Drosophila Tropomodulin is required for multiple actin-dependent processes within developing myofibers Free

Skeletal muscle cells (myofibers) are essential for survival. Myofibers are composed of myofibrils, which consist of arrays of repeating units known as sarcomeres, the fundamental contractile machinery of the muscle. These contain three main structural elements: thin filaments, thick filaments and Z-discs [reviewed by Sweeney and Hammers (2018)]. Z-discs are the thin filament attachment site and are composed of many proteins, for example α-Actinin and Zasp (also known as Zasp66). Thin filaments are composed primarily of actin filaments (F-actin) and actin binding proteins, including proteins important for anchorage to the Z-disc (for example CapZ; Cpb), for contraction (troponin and tropomyosin) and for F-actin length regulation (nebulin and Tmod) (reviewed by Szikora et al., 2022; Ghosh and Fowler, 2021; Fowler and Dominguez, 2017). Thick filaments are composed primarily of myosin and myosin-associated proteins. Thin filaments sliding past thick filaments brings Z-discs closer together (contraction), generating force in a calcium-regulated manner. Although much is known about protein composition and function of sarcomeres and myofibrils, the mechanisms controlling their formation and maintenance remain elusive.

Although myofibrillogenesis is not well understood, the most widely accepted model suggests tension is essential. According to this model, once the myofiber has attached to tendon cells, short F-actin stochastically attach to each other and to the myofiber ends via tension-activated proteins such as β-integrin and Talin. Through a positive feedback loop, initial tension attracts more tension-activated proteins, leading to further tension and myofibril assembly and maturation along the longitudinal axis of the myofiber (Lemke and Schnorrer, 2017; Lemke et al., 2019; Pines et al., 2012; Weitkunat et al., 2014; Sparrow and Schöck, 2009). Subsequently, additional myofibrillar growth and organization also require tension (LeGoff and Lecuit, 2015). Several other processes regulate myofibrillogenesis and myofibril maintenance. First, mitochondria influence myofibrillogenesis and vice versa via the mechanical interplay between mitochondrial dynamics and myofibril morphogenesis (Avellaneda et al., 2021). Second, myofibrils undergo protein turnover without compromising their organization during contraction and myofibrillogenesis (Sanger and Sanger, 2008; Ono, 2010; Littlefield and Fowler, 2008). Third, microtubules have been shown to play guidance roles in the establishment of myofibrils in adult flight muscles (Dhanyasi et al., 2021). Finally, actin regulatory proteins that control monomeric (G-actin) to F-actin ratios in the cell are crucial for myofibril assembly and turnover (Littlefield and Fowler, 2008; Deng et al., 2021). Although contributing factors to myofibril formation and maintenance have been identified, their interplay and molecular mechanisms are incompletely understood.

Mutations in sarcomere genes are linked to skeletal muscle diseases such as nemaline myopathy (NM), which presents with muscle weakness (de Winter and Ottenheijm, 2017; Sewry et al., 2019). NM is associated specifically with mutations in thin filament genes, including ACTA1, NEB, LMOD3, CFL2, TPM2, TPM3, TNNT1, TNNT3 and MYPN (reviewed by Laitila and Wallgren-Pettersson, 2021; Christophers et al., 2022). In addition to the growing list of mutations identified in human patients, Tmod mutant zebrafish show an NM-like phenotype (Berger et al., 2014). Studies in model systems have demonstrated that Tmod regulates F-actin length by capping pointed-ends to inhibit monomer addition/dissociation (Molnár et al., 2014; Berger et al., 2022; Littlefield et al., 2001). Tmod is related in sequence and structure to leiomodin (Lmod), an F-actin nucleator (Chereau et al., 2008; reviewed by Fowler and Dominguez, 2017; Yamashiro et al., 2012), deficiency of which has been linked to NM (Yuen et al., 2014; Cenik et al., 2015). Some Tmods also have low nucleation activity (Fischer et al., 2006; Yamashiro et al., 2010, 2014). Although humans have four TMOD and three LMOD genes that cap and nucleate F-actin, respectively, Drosophila has no Lmod, but expresses 18 Tmod splice variants. It is unknown, however, which Tmod isoforms are most abundant in Drosophila muscle and whether individual isoforms cap or nucleate F-actin. It is also unknown whether and how the disruption of Tmod isoforms in Drosophila would result in the NM-like defects observed in other models.

Previous muscle research focused on the role of Tmod and Lmod in regulating sarcomeric F-actin. However, similar to other cell types, cytoplasmic F-actin plays several essential roles in myofibers: maintenance of intracellular structure by tethering myofibrils to the plasma membrane and extracellular matrix (ECM), organization of myofiber organelles and vesicle trafficking, and movement and anchoring of myonuclei at the myofiber periphery (reviewed by Davidson and Cadot, 2021; Charvet et al., 2012; Kee et al., 2009). The disruption of cytoplasmic F-actin dynamics could contribute to muscle deterioration and weakness and has not been comprehensively studied. Here, we analyze the effects of Tmod knockdown (Tmod-KD) in Drosophila larval muscle. We observe progressive muscle deterioration structurally and functionally, with defects in muscle size, sarcomere length, myofibril homeostasis, Integrin-based myofibril attachment, and myonuclear shape and activity. Alongside myofibril organization breakdown, Tmod-KD myofibers show ectopic F-actin accumulations. Transcriptomic analyses reveal downregulation of proteasomal degradation and upregulation of oxidative metabolism. Biochemically, Drosophila Tmod exclusively acts as an actin capping protein which, in addition to regulating thin filament length, is involved in establishing tension along the myofiber. The phenotypes generated by Tmod loss support and extend the tension-driven self-organizing myofibrillogenesis model and suggest a role for Tmod in myofibril assembly and maintenance. We propose that the function of Tmod as an F-actin capping protein is crucial in a variety of locations within the myofiber that directly and indirectly maintain muscle function.

To identify genes with a crucial role in muscle development and disease, we performed a limited screen in Drosophila melanogaster. We used a muscle-specific driver (Dmef2-Gal4) and available RNAi lines to knock down homologues of human proteins implicated in myofibril/sarcomere structure and function (Schnorrer et al., 2010) (Table S1). We screened for embryonic lethality, muscle defects in late third instar larvae, pupal lethality and adult flight (Table S2). Expression of tmodRNAi resulted in phenotypes that suggested Tmod maintains muscle integrity and function. We confirmed the KD phenotypes using three different RNAi lines and a homozygous tmod mutant, and assessed KD levels using qRT-PCR, immunofluorescence and western blot (Fig. S1). Based on these results, we selected tmod for further study and used one RNAi line, dsRNA-HMS02283 (Perkins et al., 2015) for subsequent KD experiments as it exhibited an intermediate phenotype and degree of mRNA KD.

