight at RT in the dark. We used TRJ244 HCR amplifier B1, RTJ245 HCR amplifier B2, RTJ256 HCR amplifier B2, RTJ 247 HCR amplifier B1. Target mRNA sequence information remains at the discretion of Molecular Instruments Company. Excess of hairpins was removed by several washes with 5XSSCT solution at room temperature.

Imaging

Samples were mounted in Vectashield with DAPI (Vector Laboratories, Inc. Burlingame, CA) and a Leica SP8 confocal microscope was used for image acquisition and for time-lapse imaging of living embryos. In vivo imaging of lateral muscle was performed from late stage 14 to stage 16 every 3 min. We used Adobe Photoshop and ImageJ for image processing.

Genome editing

To generate the Drosophila model of human RYR1 VUS p.Met4881Ile we applied the CRISPR-Cas9 homologous recombination genome editing approach with the use of single strand oligo donor (ssODN) and gRNA. A 20nt 5’-AAACGCTTCGTGTTCCATCTGTAC-3’ gRNA targeting the dRYR region close to the mutation site was designed using the OPTIMAL Target Finder platform (Gratz et al., 2014) and cloned into PCFD5 plasmid (Adgene). Single strand ssODN donor of 115 nt in size with sequence modification TTG to ATC was generated by IDT Company. Both components were injected at a concentration of 100 ng/μl by Best Gene into nos-Cas 9 (III-attp2) flies. PCR molecular screening with a pair of primers: Forward 5’-TGCAGAGCAGCCGGAGGATGAC; Reverse 5’-ATCAGACGCGGCGAATCCGCAG and Sanger sequencing were used to identify progenies carrying the edited sequence. Genetic crosses established homozygous Drosophila lines carrying the RyR1 p.Met4881Ile variant mutation.

Functional tests of muscle performance

Motility tests were carried out on 30 3^rd instar larvae as previously described (Picchio et al., 2013). The motility test was performed by recording the number of peristaltic contractions executed by the larva in 1 min on a grape medium plate. The righting test consisted in placing the larva on its back and measuring the time it took to revert to its crawling position. For the locomotor test, a track 2 mm wide, 5 mm deep and 5 cm long was prepared on a grape medium plate. Larvae were placed on the test track and the time they took to crawl a distance of 5 cm was recorded.

Heartbeat analyses

The cardiac activity analyses of adult Drosophila hearts were performed on 1-week-old females flies using the Semi-automated Optical Heartbeat Analysis (SOHA) approach protocol of Fink et al. (2009). For each experiment about 20 flies were analyzed. The flies were anesthetized with Fly Nap, dissected in an oxygenated, artificial hemolymph composed of 108 mM NaCl, 5 mM KCl, 2 mM CaCl2, 8 mM MgCl2, 1 mM NaH2 PO4, 4 mM NaHCO3, 10 mM sucrose, 5 mM trehalose, 5 mM HEPES (pH 7.1). The beating hearts were filmed by digital camera on 30s movie with the speed of 150 frames/s (Digital camera C9300, Hamamatsu, McBain Instruments, Chatsworth, CA). The SOHA program based on Matlab R2009b software was used for film analysis (Fink et al., 2009).

Mitochondria area quantifications

The total area of mitochondria was identified by Otsu thresholding on Fiji for internal muscle z-planes. The quantifications for z-planes were averaged for each of 25-30 different VL3 muscle according to Zhang et al., (2024).

Statistic

All statistical analyses were performed using Prism (v9.5.1, GraphPad, Software, La Jolla, CA, USA). The t test was used to compare control to variant context and one-way ANOVA tests were used for comparisons with more than two datasets. Bar plot represent the mean and the standard deviation. On the figures, statistical comparisons of sample vs control are indicated as ****: P ≤ 0.0001; ***: P ≤ 0.001; **: P ≤ 0.01; *: P ≤ 0.05; ns > 0.05.

Ethical approval

This study was conducted in accordance with the ethical standards of the institutional and national research committees. Ethical approval was obtained from the appropriate institutional ethics committee under the reference 38RC21.0399

Data availability

All behavioral, phenotypical and image analyses datasets that were used to generate graphs in Figures 1, 2, S2, 3, 5 and 6 are available and will be provided as source.

Acknowledgements

We thank Drosophila Bloomington Stock Centers for Drosophila lines and L. Mouty for the technical assistance. This study was supported by the « Priority Research Programme on Rare Diseases » of the French Investments for the Future Programme, by the AFM-Telethon strategic grant to MyoNeurAlp consortium, the Reference Centre for Neuromuscular Diseases grant to MZ and CS, ANR-iSITE-2025 grant to MZ, by the French Government Scholarship grant to MMP and The Polish National Agency for Academic Exchange within The Bekker Programme (grant no. BPN/BEK/2021/2/00006) to MDM.

Additional information

Funding

Institut National de la Santé et de la Recherche Médicale (Inserm) (RYR-ClassifAI)

• John Rendu

• Krzysztof Jagla

Department of Agriculture, Food and the Marine, Ireland (DAFM) (MyoNeuralp)

• Monika Zmojdzian

Agence Nationale de la Recherche (ANR) (iSITE CAP2025)

• Krzysztof Jagla

• Catherine Sarret

Bekker Programme (BPN/BEK/2021/2/00006)

• Magda Dubinska-Magiera

• Marta Migocka-Patrzałek