Drosophila larvae are segmented organisms with three thoracic and eight abdominal body segments (A1-A8), each consisting of ∼30 unique body muscles (Bate, 1990). Here, we focused on two ventral longitudinal (VL) muscles, VL3 and VL4 (also named muscles 6 and 7), which are well characterized, easily assessable, flat rectangular cells with nuclei located on one cell surface (Fig. 1A-C) (Windner et al. (2019). Muscle-specific Tmod-KD resulted in a striking phenotype at the end of larval development, with a significant reduction in myofiber size (Fig. 1D,E). Although the myofiber length was conserved, the width was reduced by 30-60% (Fig. 1D-F). Furthermore, we observed several distinct morphological features: increased sarcomere length (Fig. 1G), disorganized misoriented myofibrils (Fig. 1H,I), ectopic F-actin accumulation (Fig. 1D,E) and misshaped internalized nuclei (Fig. 1J). Strikingly, intact myofibrils with changes in sarcomere length were restricted to the cuticular side of the myofiber. In contrast, the visceral surface, which includes the myonuclei, was disorganized (Fig. 1E; control in Fig. S1). These observations supported a crucial function for Tmod in regulating sarcomeric actin and myofibril stability. Furthermore, they suggested additional Tmod role(s) in the regulation of cytoplasmic actin, including nuclear stabilization and other cellular processes.

Drosophila larval development can be divided into three instar stages during which body wall muscles grow dramatically in size (25- to 40-fold): first, second, and third instars (this last stage can be subdivided in early and late third instars, the last 2 h of which is called the ‘wandering’ stage). Over this 4 day period, new sarcomeres and myofibrils are added, while nuclear number remains constant (Demontis and Perrimon, 2009; Bai et al., 2007; Balakrishnan et al., 2020). Although muscle phenotypes were found in all Tmod-KD late third instars (see also Fig. S1B), not all segments were affected equally. Instead, we found a gradient of muscle phenotypes along the larval anterior-posterior (AP) axis. Myofibers in more anterior segments (A1, A2) had severely disrupted myofibrils and nuclei, whereas posterior ones (A5, A6) maintained nuclear shape and position and exhibited a milder myofibrillar phenotype (Fig. S2).

We analyzed Tmod-KD phenotypes along the larval AP axis throughout larval development. In first instars, we observed wildtype (WT)-like muscles, which contained the typical conserved arrays of sarcomeres. The first observed defects in the Tmod-KD myofibers were diagonal myofibrils at the myofiber ends of segments A1 and A2 in second instars (Fig. 2A; Fig. S2). In early third instars, these defects also appeared in mid-segments (A4, A5) and became even more pronounced in the anterior segments, which displayed ectopic diagonally intersecting myofibrils (Fig. 2B,D; Fig. S2). In late third instars, the sarcomeric pattern was disrupted almost entirely on the visceral side of anterior segments and large bundles of F-actin replaced organized myofibrils (Fig. 2C,F; Fig. S2). Occasionally, interlaced ‘braided’ myofibrils formed at the beginning of late third instar, and ectopic radial F-actin accumulations arose by the end of late third instar (Fig. 2E,G). In addition, once the first diagonally misoriented myofibrils were observed, perpendicular myofibrils were found adjacent to the myotendinous junction (MTJ) at the posterior myofiber end (Fig. 1H; Fig. 2B). These perpendicular myofibrils also later developed at the anterior end of the myofibers (Fig. 2C). Nuclear irregularities appeared in anterior segments during late third instar stage and, together with ectopic F-actin structures, were only observed in anterior segments. These data indicated that Tmod-KD myofibers experience a specific temporal and spatial pattern of structural changes, which appeared in parallel with larval growth, with the most severe disruptions occurring in dramatically growing third instar larvae.

To understand how Tmod loss generates these phenotypes, we studied the biochemical activity of Drosophila Tmod using purified proteins in vitro. The canonical role of Tmod in vertebrates is filament pointed-end capping (Dye et al., 1998; Weber et al., 1994; Gregorio et al., 1995; Mardahl-Dumesnil and Fowler, 2001; Fowler and Dominguez, 2017). However, some Tmods also have limited actin filament nucleation activity (Fischer et al., 2006; Yamashiro et al., 2010, 2014), mimicking the activity of the Tmod structural homolog Lmod that has strong nucleation activity (Chereau et al., 2008). Lmod is not found in Drosophila. Nonetheless, while mammals express only four Tmod isoforms, Drosophila expresses 18 different splice variants. Moreover, several Drosophila Tmod isoforms differ substantially in sequence from their mammalian counterparts, displaying N- and/or C-terminal extensions reminiscent of those present in mammalian Lmods (Fig. S3F). These observations led us to ask whether Drosophila Tmod isoforms capped pointed-ends or nucleated filaments.

We chose two isoforms for in vitro analysis, the selection of which was based on several criteria: the RNAi target sites, transcript abundance in control muscle, the degree of KD, sequence analysis and the unique structures of the isoforms (Fig. S3A-F, Table S3). Based on these factors, isoforms Q, O and K, representing two different N-terminal subgroups, were chosen for in-depth biochemical analysis (note that isoforms O and K have identical protein sequences). We expressed and purified TmodQ, TmodO/K, and human TMOD3 (a control for capping activity) in Escherichia coli and monitored the effect of TmodQ and TmodO/K on actin assembly using the pyrene-actin polymerization assay (Doolittle et al., 2013). Because the concentration of actin monomers (1.5 µM) is higher than the critical concentration for monomer addition at the pointed-end (∼0.6 µM), pointed-end capping under these conditions would be observed as a decrease in fluorescence with increasing Tmod concentrations, resulting from inhibition of monomer addition at the pointed-end. In contrast, if Tmod nucleated filaments from monomers, we would observe an increase in fluorescence (i.e. polymerization), resulting from the formation and elongation of new barbed-ends. Similar to human TMOD3, TmodQ and TmodO/K reduced polymerization in a concentration-dependent manner (Fig. 3A; Fig. S3G,H). These results suggested that the two representative Drosophila Tmod isoforms analyzed function as bona fide pointed-end cappers like their mammalian Tmods, and not nucleators like mammalian Lmods. Therefore, the observed KD phenotypes are likely due to a deficiency in pointed-end capping.

Previous work has shown that Tmod regulates sarcomere length by capping the pointed-end of sarcomeric F-actin (reviewed by Littlefield and Fowler, 2008; Ghosh and Fowler, 2021). To assess the effects of Tmod-KD in Drosophila larval myofibers, we quantified muscle length, Z-disc number and sarcomere lengths in second and late third instars. At both time points, Tmod-KD myofibers had fewer Z-discs than controls with the same myofiber length (Fig. 3C; Fig. S4A,B). Z-disc numbers were reduced by 15% in the second instar and by 26% in late third instar, indicating that sarcomere size increased in growing Tmod-KD myofibers. Accordingly, the distance between Z-discs revealed a significant 22% and 31% increase in sarcomere length in Tmod-KD myofibers at the second and late third instar stages, respectively (Fig. 3D). To confirm that Tmod regulates sarcomere size via its effect on thin filaments, we quantified sarcomeric F-actin length in both second instar and late third instars. Thin filament length increased by 7% in second instar and 35% in late third instars in Tmod-KD myofibers. Interestingly, the proportion of thin filament length to sarcomere length was similar to control at each time point, whereby thin filaments account for ∼24-27% of sarcomere length (Fig. 3B,F; Fig. S4). Together, these data suggested that capping of sarcomeric F-actin by Tmod is required to prevent increases in sarcomere size during myofiber growth. The same thin filament to sarcomere length ratio in Tmod-KD and control myofibers indicates that independent mechanisms maintain sarcomere proportions.

Sarcomere size is highly conserved for optimal contraction, and changes in its size have been linked to reduced muscle function (Fernandes and Schöck, 2014; Molnár et al., 2014; Spletter et al., 2018, 2015). To assess the functional consequences of muscle-specific Tmod-KD, we performed larval locomotion assays. We analyzed larval velocity throughout larval development, and myofiber function significantly decreased, starting at second instar stage and over larval development (Fig. 3E). These data suggested Tmod-KD caused myofiber weakness even before obvious muscle deterioration, and that changes in thin filament length could be the first effect of Tmod-KD to reduce muscle function.

The current model of myofibrillogenesis suggests that tension is crucial for this process. Specifically, the muscle attachment sites at the MTJs play a fundamental role in generating tension within a myofiber. The MTJ represents an analogous structure to focal adhesion sites (Atherton et al., 2016; Martino et al., 2018; Pardo et al., 1983; Quach and Rando, 2006; Mohammad et al., 2012), which are composed of the Vinculin-Talin-Integrin complex tethering the actin cytoskeleton to the ECM, or in the case of the myofiber, the myofibrils to the muscle attachment site. We hypothesized that Tmod functions, via its role as an actin filament capping protein, at the MTJs to stabilize myofibril attachment during myofiber growth.

In agreement with this hypothesis, we detected Tmod localized at the ends of control myofibers, adjacent to myofibrillar F-actin and co-localizing with β_PS-integrin (Mys) (Fig. 4A-C; Fig. S5G). We next evaluated protein enrichment of the tension-mediating proteins β_PS-integrin and its binding partner Talin (Rhea) in Tmod-KD MTJs. β_PS-integrin and Talin were significantly decreased in Tmod-KD anterior segments (Fig. 4D-G; Fig. S5A-C′). To determine whether the strong decrease in signal was due to mislocalization or an overall decrease in protein levels, we performed western blot analysis. Talin levels did not decrease in Tmod-KD wandering larvae muscle-enriched lysates (Fig. 4I; Fig. S5E), suggesting that Talin is mislocalized. Upon further examination, we observed pools of β_PS-integrin and Talin in ectopic areas (Fig. 4E). These data suggested that the Tmod F-actin capping function is required for establishing the proper anchorage sites at the myofiber ends by stabilizing the interaction between F-actin and tension-mediating proteins.

To address whether mislocalization of β_PS-integrin and Talin correlated with tension loss at the MTJ, we assessed phospho-tyrosine (pTyr), a marker of active cytoskeletal tension sites (Chrzanowska-Wodnicka and Burridge, 1994; Martino et al., 2018; Parsons et al., 2010) that has been shown to be enhanced under tension in several contexts (Sawada et al., 2006; Tamada et al., 2004; Yu and Zallen, 2020; Bays et al., 2014). pTyr was significantly decreased in Tmod-KD MTJs (Fig. 4E,H; Fig. S5D,D′), indicating reduced tension at the MTJ. Together, these experiments suggested a decrease in mechanosensory proteins and a reduction of tension at the muscle ends. A lack of tension could cause myofibrillar disorganization and impaired myofibrillogenesis in the Tmod-KD myofibers.

Control Drosophila larvae displayed multiple, evenly spaced, round disc-shaped nuclei on the visceral side of the myofiber. In contrast, many Tmod-KD nuclei were elongated, with an increased perimeter compared with control (Fig. 5A-C). The most irregular-shaped nuclei were located towards the middle of the myofiber along the AP axis (Fig. 5D). It is possible Tmod-KD directly affected F-actin dynamics that stabilize nuclear shape and position. Unfortunately, imaging of cytoplasmic and/or nuclear F-actin is obstructed by the bright signal of the underlying sarcomeres, which prevented the further analysis of direct effects of Tmod-KD on cytoskeleton and nucleoskeleton in this study.

Stable radial microtubule (MT) arrays around the nuclei maintain positioning and nuclear integrity during myofiber contraction (Zheng et al., 2020; Becker et al., 2020; Manhart et al., 2018; Metzger et al., 2012). In addition, MTs physically interact with actin (Farina et al., 2016; Dogterom and Koenderink, 2019), which raises the possibility that actin-MT crosstalk supports nuclear shape and position. To assess whether perinuclear MTs were affected in Tmod-KD muscles, we quantified the number of intact radial MT arrays in control and Tmod-KD myofibers. We observed a significant decrease in intact perinuclear MT arrays in Tmod-KD muscles and noted a decrease in longitudinal MT-tracks (Fig. 5E; Fig. S6A). These data indicated that Tmod affects essential MT networks in myofibers, which we hypothesize occurs via its role in stabilizing F-actin, and thus mediating the interactions of MTs with actin filaments around the nuclei. A loss of perinuclear MTs could subsequently result in destabilization of nuclear shape and position.

To test whether nuclear activity was affected by nuclear morphology, we analyzed the transcriptional and translational potential of Tmod-KD myonuclei. To assess transcription, we quantified H3K9ac, a marker of open chromatin and transcriptionally active regions in the DNA (Kharchenko et al., 2011). Nuclei with the most irregular shapes (roundness<0.6) exhibited decreased H3K9ac levels, whereas wild-type (WT)-shaped nuclei (roundness>0.6) showed similar H3K9ac levels to control (Fig. 5F,H; Fig. S6B). To assess translation, we quantified nuclear intensity of Fibrillarin (Fib), a nucleolar protein that plays an essential role in ribosome biogenesis and is a commonly used readout for protein synthetic ability (Gillery et al., 1996; Lefèvre et al., 2001). Misshaped nuclei (roundness<0.6) exhibited similar Fib intensity levels to control, whereas WT-shaped nuclei (roundness>0.6) showed higher levels than control (Fig. 5G,I; Fig. S6C). These data suggested that loss of nuclear shape and position in Tmod-KD myofibers reduced transcriptional output but increased translational potential. While deformed nuclei are functionally compromised, we propose that WT-shaped nuclei in the Tmod-KD muscle compensate for this deficiency by increasing translation.

We analyzed the transcriptome of Tmod-KD larvae (Fig. 6, Fig. S7) to better understand the mechanisms behind the observed phenotypes and uncover additional consequences. Consistent with the morphological features found in Tmod-KD muscle, we detected misregulation of actin genes. Sarcomeric actin isoforms (Act57B, Act87E) (Röper et al., 2005; Fyrberg et al., 1983) were upregulated (Fig. 6D,E; Table S4), consistent with the observed ectopic F-actin bundles (Fig. 2F,G). In contrast, cytoplasmic actin isoforms (Act5C, Act42A) were downregulated in Tmod-KD myofibers (Fig. 6D,E; Table S4), consistent with the alterations in nuclear shape detected in Tmod-KD.

We also performed an over-representation analysis (ORA) to determine whether a set of shared genes was differentially expressed based on Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) gene sets. These data indicated that degradation-related genes (particularly proteasomal genes) and/or pathways were downregulated, whereas metabolism-linked genes (particularly oxidative metabolism genes linked to mitochondrial ATP production) and/or pathways were upregulated in Tmod-KD muscles (Fig. 6A,B; Fig. S7G,H). To test whether ATP production was indeed altered in Tmod-KD muscles, we performed a functional ATPase assay. Consistent with our RNA-seq data, ATP levels significantly increased in Tmod-KD myofibers (Fig. 6C). Together, these data suggested a role for Tmod, either directly (i.e. G-actin's role in transcription) or indirectly (i.e. due to nuclear deformities caused by myofibril disorganization) in gene expression. We speculate that these changes reflect a compensatory mechanism to promote growth in a degenerating muscle.

Tmod function has been studied using various cell types (Fowler, 1997). Here, we investigated the effects of Tmod-KD in Drosophila larval myofibers during development using a combination of detailed in vivo analyses, in vitro experiments and transcriptomic approaches. We propose that, through its capping function, Tmod affects actin dynamics in different locations within the myofiber and is ultimately essential for muscle function.

We suggest the following model for Tmod pleiotropic effects in muscle cells (Fig. 7). In the sarcomere, the barbed end of F-actin is capped by CapZ and anchored to the Z-disc, whereas the pointed-end is capped by Tmod. However, due to the low affinity of Tmod for F-actin (Weber et al., 1994), there are opportunities for monomer addition and therefore, the pointed-end remains suitable for sarcomere growth. When Tmod is knocked down, thin filaments grow in length. We suggest this additional polymerization causes sufficient imbalance to favor the production of muscle-specific larval G-actin isoforms (Act57B, Act87E); this increased G-actin availability and continuously growing sarcomeric F-actin would result in ectopic F-actin bundles and radial structures. In the cytoplasm, F-actin growth occurs through the canonical pathway, which is, generally, by the addition of G-actin monomers at the barbed end and depolymerization at the pointed-end. When Tmod is reduced, excessive depolymerization occurs, due to loss of Tmod capping activity. This excess of G-actin monomers decreases the production of cytoplasmic-specific actin isoforms (Act5C, Act42A). Loss of cytoplasmic perinuclear and nuclear F-actin would contribute to the loss of nuclear organization and directly or indirectly affect nuclear output. At the MTJ, myofibrils are anchored to the muscle ends via cytoplasmic F-actin. In the absence of Tmod, lack of stable attachments via F-actin impairs tension along the myofibrils, leading to myofibril disorganization and deterioration, and defects in myofibrillogenesis. Protein degradation and metabolism would be affected similarly by destabilization of required structures for myofiber organization or via transcriptional changes. Together, we suggest that loss of Tmod at these different sites contributes directly and/or indirectly to the observed phenotypes, ultimately leading to impaired muscle function.

Although every myofiber experiences the same Tmod-KD effect and follows the same pattern of deterioration, not every myofiber is affected equally. Along the larval AP axis, we see that the tmod phenotypes appear first and are most severe in anterior abdominal segments, which contain larger muscles than posterior segments, suggesting that growth and/or myofiber size may be a contributing factor. However, we observe that some muscles at the same axis position are preferentially affected, which suggests a more complex situation. Freely moving Drosophila larvae constantly pivot and sweep their heads, which are supported by muscles of segments A1-A3 (Lahiri et al., 2011); hence, muscle usage may affect muscle deterioration in our model. In parallel, the timing and distribution of muscle pathology in many human muscle diseases, as well as animal models, follow specific patterns (Barohn et al., 2014; Sewry et al., 2019). Studies in vertebrate models suggest that muscle fiber type might play a role (reviewed by Talbot and Maves, 2016). The factor(s) that make specific muscles more susceptible to deterioration remain unknown. The Drosophila larval musculature, consisting only of one fiber type, provides a useful system for further studies in this context.

Alternative splicing results in a total of 18 Tmod isoforms. Why the fly produces so many different isoforms is unclear; a crucial question remains as to whether a pointed-end nucleator exists in Drosophila. We show that isoforms Q and O/K display the Tmod characteristic F-actin capping function, despite having N-terminal extensions comparable with human LMOD. We did not find biochemical evidence of F-actin nucleation. It is possible that one of the remaining Tmod isoforms (most notably the isoforms with C-terminal proline-rich extensions) may possess nucleation activity like that of human LMODs. However, the extremely low levels of such isoforms in our RNA-seq data suggest they are not expressed in myofibers, or at the tested developmental stages. In muscle, a recent study identified Sals, an actin binding protein that promotes sarcomeric F-actin elongation from pointed-ends during muscle growth. Despite Sals being depicted as an elongator as opposed to a nucleator, it contains key regions (i.e. WH2 and PRD domains) (Tóth et al., 2016) that are found in Lmods and thought to confer its nucleating activity (Bai et al., 2007). It is unclear whether Sals also achieves a nucleator role in non-muscle tissues, as Sals appears to be expressed only in Drosophila muscle (Graveley et al., 2011; Robinson et al., 2013). Further research will be necessary to discover this.

Another important open question in the field is the mechanism by which thin filaments elongate. Both ‘competition’ (Tsukada et al., 2010) and ‘nucleation’ (Molnár et al., 2014) models have been proposed; nevertheless, it appears likely that there are elements of both in vivo. It would be interesting to test for genetic interactions of Tmod and Sals to address whether there is competition for thin filament extension regulation similar to the one hypothesized between Tmod and Lmod. As a test of Tmod and the nucleation model, one could test whether Tmod genetically interacts with barbed-end formins (such as DAAM and Fhos).

Tmod is suggested to define thin filament and sarcomere length in a variety of model systems (Szikora et al., 2022). In adult Drosophila, Tmod overexpression results in shorter sarcomeres and thin filaments (Molnár et al., 2014; Mardahl-Dumesnil and Fowler, 2001). In vertebrate systems, such as mouse Tmod-null mutants, thin filament length increases (Littlefield and Fowler, 2008). In vitro rat cardiomyocytes show increased or decreased Tmod levels resulting in shorter or longer sarcomeric actin filaments, respectively (Sussman et al., 1998). Our data demonstrate that Tmod-KD in Drosophila larval myofibers increases thin filament and sarcomere length. Thin filament to sarcomere length ratio was maintained in Tmod-KD myofibers compared with controls during larval development. This ratio was similar to measurements in frog sartorius (26%) and human pectorialis (30%) muscles (Gokhin and Fowler, 2013). This suggests that mechanisms that regulate sarcomere proportions are conserved and regulated independently of Tmod function. We further find sarcomere size increases throughout larval growth, indicating that the uncapped pointed-ends of sarcomeric actin filaments do not respect a specified size, exhibiting continuous polymerization in the absence of Tmod. In healthy muscle, sarcomere length is fixed to optimize muscle contraction, forcing growing cells to add sarcomeres rather than increasing their length (Balakrishnan et al., 2020; Williams and Goldspink, 1971; Dix and Eisenberg, 1990; Goldspink, 1983; Haas, 1950). We show an inverse relationship between sarcomere number and length in Tmod-KD myofibers, suggesting that myofibrils could grow by increasing their length via sarcomere growth. Consistent with recently published work, our data indicates that muscle length and sarcomere number are regulated independently of one another (Brooks et al., 2022). Loss of this adaptive behavior is associated with myopathy progression (Ottenheijm et al., 2009; Winter et al., 2016). Moreover, the decrease in myofiber size is suggestive of compromised myofibril addition to increase muscle mass, which further supports that mechanisms of muscle growth (addition of new sarcomeres and hypertrophy) are affected in Tmod-KD larvae. Our analyses show locomotive defects (first instar) before discernable changes in myofiber structure (second instar). Although technical limitations prevented the detailed analysis of first instar sarcomeres, we suggest that changes in thin filament length might be the first effects of Tmod-KD that directly affect muscle function. In parallel with disruption of sarcomeres and myofibrils, Tmod-KD myofibers display accumulation of ectopic and disorganized F-actin. The location of these ectopic bundled and radial F-actin accumulations near disorganized myofibrils indicates that continuously growing thin filaments could contribute to these ectopic structures. Whether the ectopic disorganized F-actin filaments were intended to serve as a myofibril template (Fenix et al., 2018; Dlugosz et al., 1984) or they result from deteriorating myofibrils remains unclear.

The currently favored hypothesis in the field describes sarcomere and myofibril formation as a tension-driven self-organizing process, whereby tension generated through muscle attachment at the MTJs is used to orient and longitudinally assemble sarcomeres and myofibrils (Lemke and Schnorrer, 2017). Here, we show that Tmod-KD results in mislocalization of two core mechanosensitive proteins and a reduction in tension at the muscle ends. According to the hypothesis, lack of tension at the muscle ends affects addition of new sarcomeres as well as homeostasis along the entire myofibril. This is consistent with our Tmod-KD data showing reduced sarcomere number and aberrant myofibril organization and stability. Previous studies using tmod mutant zebrafish also reported defects in myofibril organization and muscle weakness (Berger et al., 2022, 2014), but a role for Tmod in sarcomere addition had not been described. We also note that the myofibers in the Tmod-KD do not detach and round up, despite having reduced Integrin adhesion complex proteins at the MTJ (Brown et al., 2000).We believe that this is because of appropriate localization of the complex on the cuticular side of the myofiber (Fig. S5F) (Maartens and Brown, 2015). In addition, other adhesion proteins, known to be localized at the MTJ, including Dystroglycan and Kon-tiki (Valdivia et al., 2017), may contribute to the maintained tendon-myofiber adhesion observed in Tmod-KD larvae. Interestingly, we observe perpendicular myofibrils exclusively localized at the myofiber ends, which demonstrate sarcomeric organization albeit incorrect orientation. This raises the possibility that owing to the lack of longitudinal stability in this growth zone, the forming sarcomeres and growing myofibrils orient towards the next best structure that provides resistance, connecting to the MTJ, existing myofibrils or the visceral plasma membrane. A similar phenotype with misoriented myofibrils was observed in Drosophila larval muscle expressing the constitutively active form of C3G, a Rap1 activator (Shirinian et al., 2010). Importantly, Rap1 has been demonstrated to recruit Talin to the cell membrane (Shattil et al., 2010; Lee et al., 2009). We find ectopic localization of mechanosensitive proteins along the plasma membrane of Tmod-KD myofibers, which could lead to ectopic F-actin accumulations. This could result in local changes in tension throughout the myofiber and contribute to myofibril misalignment. Together, our study suggests that Tmod is crucial for maintaining alignment, stability and growth of myofibrils. Our data are consistent with the tension-driven self-organizing myofibrillogenesis hypothesis and adds a role for Tmod in tension mediation.

Our data implicates Tmod in nuclear positioning in the z-axis and maintaining nuclear morphology. Mispositioned and deformed nuclei are common in many myopathies and laminopathies (Romero, 2010; Steele-Stallard et al., 2018; Ross et al., 2019), highlighting the relevance of correct nuclear positioning for proper muscle function. Actin plays a structural role in and around the nucleus (Bettinger et al., 2004; Davidson and Cadot, 2021; Caridi et al., 2019). We hypothesized that Tmod-KD affects cytoplasmic F-actin to generate the nuclear phenotypes that we observed; we propose that Tmod, through its capping function, stabilizes nuclear and/or perinuclear actin to directly support nuclear integrity. However, we found defects in the MT networks surrounding the nuclei of Tmod-KD myofibers. MTs interact with actin filaments and are crucial in myonuclear positioning and integrity. Thus, we suggest that Tmod-KD has indirect effects on myonuclei by disrupting actin-MT interactions. In addition, defects in nuclear positioning and morphology could be indirectly affected by the loss myofibril structure and orientation. In Tmod-KD myofibers, nuclei are initially located at the periphery, but subsequently sink as myofibril structure is lost. Force exertion on the internalized nuclei could then lead to deformation of nuclear shape. These nuclei also show defects in transcriptional activity. Interestingly, WT-shaped nuclei in Tmod-KD myofibers display increased translation makers, consistent with a compensatory mechanism. In addition to the effects of nuclear morphology, it is possible that Tmod regulates transcription and/or translation via its effect on actin dynamics. G-actin has many important functions in the nucleus and nucleolus and directly regulates gene transcription (Visa and Percipalle, 2010; Bettinger et al., 2004). More research is needed to determine how deviation from normal positioning in three dimensions (x,y,z) results in impaired muscle function. To this end, future studies focused on the distribution of gene products throughout the myofiber when nuclear positioning is aberrant would be informative.

Our transcriptome analyses suggest that Tmod-KD affects two main processes, proteasomal degradation and mitochondrial metabolism, both essential for myofiber growth (Schiaffino et al., 2013). However, the decrease in degradation and increase in mitochondrial metabolism detected in Tmod-KD myofibers is inconsistent with the observed impaired muscle growth and muscle weakness. We suggest that this transcriptomic profile indicates a compensatory attempt of the myofibers to grow, counteracting the lack of tension along the myofiber and structural deterioration. It is possible that actin inside the nucleus causes these changes through structural and/or transcriptional changes (Chase et al., 2013; Visa and Percipalle, 2010). However, similar myofiber phenotypes and changes in metabolism have been observed in Drosophila TRIM32 (thin) mutant larval muscles (LaBeau-DiMenna et al., 2012; Bawa et al., 2020). Loss of Drosophila TRIM32, an E3-ubiquitin ligase, resulted in a decrease in glycolytic intermediates, indicating that TRIM32 switches metabolism from oxidative to glycolytic. The mechanisms by which TRIM32 and Tmod are connected remain to be elucidated, yet the similarities in phenotype and transcriptome suggest they might be interacting genetically or biochemically. This would indicate a role for Tmod in dampening oxidative metabolism and, therefore, ATP production. Strikingly, structural components involved in these pathways, such as mitochondria, are located in high densities at the visceral muscle surface and around myofibrils; hence, they could be affected indirectly via the destabilization of the underlying myofibrils.

Myopathies are often devastating disorders characterized by impaired muscle function that are frequently associated with sarcomeric mutations. The molecular details underpinning this relationship are poorly understood. Our study shows a series of phenotypes associated with KD of one essential thin filament component, Tmod, that reach far beyond the sarcomere. Our findings have important implications, as they suggest a key role for Tmod in several cellular processes, including the regulation of actin isoform expression, the stabilization of tension-mediating proteins, and nuclear integrity and positioning, therefore providing additional insight to the pathogenesis of myopathies. The mechanisms directly linking Tmod and crucial myofiber processes, including proteasomal degradation and oxidative metabolism, remain to be fleshed out. Furthermore, our findings support and strengthen the tension-driven self-organizing myofibrillogenesis model. Taken together, this study adds significantly to the understanding of Tmod in muscle development, maintenance and disease, making it a potential target for new therapeutics that address muscle weakness.

Drosophila stocks and experimental crosses were grown on standard cornmeal medium at 25°C in 12 h light/12 h dark conditions under humidity control. The Gal4-UAS system (Brand and Perrimon, 1993) was used for RNA interference studies. Details on the screen that lead to our analysis of tmod are found in Tables S1 and S2. For muscle-specific expression and RNAi potentiation we tested different driver lines (UAS-dicer2;;Mhc-Gal4,Zasp66::GFP, UAS-dicer2;;Dmef2-Gal4,Zasp66::GFP/Tm6,Hu,Tb and UAS-dicer2;;Dmef2-Gal4/Tm6,Hu,Tb) in combination with 3 UAS-tmod-RNAi lines [Bloomington Drosophila Stock Center (BDSC) 41718, BDSC 31534 and Vienna Drosophila Resource Center 108389] to validate on-target effects and explore KD levels. The driver lines were crossed with UAS-mCherryRNAi (BDSC 35785) as the control RNAi. In all examples, larvae displaying a Tb phenotype (indicatory of balancer chromosome) were excluded from the F₁ analyzed. The mutant stock tmod^MI02468 (BDSC 36446) was also used for phenotype validation with yw as control. tmod^MI02468 is a P-element insertion with a gene-trap cassette consisting of stop codons in all three reading frames in the first intronic region of 12 splice variants of tmod. It is homozygous viable, but not fertile. Except where specified, we used a Dmef2-Gal4 driver and one UAS-tmodRNAi line (BDSC 41718) for analyzing Tmod-KD phenotypes (the line is referred to as ‘tmod’). Table S5 contains details on fly stocks.

For all larval experiments, larvae were kept at 25°C and both male and female larvae were analyzed. Staging was performed on apple juice agar plates, either in the embryo through gut shape auto-fluorescence or by selecting larvae hatched within a 2 h period. First instars intended for experiments were maintained in an apple juice agar plate with yeast paste for ease of manipulation. Larvae for experiments at later stages (locomotion, muscle analysis) were transferred to standard food vials. Larvae were removed from food vials using 30% sucrose solution at second instar stage (∼24 h after transfer), early third instar (∼48 h after transfer), late third instar (∼96 h after transfer) and wandering (∼118 h after transfer) stage. Myofiber structure time-course analysis was performed on all four distinct larval stages, in the VI1 muscle or VL3/VL4 muscles of anterior segments (A1 and A2). For the RNA-seq experiment, late third instars (∼90 h after transfer) were processed for dissection. For all other experiments at late third instar stage (∼3.5 days), staging was confirmed by using developmental landmarks such as mouth hooks and spiracle morphologies (Bodenstein, 1950).

Ten third instar wandering larvae were dissected and rinsed in ice cold HL3.1 medium (Feng et al., 2004) as previously described (Brent et al., 2009). Total RNA was extracted from the muscle-enriched larval fillets using TRIzol reagent (Thermo Fisher Scientific, 15596026), followed by cleanup with TURBO DNA-free Kit (Ambion, AM1907). The SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen, 18080-051) was used to synthesize cDNA, and qRT-PCR reactions were performed on the BioRad CFX96 Real-Time PCR system using the SYBR Select Master Mix for CFX (Applied Biosystems, 4472937). Three independent mRNA preparations per genotype were collected and were run in triplicate. Fold changes were calculated based on values obtained using the delta-delta Ct method (Livak and Schmittgen, 2001; Schmittgen and Livak, 2008). Product size and uniformity was assessed via melt curve analysis. Rpl32 was used as a normalization control. Table S3 contains details on the oligonucleotides used.

Seven to ten late third instar wandering larvae were dissected (See Immunostaining and confocal imaging) and transferred into larval lysis buffer [50 mM HEPES (pH 7.5), 150 mM NaCl, 0.5% NP40, 0.1% SDS, 2 mM DTT] supplemented with Complete mini protease inhibitor cocktail (one tablet in 10 ml of lysis buffer; Roche, 11836153001). Lysates were homogenized and protein concentrations were determined using the Bradford assay. Equal amounts of protein (20 µg) were run on a 5% (Talin) or 10% (Tmod) polyacrylamide gel and then transferred onto a nitrocellulose membrane (Thermo Fisher Scientific, 88018). Membranes were blocked with either 5% milk or 5% bovine serum albumin (BSA) in TBST (Tris-buffered saline+0.1% Tween) for 1 h at room temperature. Table S5 contains details on primary and secondary antibodies. Immunoreactions were visualized in a KwikQuant Imager (Kindle Biosciences, D1001) using 1-Shot Digital-ECL (Kindle Biosciences, R1003) and intensities were quantified using Fiji (Abràmoff et al., 2004). Protein expression was normalized to GAPDH (loading control) within each sample. Figures and quantifications are representative of three experiments.

Larvae were staged and extracted from food vials (see Larval staging) at first instar, second instar, early third instar and late third instar. One larva at a time was placed at the center of an apple juice agar plate (8.5 cm diameter) and recorded for 45 s (Samsung Galaxy S8+). Movies were converted into image sequences (1 image per second) and each larva was tracked using the Manual Tracking plug-in on Fiji (Abràmoff et al., 2004). Average velocity was calculated as in Balakrishnan et al. (2021). For each genotype, we performed three replicates with ∼10 larvae each. Larval length was equivalent in both phenotypes for each developmental stage.

The cDNA for TmodO/K (NCBI Reference Sequences: NP_001247373.1/NP_001189319.1) was acquired from DNAsu clone DmCD00765633. The cDNA for TmodQ (NCBI Reference Sequence: NP_001263112.1) was synthesized by GeneWiz. Each cDNA was PCR-amplified and ligated between the SacI and KpnI sites of pRSF_Duet1 (Millipore Sigma) with an N-terminal 6× histidine tag and a C-terminal Strep-II tag.

Tmod isoforms in pRSF_Duet1 were expressed in ArcticExpress(DE3) RIL cells (Agilent Technologies), grown in Terrific Broth (TB) medium for 6 h at 37°C to an optical density of ∼1.5 at 600 nm (OD600), followed by 24 h at 10°C in 0.4 mM isopropyl-β-D-thiogalactoside (IPTG). Cells were harvested by centrifugation (4000 g), resuspended in nickel buffer [20 mM HEPES (pH 7.5), 300 mM NaCl, 10 mM imidazole, 1 mM PMSF] and lysed using a microfluidizer apparatus (Microfluidics). Proteins were first purified on nickel-NTA resin, washed with 20 column volumes of nickel buffer and eluted using nickel buffer supplemented with 300 mM imidazole. Elution fractions containing the Tmod proteins were then loaded onto Streptavidin XT resin (IBA Life Sciences) and washed with 20 column volumes of strep buffer [20 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM EDTA]. Proteins were eluted using strep buffer supplemented with 40 mM biotin, and further purified through a SD200HL 26/600 gel filtration column (GE Healthcare) in strep buffer supplemented with 1 mM DTT. Human CapZ (Rao et al., 2014), rabbit skeletal muscle actin (Pardee et al., 1982) and rabbit skeletal muscle tropomyosin (Smillie, 1982) were purified as described.

Sequences of Drosophila Tmod isoforms and human TMOD3 were aligned using the program Clustal Omega (Sievers et al., 2011). The alignment was visualized using the program Jalview (Waterhouse et al., 2009) and colored using the Clustal X color scheme.

Capping assays were carried out as previously described (Rao et al., 2014; Kumari et al., 2020). Briefly, 1.5 μM of pre-formed F-actin seeds, coated with 1 μM Tropomyosin and capped at the barbed end with saturating amounts of CapZ (25 nM), were incubated and assayed alone or with four different concentrations of Tmod (25, 100, 400, 800 nM). Upon addition of 1.5 μM G-actin (6% pyrene-labeled), reactions were monitored as the fluorescence increase resulting from the incorporation of pyrene-actin monomers at the pointed-end. Pyrene fluorescence (excitation 365 nm, emission 407 nm) was detected using a plate reader (BioTek Cytation 5), and for each Tmod concentration four replicates were recorded. Maximum polymerization rates were determined by taking the first derivative of each curve, identifying the time interval corresponding to the greatest slope and performing a linear regression on the original data within that time interval.

Larvae were dissected in ice-cold HL3.1 medium as described in Brent et al. (2009) and fixed with 10% formalin (Sigma-Aldrich, HT501128) for 20 min. Larval filets were blocked with PBT-BSA [1× PBS supplemented with 0.3% Triton X-100 (Sigma-Aldrich, X100) and 0.1% BSA (Sigma-Aldrich, A7906)] for 30 min. Larval filets were then incubated with primary antibody (Table S5) overnight at 4°C, followed by washes in PBT-BSA. Incubation with Alexa Fluor-conjugated secondary antibodies (Table S5), Phalloidin and/or Hoechst 3342 were performed at room temperature for 2 h, followed by washes in PBT. Subsequently, larval filets were mounted in ProLong Gold (Invitrogen, P36930) and slides were cured at room temperature for 24 h. Z-stacks were acquired using an SP5 laser-scanning microscope (Leica Microsystems) with either dry objectives (HC PL APO CS 10×/0.40 and HC PL APO CS2 20×/0.75) or oil-immersion objectives (HC PL APO CS 20×/0.70, HCX PL APO CS 40×/1.25-0.75, HCX PL APO CS 63×/1.40-0.60 and HCX PL APO CS 100×/1.46). All samples intended for direct comparison were imaged using the same confocal settings. Maximum intensity projections of confocal z-stacks were rendered using Fiji (Abràmoff et al., 2004). Approximately seven larvae (range: 5-10) of each genotype were dissected and experiments were repeated at least twice.

Muscle defects in second instar, early third instar and late third instar larvae were scored based on ectopic and abnormal actin structures using a confocal microscope. A score was given for each VL3/4 muscle pair: 3 – WT-like; 2 – mild structural defects (diagonal myofibrils); 1 – severe defects in muscle structure. We evaluated muscles in both hemisegments of the larva in abdominal segments 1-6. This analysis was performed in triplicate with 3-4 larvae per replicate.

Larvae expressing a Zasp66::GFP protein trap were fixed by brief (∼5 s) submersion in 65°C water and mounted on a slide with halocarbon oil. Z-stacks of the cuticular side of the VL3 or VL4 muscles from abdominal segment 2 were acquired using a SP5 laser-scanning microscope (Leica Microsystems) with either a HC PL APO CS2 20×/0.75 (late third instars) or HCX PL APO CS 40×/1.25-0.75 (second instars) oil immersion objective.

Muscle length, sarcomere size and number were quantified as described in step 32 of Balakrishnan et al. (2021). For accuracy and to assess potential regional differences, sarcomere size was also quantified by an alternative method. Z-stacks were opened in Fiji and a line that spanned six Z-discs (five sarcomeres) was made using the line tool. Subsequently, the line intensity plot was obtained, and the x-coordinates (microns) of the peaks were acquired with the multi-point tool. The distance in microns between Z-disks was obtained by calculating the difference between the x-coordinates of the six peaks. This analysis was carried out in triplicate. In each replicate, six muscles were analyzed, each belonging to an individual larva. Three lines were traced per muscle: in the anterior fraction of the muscle, in the medial region and in the posterior end, with a total of ∼90 sarcomere size measurements per replicate (Fig. S4C).

Sarcomere length was obtained from Zasp66::GFP heat-fixed larvae, which was performed by a direct method from Z-disc to Z-disc distance. Due to technical challenges in some areas of the muscles, two confirmatory methods were also employed. First, measurements were confirmed by comparing the heat-fixed samples with a formalin fixed sample. Second, we performed a sarcomere length average by taking the muscle length divided by the total number of Z-discs.

Thin filament length was assessed in formalin fixed samples that displayed a clear striation pattern and a clear M-line (therefore, we conclude the muscles were in a stretched configuration). Zasp66-GFP was used as a marker for Z-discs; we traced a line spanning three contiguous Z-discs. The line intensity plot of Phalloidin was obtained and was normalized by calculating deviation from the mean to account for background noise; those with values above zero were used to calculate thin filament length.

Maximum intensity projections of confocal z-stacks of β_PS-integrin, Talin and pTyr immunostainings were generated in Fiji (Abràmoff et al., 2004). A line spanning a set of three contiguous sarcomeres from the posterior end of the muscle, the MTJ and another set of three contiguous sarcomeres from the anterior end of the contiguous muscle was drawn with the line tool. Sarcomeres were used as a landmark/reference. MTJ was centered in the middle of the line. Line intensity plots were obtained with the intensity value for each pixel in the line. These values were split into five bins (each bin encompassing 20% of the intensity values) along the AP axis and an average was calculated for each bin. Normalization was accomplished by calculating the deviation from the mean value. We traced two lines per MTJ per larva (16-20 measurements per genotype from 4-5 different larvae) and per genotype. This analysis was carried out in duplicate.

All images were processed and analyzed with Fiji (Abràmoff et al., 2004). 2D quantifications of VL3 and VL4 muscles were performed with maximum intensity projections (as previously described in Windner et al., 2019). Muscle areas were traced by hand with the polygon tool. Automated thresholding of fluorescence intensities of anti-Lamin and/or Hoechst was used to generate binary images of VL nuclei. The number, size (area), position (x- and y-coordinates) and shape descriptors (roundness and circularity) of all nuclei within each cell was obtained and the binary masks were used to quantify Hoechst and H3K9ac pixel intensity (mean gray value). Automated thresholding of anti-Fibrillarin labeling was used to generate binary images of nucleoli and measure nucleolar sizes (areas) and to quantify Fibrillarin pixel intensity (mean gray value). The 3D rendering of nuclei was processed with Imaris. Mean gray value is defined as average gray value within the selection; this is the sum of the gray values of all the pixels in the selection divided by the number of pixels (Abràmoff et al., 2004; Schneider et al., 2012).

Seven wandering stage larvae for each genotype and per experiment were dissected as described above. Staining with α-tubulin was performed and myofibers of segments A2-6 were assessed based on the presence or absence of an array of perinuclear-MTs for each individual myonucleus within a myofiber. In control and mildly affected Tmod-KD myofibers, intact perinuclear-MTs were counted in absence of nuclear marker Hoechst to avoid bias. Quantification became more challenging due to the cell surface irregularities observed in severely affected Tmod-KD myofibers; hence, a conservative approach was used to count ‘intact’ perinuclear MT arrays: clearly circular and radial structures colocalizing with the nucleus were defined as ‘intact’. The total number of intact perinuclear-MTs was then divided by the total number of nuclei per cell, which was determined by Hoechst staining. These results were expressed as the average percentage of intact perinuclear-MTs.

Eight to ten late third instar larvae were dissected as described above per genotype in each replicate. The experiment was carried out in triplicate. The muscle-enriched larval filets were transferred to TRIzol and homogenized. The lysates were submitted to the Memorial Sloan Kettering Cancer Center (MSKCC) Integrated Genomics Operations (IGO) core for further total RNA extraction, quality assessment and Illumina next-generation sequencing. The sequencing libraries were constructed using SMARTerAmpSeq after the mRNAs were enriched by oligo-dT-mediated purification. The libraries were then sequenced on the Illumina HiSeq 4000 in a PE50 run. Over 40 million reads were generated per library. FASTQ data was read into R with the ShortRead package (Morgan et al., 2009) and quality control was performed with the Rfastp package (Wang and Carroll, 2022). Reads were mapped and aligned to the Drosophila genome using Rsubread (Liao et al., 2019). Next, the reads were counted with the GenomicAlignments package (Lawrence et al., 2013). Gene differential expression tests were performed using DESeq2 (Love et al., 2014). Finally, enrichment analysis was performed through goseq (Young et al., 2010) using both GO and KEGG databases. Plots were drawn using R with the ggplot2 package (Wickham, 2016). Lastly, transcript isoform analysis was performed using RSEM-1.3.3. (Li and Dewey, 2011), a commonly used tool that quantified isoforms from short-reads.

An ATP calibration curve was performed using the stabilized ATP standard stocks provided in the kit ATP Bioluminescence Assay Kit HS II (Millipore Sigma, 11699709001) and following the supplied protocol. Five to ten late third instar wandering larvae from each genotype were dissected (See Immunostaining and confocal imaging) and transferred into the cell lysis reagent provided with the kit. Lysates were homogenized and protein concentrations were determined using the Bradford assay. Equal amounts of protein (10 µg) were used for bioluminescence measurements. Blank measurements were subtracted from the raw data and ATP concentrations were calculated from the standard curve data. The experiment was carried out in triplicate.

Statistical significance was determined by either two-tailed paired or unpaired Student's t-test (comparisons between two groups), one-way ANOVA or two-way ANOVA (comparisons among multiple groups) in the GraphPad Prism software. For each statistical analysis, the specific test used is indicated in the relevant figure legend. Data are presented as mean±s.d., asterisks are used to denote significance. Graphs were made using either GraphPad Prism, Microsoft Excel or R.

We thank the Baylies lab members, F. Schnorrer, G. Tanentzapf, A. Beggs and V. Gupta for helpful discussions and H. Bellen and F. Schöck for providing antibodies. We thank the members of the IGO and the Bioinformatics Cores at MSKCC.